Handbook of Zoology: Volume 3 Gastrotricha and Gnathifera 9783110274271, 9783110273816

Table of contents :
List of contributing authors
1 Gastrotricha
1.1 Introduction
1.2 Morphology
1.2.1 General and external morphology
1.2.2 Integument
1.2.3 Musculature
1.2.4 Nervous system
1.2.5 Sensory structures
1.2.6 Intestinal system
1.2.7 Body cavities and connective tissue
1.2.8 Excretory system
1.2.9 Reproductive organs
1.2.9.1 Female gonad
1.2.9.2 Male gonad
1.2.9.3 Frontal and caudal organ
1.2.9.4 Reproductive system of the Paucitubulatina
1.2.9.5 Additional accessory structures
1.2.10 Gametes
1.2.10.1 Spermatozoa
1.2.10.2 Spermatogenesis and spermiogenesis
1.2.10.3 Eggs
1.2.10.4 Oogenesis
1.3 Reproduction and development
1.3.1 Reproductive biology
1.3.2 Cleavage and development
1.4 Physiology
1.5 Phylogeny
1.6 Systematics
1.6.1 Order Macrodasyida Brunson, 1950
1.6.1.1 Family Cephalodasyidae Hummon & Todaro, 2010
1.6.1.2 Family Dactylopodolidae Strand, 1929
1.6.1.3 [Family Lepidodasyidae Remane, 1927]
1.6.1.4 Family Macrodasyidae Remane, 1926
1.6.1.5 Family Planodasyidae Chandrasekhara Rao & Clausen, 1970
1.6.1.6 Family Thaumastodermatidae Remane, 1927
1.6.1.7 Family Turbanellidae Remane, 1927
1.6.1.8 Family Xenodasyidae Todaro, Guidi, Leasi & Tongiorgi, 2006
1.6.1.9 Family Redudasyidae Todaro, Dal Zotto, Jondelius, Hochberg, Hummon, Kånneby & Rocha, 2012
1.6.2 Order Chaetonotida Brunson, 1950
1.6.2.1 [Suborder Multitubulatina d’Hondt, 1971]
1.6.2.1.1 [Family Neodasyidae Remane, 1929]
1.6.2.2 Suborder Paucitubulatina d’Hondt, 1971
1.6.2.2.1 Family Muselliferidae Leasi & Todaro, 2008
1.6.2.2.2 Family Xenotrichulidae Remane, 1936
1.6.2.2.3 Family Chaetonotidae Gosse, 1864
1.6.2.2.4 Family Dasydytidae Daday, 1905
1.6.2.2.5 Family Neogosseidae Remane 1927
1.6.2.2.6 [Family Dichaeturidae Remane, 1927]
1.6.2.2.7 Family Proichthydidae Remane 1927
1.6.2.3 Gastrotricha incertae sedis
1.7 Biogeography
1.8 Ecology
Acknowledgments
Literature
2 Phylum Gnathostomulida
2.1 Introduction and history of research (Fig. 2.1)
2.2 Morphology
2.2.1 General and external morphology (Figs. 2.2–2.6)
2.2.2 Integument (Figs. 2.7 and 2.8)
2.2.3 Musculature (Figs. 2.9 and 2.10)
2.2.4 Nervous system (Fig. 2.11)
2.2.5 Sensory structures (Fig. 2.12)
2.2.6 Intestinal system (Figs. 2.13–2.36)
2.2.6.1 Pharynx
2.2.6.2 Jaws
2.2.6.3 Basal plate and jugum
2.2.7 Body cavities and connective tissue
2.2.8 Excretory system (Fig. 2.37)
2.2.9 Reproductive organs (Figs. 2.38–2.41)
2.2.9.1 Female organs
2.2.9.2 Male organs
2.2.10 Gametes
2.2.10.1 Eggs and oogenesis
2.2.10.2 Sperm and spermiogenesis (Figs. 2.39 E and 2.42–2.46)
2.2.10.2.1 Spermiogenesis
2.3 Reproduction and development
2.3.1 Reproductive biology
2.3.2 Oviposition and cleavage
2.3.3 Development (Fig. 2.47)
2.4 Biology and physiology (Figs. 2.48 and 2.49
2.5 Ecology
2.6 Phylogeny
2.7 Systematics
2.7.1 Phylum Gnathostomulida (Ax, 1956) Riedl, 1969
2.7.1.1 Order Filospermoidea Sterrer, 1972
2.7.1.1.1 Family Haplognathiidae, Sterrer 1972
2.7.1.1.2 Family Pterognathiidae Sterrer, 1972
2.7.1.2 Suborder Conophoralia Sterrer, 1972
2.7.1.2.1 Family Austrognathiidae Sterrer, 1971
2.7.1.3 Order Bursovaginoidea Sterrer, 1972
2.7.1.4 Suborder Scleroperalia Sterrer, 1972
2.7.1.4.1 Family Gnathostomariidae Sterrer, 1972
2.7.1.4.2 Family Rastrognathiidae Kristensen & Nørrevang, 1977
2.7.1.4.3 Family Agnathiellidae Sterrer, 1972
2.7.1.4.4 Family Mesognathariidae Sterrer, 1972
2.7.1.4.5 Family Paucidentulidae terrer, 1998
2.7.1.4.6 Family Clausognathiidae Sterrer, 1992
2.7.1.4.7 Family Problognathiidae Sterrer & Farris, 1975
2.7.1.4.8 Family Onychognathiidae Sterrer, 1972
2.7.1.4.9 Family Gnathostomulidae Sterrer, 1972
2.8 Biogeography
2.9 Paleontology
Acknowledgments
Literature
3 Micrognathozoa
3.1 Introduction
3.2 Morphology
3.2.1 General and external morphology
3.2.2 Integument
3.2.3 Musculature
3.2.4 Nervous system
3.2.5 Sensory structures
3.2.6 Intestinal system
3.2.7 Body cavities and connective tissue
3.2.8 Excretory system
3.2.9 Reproductive organs
3.2.10 Gametes
3.3 Reproduction and development
3.4 Physiology
3.5 Phylogeny
3.6 Systematics
3.7 Biogeography
3.8 Paleontology
3.9 Ecology
Literature
4 Rotifera
4.1 Introduction
4.2 Morphology
4.2.1 General and external morphology
4.2.1.1 Morphology of the female
4.2.1.2 Coloniality
4.2.1.3 Corona
4.2.1.4 Morphology of the male
4.2.2 Integument
4.2.2.1 Pedal glands
4.2.2.2 Retrocerebral organ
4.2.2.3 Sheats and tubes
4.2.3 Musculature
4.2.3.1 Musculature of the female
4.2.3.2 Musculature of the male
4.2.4 Nervous system
4.2.5 Sensory structures
4.2.6 Intestinal system
4.2.6.1 Intestinal system of the female
4.2.6.2 Trophi
4.2.6.3 Intestinal system of the male
4.2.7 Body cavity
4.2.8 Excretory system
4.2.8.1 Excretory system of the female
4.2.8.2 Excretory system of the male
4.2.9 Reproductive organs
4.2.9.1 Reproductive organs of the female
4.2.9.2 Reproductive organs of the male
4.2.10 Gametes
4.3 Reproduction and development
4.3.1 Reproductive biology
4.3.2 Cleavage
4.3.3 Development
4.4 Physiology
4.5 Phylogeny
4.6 Systematics
4.6.1 Classification
4.6.2 Keys
4.6.2.1 Key to higher taxa
4.6.2.2 Key to families of Bdelloidea
4.6.2.3 Key to families of Ploima
4.6.2.4 Key to families of Flosculariacea
4.6.2.5 Key to families of Collothecacea
4.6.3 Characterization of families
4.7 Biogeography
4.8 Paleontology
4.9 Ecology
4.9.1 Feeding ecology
4.9.2 Habitat
4.9.2.1 Freshwater and limnoterrestrial habitats
4.9.2.2 Saline environments
4.9.2.3 Symbiotic associations
Literature
5 Acanthocephala: functional morphology
5.1 Introduction
5.2 General aspects
5.3 Body wall: host-parasite interface
5.4 Acanthocephalan muscles
5.5 Reproductive system and reproduction
5.6 Development: host-parasite interface in the intermediate host
5.7 Excretion and osmoregulation
5.8 Nervous and sensory receptor system
5.9 Phylogenetic considerations
Literature
6 Acanthocephala
6.1 Introduction
6.2 Phylogeny
6.3 Systematics
6.3.1 Background to classification
6.3.2 Selection of diagnostic characters
6.3.3 Characterization of the classes of the Acanthocephala
6.3.4 Characterization of orders and families of the Acanthocephala
6.3.4.1 Archiacanthocephala
6.3.4.1.1 Apororhynchida: Apororhynchidae Shipley, 1899 (Fig.6.2)
6.3.4.1.2 Gigantorhynchida: Gigantorhynchidae Hamann, 1892 (Fig.6.3)
6.3.4.1.3 Moniliformida: Moniliformidae Van Cleave, 1924 (Figs.6.1 and 6.3)
6.3.4.1.4 Oligacanthorhynchida: Oligacanthorhynchidae Southwell & MacFie, 1925 (Fig.6.3)
6.3.4.2 Eoacanthocephala
6.3.4.2.1 Gyracanthocephalida: Quadrigyridae Van Cleave, 1920 (Fig.6.4)
6.3.4.2.2 Neoechinorhynchida: Dendronucleatidae Sokolovskaia, 1962
6.3.4.2.3 Neoechinorhynchida: Neoechinorhynchidae Ward, 1917 (Fig.6.4)
6.3.4.2.4 Neoechinorhynchida: Tenuisentidae Van Cleave, 1936 (Fig.6.5)
6.3.4.3 Palaeacanthocepha
6.3.4.3.1 Echinorhynchida
6.3.4.3.2 Echinorhynchida: Arhythmacanthidae Yamaguti, 1935 (Fig. 6.5)
6.3.4.3.3 Echinorhynchida: Cavisomidae Meyer,1932 (Fig.6.5)
6.3.4.3.4 Echinorhynchida: Diplosentidae Tubangi & Masilungan, 1937 (Fig. 6.6)
6.3.4.3.5 Echinorhynchida: Echinorhynchidae Cobbold, 1876 (Fig.6.5)
6.3.4.3.6 Echinorhynchida: Fessisentidae Van Cleave, 1931 (Fig.6.5)
6.3.4.3.7 Echinorhynchida: Heteracanthocephalidae Petrochenko, 1956 (Fig. 6.6)
6.3.4.3.8 Echinorhynchida: Illiosentidae Golvan, 1960 (Fig.6.6)
6.3.4.3.9 Echinorhynchida: Isthmosacanthidae Smales, 2012 (Fig.6.6)
6.3.4.3.10 Echinorhynchida: Pomphorhynchidae Yamaguti, 1939 (Fig.6.7)
6.3.4.3.11 Echinorhynchida: Rhadinorhynchidae Travassos, 1923 (Fig.6.7)
6.3.4.3.12 Echinorhynchida: Sauracanthorhynchidae Bursey, Goldberg & Kraus,2007 (Fig. 6.9)
6.3.4.3.13 Echinorhynchida: Transvenidae Pichelin & Cribb, 2001 (Fig.6.7)
6.3.4.3.14 Heteramorphida: Pyrirhynchidae Amin & Van Ha, 2008 (Fig.6.8)
6.3.4.3.15 Polymorphida: Centrorhynchidae Van Cleave, 1916 (Fig.6.8)
6.3.4.3.16 Polymorphida: Plagiorhynchidae Golvan, 1960 (Fig.6.8)
6.3.4.3.17 Polymorphida: Polymorphidae Meyer,1931 (Fig.6.8)
6.3.4.4 Polyacanthocephala
6.3.4.5 Uncertain assignment
6.5 Taxonomic keys
6.5.1 Key to the classes of the Acanthocephala
6.5.2 Key to the orders and families of the Archiacanthocephala
6.5.3 Key to the orders and families of the Eoacanthocephala
6.5.4 Key to the orders and families of the Palaeacanthocephala
Literature
7 Ecology of the Acanthocephala
7.1 Introduction
7.2 Life cycle
7.3 Behavior
7.4 Population ecology
7.5 Community ecology
7.6 Environmental parasitology
Literature
Index

Citation preview

Handbook of Zoology Gastrotricha, Cycloneuralia and Gnathifera Volume 3: Gastrotricha and Gnathifera

Handbook of Zoology Founded by Willy Kükenthal Editor-in-chief Andreas Schmidt-Rhaesa

Gastrotricha, Cycloneuralia and Gnathifera Edited by Andreas Schmidt-Rhaesa

DE GRUYTER

Gastrotricha, Cycloneuralia and Gnathifera Volume 3: Gastrotricha and Gnathifera Edited by Andreas Schmidt-Rhaesa

DE GRUYTER

Scientific Editor Andreas Schmidt-Rhaesa Centrum für Naturkunde (CeNak), Universität Hamburg, Zoologisches Museum Martin-Luther-King-Platz 3 20146 Hamburg, Germany

ISBN 978-3-11-027381-6 e-ISBN (EPUB) 978-3-11-027427-1 e-ISBN (EPUB) 978-3-11-038556-4 ISSN 2193-4231 Library of Congress Cataloging-in-Publication Data A CIP catalogue record for this book is available from the Library of Congress. 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.d-nb.de.

© 2015 Walter de Gruyter GmbH, Berlin/Munich/Boston Printing and Binding: Hubert & Co. GmbH & Co. KG, Göttingen Printed in Germany www.degruyter.com

Introduction This third subvolume of the volume “Gastrotricha, Cycloneuralia and Gnathifera” deals with the taxa Gastrotricha and Gnathifera. Gastrotrichs have become a phylogenetically mysterious group because despite some corresponding characters between gastrotrichs and cycloneuralian taxa, their DNA tells a different story. According to DNA comparisons so far, gastrotrichs cluster within Spiralia, often close to platyzoan groups (see Schmidt-Rhaesa 2013). Therefore, gastrotrichs are presented here in a more or less isolated way. Gnathiferans include animals that are morphologically and ecologically very disparate at first view. Specimens are small or large, free-living or parasitic, and display a number of morphological differences. This has made it quite difficult to recognize relationships among these taxa, until the right characters were put into focus. It is especially the jaw apparatus and the structure of the epidermis that provide the key to the understanding of gnathiferan relationships. Rotifers and acanthocephalans, so different they might appear, share a corresponding structure of the (syncytial) epidermis, which has led to the common name Syndermata. Gnathostomulida are related to Syndermata based on the fine structure of the jaw apparatus, and the discovery of Micrognathozoa supported these hypotheses (see Schmidt-Rhaesa 2012 for details and literature).

Most open questions concern the relationship between rotifers and acanthocephalans. Rotifers include three subtaxa (Bdelloidea, Monogononta, and Seisonacea), and it is still under debate whether acanthocephalans are related to Rotifera as a whole, to a part of them, or to one of its subgroups. We decided here to present rotifers (Bdelloidea and Monogononta) and Seisonacea in two different chapters. Unfortunately, the Seisonacea chapter could not be completed in time and will occur in a later volume of the Handbook of Zoology series. Acanthocephala are split into three single chapters on morphology, systematic, and ecology. With this third subvolume, we finish the volume “Gastrotricha, Cycloneuralia and Gnathifera” and hope that we could present an up-to-date review and overview on these often neglected but nevertheless important and fascinating animal groups. I thank all the authors for their contribution and De Gruyter for the enthusiasm to realize these volumes. Andreas Schmidt-Rhaesa

Literature Schmidt-Rhaesa, A. (2013): Gastrotricha, Cycloneuralia and Gnathifera: general history and phylogeny. In: SchmidtRhaesa, A. (ed.) Handbook of zoology, volume Nematomorpha, Priapulida, Kinorhyncha, Loricifera, pp. 1–10. De Gruyter, Berlin.

Contents List of contributing authors  

  XI

Alexander Kieneke and Andreas Schmidt-Rhaesa 1 Gastrotricha   1 1.1 Introduction 1 1.2 Morphology 1 1.2.1 General and external morphology 1 1.2.2 Integument 4 1.2.3 Musculature 8 1.2.4 Nervous system 24 1.2.5 Sensory structures 26 1.2.6 Intestinal system 34 1.2.7 Body cavities and connective tissue 38 1.2.8 Excretory system 39 1.2.9 Reproductive organs 45 1.2.9.1 Female gonad 49 1.2.9.2 Male gonad 50 1.2.9.3 Frontal and caudal organ 51 1.2.9.4 Reproductive system of the Paucitubulatina 56 1.2.9.5 Additional accessory structures 56 1.2.10 Gametes 57 1.2.10.1 Spermatozoa 57 1.2.10.2 Spermatogenesis and spermiogenesis 70 1.2.10.3 Eggs 74 1.2.10.4 Oogenesis 75 78 1.3 Reproduction and development 1.3.1 Reproductive biology 78 1.3.2 Cleavage and development 84 1.4 Physiology 87 1.5 Phylogeny 87 1.6 Systematics 92 1.6.1 Order Macrodasyida Brunson, 1950 93 1.6.1.1 Family Cephalodasyidae Hummon & Todaro, 2010 93 1.6.1.2 Family Dactylopodolidae Strand, 1929 95 1.6.1.3 [Family Lepidodasyidae Remane, 1927] 96 1.6.1.4 Family Macrodasyidae Remane, 1926 97 1.6.1.5 Family Planodasyidae Chandrasekhara Rao & Clausen, 1970 99 1.6.1.6 Family Thaumastodermatidae Remane, 1927 100 1.6.1.7 Family Turbanellidae Remane, 1927 105

Family Xenodasyidae Todaro, Guidi, Leasi & Tongiorgi, 2006 108 1.6.1.9 Family Redudasyidae Todaro, Dal Zotto, Jondelius, Hochberg, Hummon, Kånneby & Rocha, 2012 109 1.6.2 Order Chaetonotida Brunson, 1950 110 1.6.2.1 [Suborder Multitubulatina d’Hondt, 1971] 110 1.6.2.1.1 [Family Neodasyidae Remane, 1929] 110 1.6.2.2 Suborder Paucitubulatina d’Hondt, 1971 111 1.6.2.2.1 Family Muselliferidae Leasi & Todaro, 2008 111 1.6.2.2.2 Family Xenotrichulidae Remane, 1936 112 1.6.2.2.3 Family Chaetonotidae Gosse, 1864 114 1.6.2.2.4 Family Dasydytidae Daday, 1905 121 1.6.2.2.5 Family Neogosseidae Remane 1927 123 1.6.2.2.6 [Family Dichaeturidae Remane, 1927] 124 1.6.2.2.7 Family Proichthydidae Remane 1927 124 1.6.2.3 Gastrotricha incertae sedis 124 1.7 Biogeography 124 1.8 Ecology 125 Acknowledgments 126 Literature 126 1.6.1.8

Wolfgang Sterrer and Martin V. Sørensen  135 Phylum Gnathostomulida Introduction and history of research (Fig. 2.1) 135 2.2 Morphology 143 2.2.1 General and external morphology (Figs. 2.2–2.6) 143 2.2.2 Integument (Figs. 2.7 and 2.8) 145 2.2.3 Musculature (Figs. 2.9 and 2.10) 149 2.2.4 Nervous system (Fig. 2.11) 151 2.2.5 Sensory structures (Fig. 2.12) 152 2.2.6 Intestinal system (Figs. 2.13–2.36) 154 2.2.6.1 Pharynx 154 2.2.6.2 Jaws 158 2.2.6.3 Basal plate and jugum 169 2.2.7 Body cavities and connective tissue 171 2.2.8 Excretory system (Fig. 2.37) 172 2.2.9 Reproductive organs (Figs. 2.38–2.41) 172 2.2.9.1 Female organs 172 2.2.9.2 Male organs 174 2 2.1

viii 

 Contents

2.2.10 Gametes 176 2.2.10.1 Eggs and oogenesis 176 2.2.10.2 Sperm and spermiogenesis (Figs. 2.39 E and 2.42–2.46) 176 2.2.10.2.1 Spermiogenesis 177 2.3 Reproduction and development 180 2.3.1 Reproductive biology 180 2.3.2 Oviposition and cleavage 181 2.3.3 Development (Fig. 2.47) 182 2.4 Biology and physiology (Figs. 2.48 and 2.49 182 2.5 Ecology 183 2.6 Phylogeny 184 2.7 Systematics 187 2.7.1 Phylum Gnathostomulida (Ax, 1956) Riedl, 1969 187 2.7.1.1 Order Filospermoidea Sterrer, 1972 187 Family Haplognathiidae, 2.7.1.1.1 Sterrer 1972 187 2.7.1.1.2 Family Pterognathiidae Sterrer, 1972 188 2.7.1.2 Suborder Conophoralia Sterrer, 1972 188 2.7.1.2.1 Family Austrognathiidae Sterrer, 1971 188 2.7.1.3 Order Bursovaginoidea Sterrer, 1972 189 2.7.1.4 Suborder Scleroperalia Sterrer, 1972 189 2.7.1.4.1 Family Gnathostomariidae Sterrer, 1972 189 2.7.1.4.2 Family Rastrognathiidae Kristensen & Nørrevang, 1977 189 2.7.1.4.3 Family Agnathiellidae Sterrer, 1972 189 2.7.1.4.4 Family Mesognathariidae Sterrer, 1972 190 2.7.1.4.5 Family Paucidentulidae Sterrer, 1998 190 2.7.1.4.6 Family Clausognathiidae Sterrer, 1992 190 2.7.1.4.7 Family Problognathiidae Sterrer & Farris, 1975 190 2.7.1.4.8 Family Onychognathiidae Sterrer, 1972 190 2.7.1.4.9 Family Gnathostomulidae Sterrer, 1972 191 2.8 Biogeography 192 2.9 Paleontology 192 192 Acknowledgments Literature 192 Martin V. Sørensen and Reinhardt M. Kristensen 3 Micrognathozoa  197 3.1 Introduction 197

197 3.2 Morphology 3.2.1 General and external morphology 197 3.2.2 Integument 200 3.2.3 Musculature 204 3.2.4 Nervous system 205 3.2.5 Sensory structures 206 3.2.6 Intestinal system 207 3.2.7 Body cavities and connective tissue 211 3.2.8 Excretory system 211 3.2.9 Reproductive organs 212 3.2.10 Gametes 212 3.3 Reproduction and development 213 3.4 Physiology 213 3.5 Phylogeny 213 3.6 Systematics 215 3.7 Biogeography 215 3.8 Paleontology 216 3.9 Ecology 216 Literature 216 Diego Fontaneto and Willem H. De Smet 4 Rotifera  217 4.1 Introduction 217 4.2 Morphology 217 4.2.1 General and external morphology 218 4.2.1.1 Morphology of the female 218 4.2.1.2 Coloniality 221 4.2.1.3 Corona 221 4.2.1.4 Morphology of the male 225 4.2.2 Integument 227 4.2.2.1 Pedal glands 228 4.2.2.2 Retrocerebral organ 228 4.2.2.3 Sheats and tubes 229 4.2.3 Musculature 231 4.2.3.1 Musculature of the female 231 4.2.3.2 Musculature of the male 233 4.2.4 Nervous system 233 4.2.5 Sensory structures 234 4.2.6 Intestinal system 238 4.2.6.1 Intestinal system of the female 238 4.2.6.2 Trophi 242 4.2.6.3 Intestinal system of the male 245 4.2.7 Body cavity 246 4.2.8 Excretory system 246 4.2.8.1 Excretory system of the female 246 4.2.8.2 Excretory system of the male 248 4.2.9 Reproductive organs 248 4.2.9.1 Reproductive organs of the female 248 4.2.9.2 Reproductive organs of the male 249 4.2.10 Gametes 249 4.3 Reproduction and development 251

Contents     ix

4.3.1 Reproductive biology 251 4.3.2 Cleavage 252 4.3.3 Development 253 4.4 Physiology 253 4.5 Phylogeny 254 4.6 Systematics 255 4.6.1 Classification 255 4.6.2 Keys 259 4.6.2.1 Key to higher taxa 259 4.6.2.2 Key to families of Bdelloidea 259 4.6.2.3 Key to families of Ploima 259 4.6.2.4 Key to families of Flosculariacea 262 4.6.2.5 Key to families of Collothecacea 262 4.6.3 Characterization of families 262 4.7 Biogeography 273 4.8 Paleontology 274 4.9 Ecology 275 Feeding ecology 4.9.1 275 4.9.2 Habitat 276 4.9.2.1 Freshwater and limnoterrestrial habitats 276 4.9.2.2 Saline environments 282 4.9.2.3 Symbiotic associations 284 Literature 286 Horst Taraschewski  301 5 Acanthocephala: functional morphology 5.1 Introduction 301 5.2 General aspects 301 5.3 Body wall: host-parasite interface 305 5.4 Acanthocephalan muscles 308 5.5 Reproductive system and reproduction 309 5.6 Development: host-parasite interface in the intermediate host 310 5.7 Excretion and osmoregulation 314 5.8 Nervous and sensory receptor system 314 5.9 Phylogenetic considerations 315 Literature 315 Lesley Smales  317 6 Acanthocephala 6.1 Introduction 317 6.2 Phylogeny 317 6.3 Systematics 318 6.3.1 Background to classification 318 6.3.2 Selection of diagnostic characters 319 6.3.3 Characterization of the classes of the Acanthocephala 321

6.3.4

Characterization of orders and families of the Acanthocephala 321 6.3.4.1 Archiacanthocephala 321 6.3.4.1.1 Apororhynchida: Apororhynchidae Shipley, 1899 (Fig. 6.2) 322 6.3.4.1.2 Gigantorhynchida: Gigantorhynchidae Hamann, 1892 (Fig. 6.3) 322 6.3.4.1.3 Moniliformida: Moniliformidae Van Cleave, 1924 (Figs. 6.1 and 6.3) 322 6.3.4.1.4 Oligacanthorhynchida: Oligacanthorhynchidae Southwell & MacFie, 1925 (Fig. 6.3) 322 6.3.4.2 Eoacanthocephala 323 6.3.4.2.1 Gyracanthocephalida: Quadrigyridae Van Cleave, 1920 (Fig. 6.4) 323 6.3.4.2.2 Neoechinorhynchida: Dendronucleatidae Sokolovskaia, 1962 323 6.3.4.2.3 Neoechinorhynchida: Neoechinorhynchidae Ward, 1917 (Fig. 6.4) 323 6.3.4.2.4 Neoechinorhynchida: Tenuisentidae Van Cleave, 1936 (Fig. 6.5) 323 6.3.4.3 Palaeacanthocepha 324 6.3.4.3.1 Echinorhynchida 324 6.3.4.3.2 Echinorhynchida: Arhythmacanthidae Yamaguti, 1935 (Fig. 6.5) 325 6.3.4.3.3 Echinorhynchida: Cavisomidae Meyer, 1932 (Fig. 6.5) 325 6.3.4.3.4 Echinorhynchida: Diplosentidae Tubangi & Masilungan, 1937 (Fig. 6.6) 326 6.3.4.3.5 Echinorhynchida: Echinorhynchidae Cobbold, 1876 (Fig. 6.5) 327 6.3.4.3.6 Echinorhynchida: Fessisentidae Van Cleave, 1931 (Fig. 6.5) 327 6.3.4.3.7 Echinorhynchida: Heteracanthocephalidae Petrochenko, 1956 (Fig. 6.6) 327 6.3.4.3.8 Echinorhynchida: Illiosentidae Golvan, 1960 (Fig. 6.6) 327 6.3.4.3.9 Echinorhynchida: Isthmosacanthidae Smales, 2012 (Fig. 6.6) 327 6.3.4.3.10 Echinorhynchida: Pomphorhynchidae Yamaguti, 1939 (Fig. 6.7) 328 6.3.4.3.11 Echinorhynchida: Rhadinorhynchidae Travassos, 1923 (Fig. 6.7) 328 6.3.4.3.12 Echinorhynchida: Sauracanthorhynchidae Bursey, Goldberg & Kraus, 2007 (Fig. 6.9) 329 6.3.4.3.13 Echinorhynchida: Transvenidae Pichelin & Cribb, 2001 (Fig. 6.7) 329 6.3.4.3.14 Heteramorphida: Pyrirhynchidae Amin & Van Ha, 2008 (Fig. 6.8) 329

x 

 Contents

6.3.4.3.15 Polymorphida: Centrorhynchidae Van Cleave, 1916 (Fig. 6.8) 329 6.3.4.3.16 Polymorphida: Plagiorhynchidae Golvan, 1960 (Fig. 6.8) 329 6.3.4.3.17 Polymorphida: Polymorphidae Meyer, 1931 (Fig. 6.8) 329 6.3.4.4 Polyacanthocephala 330 6.3.4.5 Uncertain assignment 330 6.5 Taxonomic keys 331 6.5.1 Key to the classes of the Acanthocephala 331 6.5.2 Key to the orders and families of the Archiacanthocephala 332 6.5.3 Key to the orders and families of the Eoacanthocephala 332

Key to the orders and families of the Palaeacanthocephala 332 Literature 334 6.5.4

Bernd Sures 7 Ecology of the Acanthocephala 7.1 Introduction 337 7.2 Life cycle 337 7.3 Behavior 339 7.4 Population ecology 341 7.5 Community ecology 341 7.6 Environmental parasitology Literature 342 Index 

 345

 337

341

List of contributing authors Willem H. De Smet University of Antwerp Department of Biology Campus Drie Eiken Universiteitsplein 1 2610 Wilrijk Belgium Dr. Diego Fontaneto National Research Council Institute of Ecosystem Study Largo Tonolli 50 28922 Verbania Pallanza Italy Dr. Alexander Kieneke Forschungsinstitut Senckenberg DZMB Südstrand 44 26382 Wilhelmshaven Germany Prof. Dr. Reinhardt M. Kristensen University of Copenhagen Natural History Museum of Denmark Universitetsparken 15 2100 Copenhagen Denmark Prof. Dr. Andreas Schmidt-Rhaesa University Hamburg Centrum für Naturkunde Martin-Luther-King-Platz 3 20146 Hamburg Germany

Prof. Dr. Lesley Smales South Australia Museum Parasitology Section North Terrace Adelaide, SA 5000 Australia Prof. Dr. Martin V. Sørensen Natural History Museum of Denmark Øster Voldgade 5-7 1350 Copenhagen K Denmark Dr. Wolfgang Sterrer Bermuda National History Museum PO Box FL145 Flatts FLBX Bermuda Prof. Dr. Bernd Sures Universität Duisburg-Essen Aquatische Ökologie 45117-Essen Germany Prof. Dr. Horst Taraschewski Karlsruher Institut für Technologie (KIT) Zoologisches Institut Kaiserstr. 12 76131 Karlsruhe Germany

Alexander Kieneke and Andreas Schmidt-Rhaesa

1 Gastrotricha 1.1 Introduction

1.2 Morphology

The microscopically small gastrotrichs are abundant in diverse aquatic habitats. Gastrotrichs must have been among the first small animals studied with early microscopes. Remane (1936) lists some reports from the 18th century, which likely represent gastrotrich specimens. At first, gastrotrichs were treated, together with protozoans, rotifers, and other tiny animals, as “infusorians”, until Ehrenberg (1830) separated gastrotrichs and rotifers from protozoans. Ehrenberg treated gastrotrichs as part of rotifers and only subsequently researchers recognized the differences between these two groups. Mečnikow (1865) introduced the name Gastrotricha. Originally, gastrotrichs were only found in freshwater, until Schultze (1853) found the first marine species, Turbanella hyalina, in sandy samples from the island Neuwerk (North Sea). Soon after, Claparéde (1867) described Hemidasys agaso from the harbour in Naples (Mediterranean Sea). The main era of the discovery of marine species started with Remane’s intensive investigations of marine sediments in the Kiel Bight (Baltic Sea) (Remane 1924, 1925a, 1926a, b, 1927a, b, see also 1927c, 1929, 1936). Today, we know about 780 species, a number that is constantly growing. Still, the gastrotrich fauna of many places is unknown or has been sampled only superficially. Gastrotrichs occur in a variety of freshwater habitats and in the sea, from the littoral region to the deep sea. In the marine environment, gastrotrichs are part of the mesopsammon, the community of animals living in the pore system between sand grains. In freshwater, gastrotrichs are either benthic (mesopsammic or epipelic, respectively) or live among vegetation, some species are swimming in the free water. The first broad account to gastrotrich morphology was made by Zelinka (1889). Because of the minute size of gastrotrichs, transmission electron microscopy (TEM) played an important role to understand their internal anatomy (see, e.g., Ruppert 1991). The position of gastrotrichs in the phylogenetic system has changed several times and still is not solved convincingly.

1.2.1 General and external morphology Most gastrotrich species are microscopically small animals in the size range of a few hundred micrometers. Especially among Paucitubulatina, there are very small species with about 70 µm length. The longest gastrotrichs belong to the macrodasyid genus Megadasys and reach up to 3.5 mm in length (Schmidt 1974). The body form varies quite a bit. Almost all species in the taxon Paucitubulatina are more or less tenpin-shaped with a clearly defined head region, a narrowing neck, and a slightly bulbous trunk (Fig. 1.1 A). Species of Macrodasyida vary much more in shape. A head and neck region is present in several species, but often not very distinct. The trunk is usually of equal diameter throughout, giving the animals a shape that is often called “strap-shaped” (Fig. 1.1 B). Some species are short and broad and can best be termed “tongue-shaped”. The chaetonotid genus Neodasys resembles the strap-shaped macrodasyids in body outline. In the posterior end, many gastrotrichs have paired extensions often called feet or, in paucitubulatinans, furca (Fig. 1.1 A, B). The feet carry one or more pairs of adhesive tubes (see below). In Paucitubulatina, only one pair of adhesive tubes is present on the furca. The exceptions are the absence of feet in swimming chaetonotids in the taxa Neogosseidae and Dasydytidae as well as reported two pairs in the genera Dichaetura and Diuronotus (see, e.g., Schwank 1990, Todaro et al. 2005). A recent description of Dichaetura filispina (Suzuki et al. 2013) found one adhesive tube and a solid spine on each foot, making a reinvestigation of the other species of the genus desirable. In macrodasyidan species, there is usually more than one adhesive tube on each foot (Fig. 1.1 B). When feet are absent, the posterior end is rounded or an unpaired tail is present. This tail is most conspicuous and several times as long as the trunk in species of the genus Urodasys (e.g., Wilke 1954, Schoepfer-Sterrer 1974). Based on their parsimonious character optimization, Kieneke et al. (2008a) reconstructed the stem species of Gastrotricha as an elongate (wormshaped), dorsoventrally flattened, benthic-marine animal with a rounded frontal and a bilobed caudal trunk end.

2 

 1 Gastrotricha

Further appendage-like extensions of the body wall occur in some species, such extensions either act as a basal structure for adhesive tubes or as sensory structures in the head region (see chapter Sensory Structures and descriptions of genera). Adhesive tubes are slender extensions, often accompanied by a cilium. They have a secretory function (see below). In chaetonotids of the subtaxon Paucitubulatina, they are restricted to the feet, but in macrodasyids and in Neodasys, adhesive tubes occur on various body regions (Fig. 1.1 A, B). Usually, adhesive tubes are divided into three major groups (see Fig. 1.1 B). Anterior adhesive tubes (usually abbreviated TbA) are the tubes in the head region. Often, they are concentrated in paired clusters on the ventral side; in some cases, they originate from a common basis. This basis is sometimes called “hand” or “fleshy base”. The lateral adhesive tubes (TbL) are positioned along the body in a lateral position; they may be few (e.g., in Dactylopodola) or very many (e.g., in different species of Turbanella). Posterior adhesive tubes (TbP) are those tubes present on feet or, when feet are absent, on the posterior end of the animals. In addition to the TbL, there may also be dorsal (TbD), dorsolateral (TbDL), ventrolateral (TbVL),

A

or even ventral (TbV) adhesive tubes arranged along the entire body or restricted to certain regions of the trunk. The stem species of Gastrotricha at least had adhesive tubes in the lateral (TbL) and posterior (TbP) arrangements (Kieneke et al. 2008a). The mouth opening is terminal or subterminal on the anterior tip of the animal. In some species, it leads to a funnel- or barrel-shaped buccal cavity. In some species (e.g., from the genera Diplodasys, Oregodasys, Ptychostomella, and Tetranchyroderma), the mouth opens very broad and occupies almost the entire anterior end, dorsally shielded by the so-called oral hood. Very characteristic and important for determination is the covering of the body by cilia and cuticular structures. The restriction of locomotory cilia to the ventral side of the animals is the name-giving feature of gastrotrichs (Fig. 1.2 D–H). Cilia occur as a broad field, as transverse rows, as isolated paired patches, or as paired longitudinal bands in the trunk region (Remane 1936). In the head region, they usually cover the entire ventral surface. An exception occurs in species of the Xenotrichulidae; here, cilia in the anterior region and in the midtrunk region are tightly packed together and form so-called cirri (Fig. 1.1 A). Further, isolated and often stiff

B TbA

cir

TbP

fu

TbL

TbP 50 µm

100 µm

Fig. 1.1: Gross body organization of Gastrotricha. (A) Xenotrichula velox (Paucitubulatina) from Tulip Beach, Lee Stocking Island (Bahamas), ventral view. (B) Turbanella hyalina (Macrodasyida) from the intertidal at Schillig, Northern Germany, ventral view. (A and B) differential interference contrast (DIC). Abbreviations: cir, ventral locomotorc cirri; fu, furca; TbA, anterior adhesive tubes; TbL, lateral adhesive tubes; TbP, posterior adhesive tubes.

1.2 Morphology 

A

B

 3

C

100 µm

D

lci 100 µm sci

100 µm

25 µm

E F

G

sci

H

lci

sci

lci

sp

lci

25 µm

25 µm

lci

50 µm

50 µm

Fig. 1.2: Epidermis and external cilia of Gastrotricha. (A) Neodasys uchidai (Multitubulatina). Dorsal view of anterior end. Light microscopic bright field (BF) image. (B) Macrodasys sp. Horizontal view of anterior end. (C) Dorsal view of a juvenile Macrodasys. Note the vacuolated epidermis cells in A–C (white triangles). (D) Anterior end of a marine Aspidiophorus sp., lateral view. Cilia are restricted to the ventral side. (B–D) DIC. (E–G) Maximum projections of confocal image stacks. In all 3 species, α-tubulin was stained with fluorescence-labeled antibodies thereby making visible locomotor and sensory cilia. (E) Pseudostomella roscovita. (F) Thaumastoderma ramuliferum. (G) Tetranchyroderma sp. (H) Scanning electron microscopic (SEM) image of the ventral surface of a Turbanella subterranea. Note the paired columns of locomotory cilia. Abbreviations: lci, locomotory cilia; sci, sensory cilia; sp, cilia of spermatozoa.

4 

 1 Gastrotricha

cilia may be seen in different locations, particularly in the anterior end. Such cilia are assumed to have sensory functions (see chapter Sensory Structures). The entire body is covered by a cuticle, and this cuticle may form various structures covering the body, especially on the dorsal side. Such coverings are spines, scales, or mixtures of these forms. In macrodasyids, scales occur, for example, in genera such as Lepidodasys and Diplodasys and spines occur in Acanthodasys. However, species of Diplodasys additionally possess spines at the lateral margins while species of Acanthodasys may display a spiny surface interspersed with tiny scales. Very conspicuous are cuticular structures in which three to five curved and pointed branches originate from a common base. Such structures are called triancres, tetrancres, and pentancres, and occur in the genera Pseudostomella, Thaumastoderma, and Tetranchyroderma. In chaetonotids, the variability of spines and scales is more diverse and characteristic for the genera. Spines can originate from scales (spined scales) or scales can be stalked, i.e., they rest on a cuticular rod that is basally connected to the cuticle that directly lines the epidermis. There are, however, also many species with a rather thin cuticle without any of the above-mentioned differentiations. This condition was probably also present in the last common ancestor of Gastrotricha (Kieneke et al. 2008a).

1.2.2 Integument The integument of gastrotrichs is composed of a layer of epidermal cells, the cuticle, and the basal extracellular matrix (ECM). Also described here are glandular structures associated with the epidermis, in particular the adhesive tubes and epidermal glands. The epidermis is cellular, and the cells differ in their structure between the dorsal and the ventral sides of the animal. The ventral cells are usually cuboidal ciliated cells; the dorsal cells are flatter, contain less cytoplasm, and lack cilia (Ruppert 1991). Epidermal cells are connected to each other by adhaerens junctions and septate junctions (Ruppert 1991). Adhaerens junctions are mechanical cell-cell connections, and they occur in various types, of which desmosomes and hemidesmosomes are the most well-known ones (see Schmidt-Rhaesa 2007). At least in species of Macrodasys, the dorsal epidermal cells are large and contain a vacuole (Teuchert 1978, Ruppert 1991; Fig. 1.2 A–C), this condition suggests either a skeletal function during

locomotion (Teuchert 1978) or possibly an adaptation to the interstitial habitat (Ax 1966). To our knowledge, gap junctions have not been shown in gastrotrichs but should be present because they are broadly distributed among eumetazoans (see, e.g., Schmidt-Rhaesa 2007). The basal lamina (ECM) is very thin or may even be absent in gastrotrichs (Ruppert 1991). The name-giving feature of gastrotrichs is the presence of locomotory cilia on the ventral side of the animals (Fig. 1.2 D). Their action allows ciliary gliding, which is the most important form of locomotion (see chapter Musculature for muscle-aided locomotion). The ventral cells have either one cilium per cell (monociliated; Fig. 1.3 A) or more cilia per cell (few up to about 40; multiciliated; Fig. 1.3 B). Monociliated cells occur in several macrodasyids and in Neodasys, multiciliated cells in all Paucitubulatina and several macrodasyids (Rieger 1976, Boaden 1985, Ruppert 1991). Cilia have the usual axonemal pattern of internal microtubules (nine peripheral duplets and two single central ones) and the usual basal structure of nine peripheral triplets of microtubuli (Fig. 1.3 E). An accessory centriole is present and a pair of ciliary rootlets anchors the cilia in the epidermal cells (Rieger 1976, Ruppert 1991; Fig. 1.3 A, B). In most species, a rostral and a caudal rootlet are present (see Rieger 1976 for length measurements), only in the investigated species of Lepidodasys and Xenotrichula is the rostral rootlet absent (Rieger 1976). Species of Xenotrichulidae are peculiar in possessing cirri, which are bundles of cilia that act as a functional unit (Ruppert 1979; Fig. 1.3 B). In Xenotrichula, each cirrus as a whole is surrounded by epicuticle (Rieger 1976, Ruppert 1991; Fig. 1.3 B), wheraes this is not the case in the xenotrichulid genus Draculiciteria (Ruppert 1991). The rootlets of all cilia from one cirrus form a bundle that is anchored in the epidermal cell (Rieger 1976, Ruppert 1991; Fig. 1.3 B). Cirri also occur in species of the genus Oregodasys, but these have not been investigated ultrastructurally to date. The epidermis is covered by a cuticle, and the pharynx also has a cuticular lining. Further cuticular structures are rare; in some species of Urodasys, a cuticular stylet is present in the reproductive system (Schoepfer-Sterrer 1974). A further presumably cuticular hard part of the reproductive system may be the recently discovered sclerotic canal inside the caudal organ of Tetranchyroderma bronchostylus (Atherton & Hochberg 2012). The body cuticle is composed of two layers, the endocuticle and the epicuticle. Please note that the outer layer is often called “exocuticle”, but for comparative reasons explained below, we prefer the

1.2 Morphology 

A

 5

B nu

* * enc

*

epc

*

lci cir 1 µm

0.5 µm

C

D

epc

epc

enc

ve

0.5 µm

E

mi

epc ve 2 µm

term “epicuticle”. The epicuticle covers the entire body, including the sensory and locomotory cilia (Fig. 1.3 D, E), which is a very peculiar feature among animals and much likely an autapomorphy of the Gastrotricha (Kieneke et  al. 2008a). Sensory cilia in Tetranchyroderma adelae are more complex; in these case, both cuticular layers, endocuticle and epicuticle, surround each cilium (Hochberg 2008). Within the endocuticular layer, 10 microvilli are embedded (Hochberg 2008). Among gastrotrich species, the cuticle varies in thickness and may be smooth or sculptured. The fine structure of the cuticle was extensively investigated by Rieger & Rieger (1977) and Ruppert (1991); if not otherwise indicated, the following data refer to these sources. The thickness of the cuticle ranges from 100  nm up to 4 µm. The two layers, epicuticle and endocuticle, can always be distinguished. The epicuticle is composed of a varying number of layers. Each such layer is usually trilaminate, which means that it is composed of an electron-dark outer and inner sublayer and an electron-lucent middle sublayer. In some species, this trilaminate substructure of the individual layers

0.25 µm

Fig. 1.3: Ultrastructure of epidermis, external cilia and cuticle of Gastrotricha. (A) Neodasys chaetonotoideus (Multitubulatina), ventral epidermis with monociliated epithelial cells. Note the cellular junctions/belt desmosomes (asterisks) and the basal bodies of 2 cilia (white triangles). (B) Xenotrichula carolinensis (Paucitubulatina), multiciliary ventral epidermis cell forming a locomotory cirrus. Note the bundle of ciliary rootlets (white triangle). (C) Dorsal epidermis of Tetranchyroderma sp. (Macrodasyida) with a pentancre formed by endocuticle and epicuticle. (D and E) Cross sections of locomotory cilia covered by epicuticle: (D) Chaetonotus maximus and (E) Neodasys chaetonotoideus. (A–E) TEM images of cross-sectioned specimens. Abbreviations: cir, locomotory cirrus (compound cilium); enc, endocuticle; epc, epicuticle; lci, locomotory cilium; mi, mitochondrium; nu, nucleus of epidermis cell; ve, electron-dark vesicles.

has not been recognized and appears to be a thin monolayer (observed in representatives of Crasiella, Dactylopodola, and Urodasys) or a thicker monolayer (observed in Neodasys, Fig. 1.3 A, E). The number of layers ranges from 1 to 25, species with an unsculptured (smooth) cuticle have 2 to 25 layers, species with a sculptured cuticle have 1–18 layers (see, e.g., Balsamo et  al. 2010a for one layer of epicuticle in Diuronotus aspetos and Musellifer delamarei). The endocuticle is granular or fibrous in fine structure, and sometimes, a subdivision is observed. In this case, there is an outer striated, a middle finely fibrous, and an inner loosely fibrous substructure. Especially in species of Paucitubulatina, there are local thickenings of the cuticle in the head region that have the appearance of cuticular plates (cephalion, pleura, hypostomium) (Fig. 1.4 J, K). The presence and shape of these plates is of taxonomic importance. Ultrastructural sections through these regions are not available, but there is a peculiar surface structure of curved ridges on the cuticular plates on the head of Lepidodermella squamata observable with the scanning electron microscope (SEM) (Hochberg 2001).

6 

 1 Gastrotricha

When cuticular structures such as spines or scales are present (Fig. 1.4 A–I), these are formations of the endocuticle. Rieger & Rieger (1977) and Ruppert (1991) recognized three different types of such structures. In Xenodasys, the cuticular structures are hollow and include processes of the epidermis. In all other cases, structures are only made of cuticular material, and no epidermal components extend into them. In macrodasyids (Lepidodasys and Thaumastodermatidae), cuticular structures are solid local thickenings in the endocuticle (see Hochberg 2008 as one example; Fig. 1.3 C). Sometimes a substructure such as a fine striation or a honeycomb pattern can be observed. In representatives of Paucitubulatina, cuticular structures are derivates of the outer sublayer of the endocuticle; they have a homogeneous or finely striated substructure. In Diuronotus aspetos and Musellifer delamarei, the solid scales are made up of two electron-dense, homogeneous layers (Balsamo et al. 2010a). Sometimes scales and spines of  Paucitubulatina are hollow structures, but in contrast to Xenodasys, they never include epidermal extensions. The gastrotrich cuticle does not contain chitin (Neuhaus et  al. 1996), and is not molted during deve-

A

B

10 µm

E

C

4 µm

F

lopment (Ruppert 1991). Despite the fact that molecular analyses do not favor a close relationship between gastrotrichs and cycloneuralians (nematodes and related groups, see Schmidt-Rhaesa 2013), the structure of the cuticle appears comparable to some extent. Cycloneuralians probably have an ancestral cuticular structure composed of three layers, a proteinaceous endocuticle, a chitinous exocuticle, and a trilaminate epicuticle (see Schmidt-Rhaesa et  al. 1998). The trilaminate epicuticle and the proteinaceous endocuticle appear comparable and could argue for a common ancestor of Cycloneuralia and Gastrotricha. During gastrotrich evolution, the epicuticle becomes multiplied; during cycloneuralian evolution, an additional layer, the chitinous exocuticle, occurs (Schmidt-Rhaesa 2002). In macrodasyids, the epidermis often contains glandular cells (glandulocytes), the so-called epidermal glands (Fig. 1.5 A–D). Epidermal glands may be arranged in paired longitudinal rows along the dorsal side of the animals. Each epidermal gland is composed of a single, flask-shaped glandulocyte and acts as an individual unit. Beside the sparsely arranged organelles and a rather big,

D

2 µm

G

5 µm

J

ce

10 µm

K 50 µm

25 µm

H

50 µm

I 20 µm

4 µm

ce ep

hp 8 µm

Fig. 1.4: Cuticular differentiations of Gastrotricha. (A) Dorsal tile-like scales of Diplodasys rothei (Macrodasyida). (B) Spined scales of Chaetonotus schulzei (Paucitubulatina). Note the pair of denticles slightly proximal to the tip of each spine. (C) Dorsal keeled scales of Aspidiophorus sp. (Paucitubulatina) (D) Pentancres of Tetranchyroderma sp. (Macrodasyida). (E) Another species of Tetranchyroderma with tetrancres. (F) Keeled scales of Lepidodasys sp. (Macrodasyida). (G) Slightly polygonal scales of Draculiciteria tesselata (Paucitubulatina). (H) Heterolepidoderma sp. (Paucilubulatina) with keeled scales, lateral view. (I) Lateral thorn of Diplodasys rothei. (J) Head region of Chaetonotus maximus (Paucitubulatina), dorsal view. (K) Head region of another species of Chaetonotus, lateral view. Note the “mouth basket” around the mouth opening. (A–D and H–K) SEM images. (E–G) DIC images. Abbreviations: ce, cephalion; ep, epipleurion; hp, hypopleurion.

1.2 Morphology 

and on the posterior end in paucitubulatinan chaetonotids. These structures are tube-like extensions of the body cuticle containing two types of glandulocytes (all information in this section from Tyler & Rieger 1980 and Ruppert 1991. These glandulocytes are basal to the tubes and extend through the tube to open at its apical end where the cuticle is broken by one or more pores. Two different types of glandulocytes are distinguished (Fig. 1.8 A–B). One produces a secretion made up of larger, electron-dense vesicles, and this is called “viscid gland cell” and is assumed to have an adhesive function. The other glandulocyte produces smaller vesicles, this is called “releasing gland cell” and is assumed to dissolve the adhesive secretion and release the attachment. In the investigated cases, one releasing gland cell and one to few viscid gland cells have been observed. They usually open through one apical pore, but in Tetranchyroderma sp., each of the two viscid cells has its own pore. The structure of the adhesive tubes corresponds to the definition of the “duo-gland adhesive system” (see, e.g., Tyler 1988). Further, glandulocytes (epidermal glands) or

basally positioned nucleus, there are few big secretion granules with irregularly staining content (Fig. 1.6). Secretion products are released through an apical pore within the cuticle (Teuchert 1977a, Ruppert 1991). In Turbanella cornuta, the apical neck of the glandulocyte is formed by a cellular protrusion that is surrounded by up to 50 microvilli, which do not penetrate the cuticle. Further proximal, 10 rings of short microvilli surround the neck of the epidermal gland (Fig. 1.6). It is hypothesized that these microvilli serve as a mechanic protection against pressure from the surrounding epidermis cells (Teuchert 1977a). In Oregodasys katharinae, the glandular system is more complex, it comprises at least three types of papillae beneath the cuticle, blunt, triangle-shaped and sensory ones (Hochberg 2010a). Furthermore, insunk glandulocytes are present. Until now, there is no convincing hypothesis for the functional role of the epidermal gland system in macrodasyidan gastrotrichs (Ruppert 1991, but see Hochberg 2010b). Conspicuous glandular structures of the Gastrotricha are the adhesive tubes (Fig. 1.7), which are present in different body regions in macrodasyids

A

B

 7

C mg

eg eg eg

eg eg

50 µm

100 µm

at

20 µm

D

4 µm

Fig. 1.5: Epidermal glands of Macrodasyida. (A) Diplodasys sp., habitus. Note the slightly colored epidermal glands. (B) Different types of epidermal glands of Diplodasys cf. meloriae. (C) Epidermal glands and adhesive tubes within the vacuolated epidermis of Turbanella bocqueti. (D) Elevated pore of an epidermal gland of Tetranchyroderma sp. (A) BF-image (B and C) DIC Images. (D) SEM image. Abbreviations: at, adhesive tube; eg, epidermal gland; mg, midgut.

8 

 1 Gastrotricha

po

sg

mv cut

cj gn mv

ed

ed sg

gc nu

er di mi

2 µm

Fig. 1.6: Epidermal gland of Turbanella cornuta, schematic. Note the circularly arranged microvilli on the surface of the neck of the gland cell. The proximal extension of the gland cell leads to the basal matrix. Abbreviations: cj, cellular junction (adhaerens junction and septate junction); cut, cuticle; di, dictyosome; ed, epidermis; er, endoplasmic reticulum; gc, gland cell; gn, neck of the gland cell; mi, mitochondria; mv, microvilli; nu, nucleus; po, pore of the epidermal gland; sg, secretory granule. (According to figure 5 of Teuchert 1977).

monociliated sensory cells can be associated with the adhesive tubes (see chapter Sensory Structures). A peculiar exception to the structure described above occurs in the genus Neodasys (Tyler et al. 1980, see also Ruppert 1991; Fig. 1.7 E). Neodasys has, from a general appearance, a distribution of adhesive tubes comparable to macrodasyidan species (however, anterior adhesive tubes are absent in Neodasys), but the fine structure of these tubes is not comparable. The papilliform lateral tubes in Neodasys contain only one glandulocyte per tube, this cell has a rudimentary cilium. A ciliated sensory cell is closely associated with this glandulocyte (Fig. 1.8 C). Tyler et al. (1980) interpret this structure as a kind of forerunner of the duo-gland adhesive tubes of other gastrotrichs. According to this model, the viscid gland cell(s) of the adhesive tubes are derived from ciliated epidermal

cells, and the releasing gland cell is derived from the associated sensory cell. Such a scenario also supports the hypothesis of Neodasys as the earliest branch within the phylogenetic system of Gastrotricha as proposed by Kieneke et  al. (2008a). The posterior adhesive organs of Neodasys are also of peculiar structure. The paired caudal feet include adhesive tubes similar in ultrastructure to the lateral ones but additionally possess a distal pore lined by a microvillar border of a specialized myoepithelial cell (Tyler et al. 1980, Ruppert 1991).

1.2.3 Musculature Solely based on light microscopic investigations, Remane (1936) already provided a quite detailed understanding of the muscular system in different species of the Gastrotricha, for instance, in Macrodasys sp., Turbanella cornuta, Dactylopodola baltica, Chaetonotus sp., Chaetonotus simrothi, Aspidiophorus paradoxus, and Dasydytes ornatus. Although he was not able to present any data on circular muscles in Gastrotricha in his slightly earlier monograph (Remane 1929), the occurrence of this muscle component could be demonstrated a few years later in taxa such as Polymerurus, Dactylopodola, Pleurodasys, and Oregodasys (Remane 1936). He concluded that Gastrotricha are characterized by a musculature consisting of a system of separate strands of longitudinal and circular muscles in contrast to the nearly closed muscular sheath below the epidermis composed of outer circular muscle layer and inner longitudinal muscle layer in many other vermiform taxa of the Bilateria, a consideration that is currently up-to-date. However, some of Remane’s (1936) findings had to be revised or complemented as well, as new techniques and methods provided a much more detailed view on muscle arrangement of microscopic animals. The resolving power of electron microscopy, primarily TEM, gave a first insight to the ultrastructure and cytomorphology of single muscle cells of Gastrotricha. Muscle ultrastructure was intensely studied in Turbanella cornuta by Teuchert (1974) and will be reviewed below. Then, the combination of specific fluorescence staining methods (e.g., staining of f-actin with fluorochrome-labeled phalloidin) with three dimensionally resolving confocal laser scanning microscopy (or initially conventional epifluorescence microscopy) provided a holistic look on the myoanatomy of many microscopic invertebrate taxa including the Gastrotricha. Richard Hochberg and Marian K. Litvaitis were the first to demonstrate the diversity of muscle arrangement among several gastrotrich species

1.2 Morphology 

A

B

C

TbVL

mo

D

eg

*

TbA

50 µm

E

F

TbA

G H

50 µm

50 µm

50 µm

TbP

100 µm

*

TbP

100 µm

TbVL

I

50 µm

50 µm

J

30 µm

4 µm

K

4 µm

Fig. 1.7: Different arrangements of adhesive tubes in Gastrotricha. (A) Tetranchyroderma sp., ventral view of the head. (B) Posterior trunk end of Macrodasys sp., ventral view. (C) Ventral view of Xenodasys riedli showing the peculiar adhesive organs or pedicles (asterisks). (D) Rear trunk end of Macrodasys caudatus that is tightly glued to the microscopic slide (white triangle). (E) Neodasys chaetonotoideus that vigorously adheres to the glass slide and to sediment particles (triangles). (F) Anterior and posterior end of Megadasys sp., ventral view. (G) Caudal furca of Draculiciteria tesselata (Paucitubulatina), ventral view. (H) Underside of Diplodasys sp. with ventrolateral adhesive tubes. (I) Ventral side of the head of Cephalodasys maximus with hand-like arranged anterior adhesive tubes on a “fleshy base”. (J) Posterior lateral adhesive tubes of Dactylopodola baltica. (K) Posterior adhesive tubes of Tetranchyroderma sp. on a pedicle. (A–C and F–H) DIC images. (D and E) BF images. (I–K) SEM images. Abbreviations: mo, mouth opening; TbA, anterior adhesive tubes; TbP, posterior adhesive tubes; TbVL, ventrolateral adhesive tubes.

 9

10 

 1 Gastrotricha

A B

cut

cut

vg rg

ed ed nu 1 µm

vg nb

C

cil

mv

bb cr

cut

nu gc

mi di

nb

sc

er

3 µm

Fig. 1.8: Adhesive organs (adhesive tubes) of Gastrotricha. (A) Longitudinal section (schematic) of a dorsolateral adhesive tube of Tetranchyroderma sp. consisting of 2 viscid glands and 1 releasing gland. (B) Cross section of that tube (level indicated by bold line in A). Note the larger and electron-dark vesicles of the viscid glands and the smaller and lighter vesicles of the releasing gland. In Tetranchyroderma sp., each gland cell has its own cuticular pore. (C) Longitudinal section (schematic) of a lateral adhesive tube of Neodasys sp. A sensory cell, probably mechanoreceptive in function, is closely associated with the single gland cell. A releasing gland is missing in Neodasys. Abbreviations: bb, basal body; cil, cilium of the sensory cell; cr, ciliary rootlet; cut, cuticle; di, dictyosome; ed, epidermis; er, endoplasmic reticulum; gc, gland cell; mi, mitochondria; mv, microvilli; nb, neurite bundle; nu, nucleus; rg, releasing gland; sc, sensory cell; vg, viscid gland. (A and B, According to a TEM micrograph of Tyler & Rieger 1980; C, modified from Tyler et al. 1980.)

representing various taxa (Hochberg & Litvaitis 2001a–d, 2003a,Hochberg 2005). Among other findings, these studies comprise the discovery of helicoidally arranged muscles that partially enwrap the gut tube (pharynx plus midgut) and represent a unique character (autapomorphy) of the Gastrotricha (Hochberg & Litvaitis 2001a). Furthermore, quite a high abundance of circular muscles (in visceral as well as somatic positions) was observed in macrodasyidan species and in Neodasys (e.g., Hochberg & Litvaitis 2001b, Hochberg 2005), unlike what was anticipated before by Remane (1936). Hochberg & Litvaitis (2001b) also tested and demonstrated the phylogenetic value of muscle characters of Gastrotricha and established a speciescharacter-matrix. This matrix, which has been expanded by current data since, provides a thorough survey of general muscle patterns of the Macrodasyida (Tab. 1.1). Taking the muscular characters of several species of Macrodasyida and Paucitubulatina and those of two investigated species of the phylogenetically important taxon Neodasys (see chapter Phylogeny) into account, Hochberg (2005) suggests a “primitive organization” of the gastrotrich musculature, i.e., the character pattern that was probably present in the last common ancestor (stem species) of Gastrotricha. According to this scenario, the stem species was provided with muscle strands in three different orientations, longitudinal, circular, and helicoidal (Figs. 1.9 and 1.10). The myoepithelial sucking pharynx (for details of the contractile elements of the pharynx see below) is surrounded by numerous consecutive visceral muscle rings followed by visceral longitudinal muscle fibers that accompany the whole gut tube from the terminal mouth opening to the ventral anus. These visceral longitudinal muscles are located dorsal, lateral, and ventral to the gut tube. In the intestinal region posterior to the pharynx, the visceral longitudinal muscle fibers are surrounded by aligned visceral muscle rings. Hence, the spatial arrangement of visceral longitudinal and visceral circular muscles is inverted form the pharynx to the intestine (inner circulars and outer longitudinals versus inner longitudinals and outer circulars, see Figs. 1.9 B–C, 1.11 C–D). Both circular and longitudinal muscle components of the gut tube are enwrapped with a muscular double helix. Such fibers are crossing on the dorsal, ventral, and lateral sides of the gut tube (Figs. 1.9 A, 1.10, 1.11 A–B, 1.12 B). The helicoidally arranged muscles do not span the whole intestine down to the anus but only reach the midtrunk region in most species (e.g., Hochberg & Litvaitis 2001b, 2003a). Such a pattern can also be assumed for the stem species of Gastrotricha. In a somatic, ventrolateral position, there is a pair of massive longitudinally arranged muscle bands composed of several closely arran-

1

1

0

0

0

1

Pseudostomella roscovita Tetranchyroderma papii

1

1

Turbanella 1 sp. Acanthodasys 0 aculeatus

0

0

1

1

1

0

0

1

Turbanella ambronensis

1

1

1

1

1

Paradasys sp. Lepidodasys ligni

0

1

1

Dolichodasys 1 elongatus

1

Semicircular muscle band on ventral side of pharynx

3

Macrodasys caudatus

1

1

Dactylopodola baltica

2

Somatic Visceral circular circular muscles muscles on pharynx

1

1

1

1

1

1

1

?

1

1

1

Visceral circular muscles on intestine (midgut)

4

1

1

1

1

1

1

1

1

1

1

Visceral longitudinal muscles on dorsal side of gut tube

5

1

1

1

1

1

1

1

1

1

1

Visceral longitudinal muscles on ventral side of gut tube

6

Character-number according to Hochberg & Litvaitis (2001b)

1

1

1

1

1

1

1

1

1

1

Somatic longitudinal muscle bands in ventrolateral position (musculus principalis according to Remane 1936)

7

Tab. 1.1: Characters related with musculature and muscle arrangement in Macrodasyida.

0

0

1

1

1

?

0

1

1

1

Splitting of musculus principalis in midtrunk region

8

On mouth rim

On mouth rim

At level of TbA On mouth rim

At level of TbA

On mouth rim

At level of TbA On mouth rim

At level of TbA

At level of TbA

Anterior insertion of musculus principalis

9

1

1

0

0

1b

0

0

0

0

1

Crossover muscles in caudal region

10

0

0

0

1

1

0

0

0

0

1

Branches of musculus principalis supply head region

11

1

1

1

1

1

1

1

1

1

1

Helicoidal muscles enwrap pharynx

12

1

1

1

1

0

0

1

1a

0

1

Helicoidal muscles enwrap intestine (midgut)

13

Oblique striation

Oblique striation

Oblique striation Oblique striation

Oblique striation

Oblique striation

Oblique striation Oblique striation

Oblique striation

Crossstriated

Muscle striation pattern (see also Tab. 1.3)

14

Hochberg & Litvaitis (2001b) Hochberg & Litvaitis (2001b) Leasi et al. (2006) Hochberg et al. (2013) Hochberg & Litvaitis (2001b) Hochberg & Litvaitis (2001b) Leasi et al. (2006) Hochberg & Litvaitis (2001b) Hochberg & Litvaitis (2001b) Hochberg & Litvaitis (2001b)

Source

1.2 Morphology 

 11

?

1

1

1

0

1

1

0

0

0

0

0

Semicircular muscle band on ventral side of pharynx

3

1

?

1

1

Visceral circular muscles on intestine (midgut)

4

1

1

1

1

Visceral longitudinal muscles on dorsal side of gut tube

5

1

1

1

1

Visceral longitudinal muscles on ventral side of gut tube

6

1

1

1

1

Somatic longitudinal muscle bands in ventrolateral position (musculus principalis according to Remane 1936)

7

?

1

0

0

Splitting of musculus principalis in midtrunk region

8

On mouth rim

On mouth rim

On mouth rim

On mouth rim

Anterior insertion of musculus principalis

9

0

0

1

1

Crossover muscles in caudal region

10

1

0

0

0

Branches of musculus principalis supply head region

11

1

1

1

1

Helicoidal muscles enwrap pharynx

12

1

1

1

1

Helicoidal muscles enwrap intestine (midgut)

13

Crossstriated

Oblique striation

Oblique striation

Oblique striation

Muscle striation pattern (see also Tab. 13.2.3.3)

14

Modified and amended from Hochberg & Litvaitis (2001b). A question mark (?) indicates an unknown character state; 0, absence; 1, presence; TbA, anterior adhesive tubes. a There is only one crossing of the helicoidal muscle on the intestine in Paradasys sp. (see Leasi et al. 2006). b Hochberg & Litvaitis (2001b) code the crossover muscle as “present” for T. ambronensis, although in the text, they report its absence.

Neodasys cirritus

Tetranchyroderma megastoma Thaumastoderma heideri Oregodasys cirratus

2

Somatic Visceral circular circular muscles muscles on pharynx

1

Character-number according to Hochberg & Litvaitis (2001b)

Tab. 1.1 (Continued)

Hochberg & Litvaitis (2001b) Hochberg & Litvaitis (2001b) Rothe & SchmidtRhaesa (2010) Hochberg (2005)

Source

12   1 Gastrotricha

1.2 Morphology 

 13

vc

A

hm

ph

ph

B

br

vc vlm hm in

vl

sc

vl

sc

vc

go

in

hm

go

vl sc vlm

vc

vlm

vc

hm

ph

sc

vl

vlm

C Fig. 1.9: Muscular system (schematic) of the last common ancestor of Gastrotricha. (A) Myoanatomy, dorsal view. (B and C) Trunk cross sections at different levels (indicated by bold lines). Note the reversal of the sequence of visceral circular and longitudinal muscles from pharyngeal (B) to intestinal (C) region. It is not sure if the stem species, like many extant gastrotrichs, possessed a splitting of the ventrolateral muscle bands in the midtrunk region (left body side in A and C). Abbreviations: br, brain; go, gonads; hm, helicoidal muscle; in, intestine; ph, pharynx; sc, somatic circular muscle; vc, visceral circular muscle; vl, visceral longitudinal muscles; vlm, ventrolateral muscle bands (musculi principales).

in

vc

vl

Fig. 1.10: Muscular system (schematic) of the last common ancestor of Gastrotricha, lateral view. Note the reversal of the sequence of visceral circular and longitudinal muscles from pharyngeal to intestinal region. Abbreviations: hm, helicoidal muscle; in, intestine; ph, pharynx; sc, somatic circular muscles; vc, visceral circular muscles; vl, visceral longitudinal muscles; vlm, ventrolateral muscle band (musculus principalis).

14 

 1 Gastrotricha

ged muscle fibers (Fig. 1.12 A–D). These paired musculi principales (according to Remane 1936, but see the notes on this terminology in Teuchert 1974) or ventrolateral muscle bands insert anteriorly close to the mouth rim and reach the caudal lobes that bear the posterior adhesive tubes (the stem species of Gastrotricha was characterized by a bilobed caudal trunk end according to Kieneke et al. 2008a). Along the whole body, there are separate somatic circular muscles that surround the ventrolateral muscle bands and probably all other muscular components (Figs. 1.9 and 1.10). Yet, it is not clear if these somatic circular muscles of the stem species of Gastrotricha represent closed rings. In many species of the Macrodasyida, the somatic circular muscles are reported to enclose the ventrolateral muscle bands “on either side of the midgut”. Such a description, in addition with the presented data, indicates the presence of incomplete muscle rings at least in these species (e.g., Hochberg & Litvaitis 2001b, Leasi et  al. 2006). In Neodasys, the somatic circular muscles are branches of the visceral circular muscles that line the gut tube (Hochberg 2005). Such a condition is generally regarded to demonstrate the evolutionary origin of the somatic circular muscles (see Leasi & Todaro 2008 and references therein). As for the stem species, there are not data on the exact numbers concerning different muscular components such as the number of visceral and somatic circular muscles, of visceral longitudinal muscles, of crossings of the helicoidal muscles, and of muscle fibers per ventrolateral muscle band. It remains to mention that there are diverse modifications (e.g., reductions, losses, branching patterns, additional muscle components) from the previously described ancestral muscular character pattern in extant species of Gastrotricha (see Tab. 1.1). These include, for instance, the presence of a peculiar semicircular muscle band on the ventral side of the pharynx in taxa such as Dactylopodola baltica, Paradasys sp., or Turbanella (Hochberg & Litvaitis 2001b, Leasi et al. 2006). When present, this muscle connects both ventrolateral muscle bands (musculi principales) close to the level of the anterior adhesive tubes (see Fig. 1.24 C). A comparable (homologous?) muscle is also present in the basal paucitubulatinan species Musellifer delamarei (Leasi & Todaro 2008, see Tab. 1.2). Additionally, there might be one or two so-called crossover muscles connecting both parts of musculi principales in the caudal region of many but not all species with a bilobed caudal end (Hochberg & Litvaitis 2001b, c, Tab. 1.1). The crossover muscle results from a splitting of one or few fibers of the ventrolateral muscle band passing over to the other side and vice versa (Hochberg & Litvaitis 2001c). There is a splitting of each ventrolateral muscle band in the midtrunk region in

several species with paired testes: Some fibers run more laterally (pars lateralis according to Remane 1936), while others are located more medially (pars ventrolateralis according to Remane 1936). Further posterior, all fibers converge forming a common muscle band (e.g., in Dactylopodola baltica, Paradasys sp., Turbanella sp., or in Crasiella fonseci, see Hochberg & Litvaitis 2001b, Leasi et al. 2006, Hochberg 2014). Such kind of splitting is mostly absent in members of the Thaumastodermatinae that only possess one single testis (Hochberg & Litvaitis 2001b, but see Rothe & Schmidt-Rhaesa 2010 for the situation in Oregodasys cirratus). This taxon is further characterized by complete absence of somatic circular muscles (Hochberg & Litvaitis 2001b, c). An exception to this pattern is Oregodasys cirratus (Rothe & Schmidt-Rhaesa 2010, Tab. 1.1). A peculiarity of whole Thaumastodermatidae is the formation of anterior branches of dorsal visceral longitudinal muscles that spread out into the oral hood (Remane 1936, Hochberg & Litvaitis 2001b, c). These muscle branches facilitate the withdrawal of the oral hood (Remane 1936). Additional variation in muscle organization concerns the anterior insertion of the ventrolateral muscle bands. In species that possess distinct, frequently hand-like anterior adhesive organs composed of several closely arranged adhesive tubes (e.g., taxa Turbanella and Paradasys), the musculi principales terminate in close proximity to these organs and hence a certain distance posterior to the anterior end. In species that do not possess those anterior adhesive organs but instead have separately arranged adhesive tubes close to the anterior trunk end (e.g., Lepidodasys, Macrodasys, Thaumastodermatidae), the ventrolateral muscle bands insert close to the mouth rim (Tab. 1.1). Species such as Dactylopodola baltica and species of Turbanella and Neodasys have thin branches of the ventrolateral muscle bands in the anterior region that supply the head (Hochberg & Litvaitis 2001b, Hochberg 2005, Tab. 1.1). These are different from the muscle branches within the head region (oral hood) of the Thaumastodermatidae, which are formations of the dorsal visceral longitudinal muscles (see above). Severe evolutionary modifications of the muscular system occurred both along the stem lineage of the gastrotrich subtaxon Paucitubulatina and within this clade. Through fluorescence staining and confocal or epifluorescence microscopy, the musculature of several species of this group has been studied so far (Hochberg & Litvaitis 2001d, 2003a, Leasi et al. 2006, Kieneke et al. 2008b, Kieneke & Ostmann 2012, Leasi & Todaro 2008, 2009). In contrast to the Macrodasyida and Neodasys, the number and arrangement of longitudinal muscles, somatic and visceral, is much more determined in Paucitubulatina.

1.2 Morphology 

Pairs of potentially homologous longitudinal muscles that have been detected in all hitherto investigated species of the Paucitubulatina are the musculi ventrales, m. ventrolaterales, m. laterales, m. dorsales, and one or two pairs of dorsodermal muscles that are branches of musculi dorsales (Hochberg & Litvaitis 2001d, 2003a, Leasi & Todaro 2008, Tab. 1.2, Figs. 1.11, 1.12 E, F). Although this is not based on a thorough phylogenetic reconstruction, we conclude that these five to six longitudinal muscle pairs probably belong to the character pattern of the stem species of Paucitubulatina (this clade is most likely a monophyletic group, see chapter Phylogeny). As most investigated basal species of Paucitubulatina (Musellifer delamarei, Draculiciteria tesselata, and most species of Xenotrichulinae) possess only one dorsodermal branch of musculi dorsales (“Rückenhautmuskel” according to Zelinka 1889, see Hochberg & Litvaitis 2003a), we suspect that this represents the ancestral condition (Figs. 1.11 and 1.12 F). Owing to their more peripheral position in the body, musculi laterales and dorsodermal muscles are considered to represent somatic components, whereas all other longitudinal muscles are visceral components (Hochberg & Litvaitis 2001d, 2003a). However, the assignment to one or the other group of musculature – visceral or somatic – is not always that explicit, and it seems that in different members of the Xenotrichulidae, the musculi ventrolaterales

A

C

ph

hm

vc

vc

ml

mvl mv

hm md

ddm vc

D

hm md

sc go

in ml

in

hm

go

mvl ddm

rather belong to the somatic musculature (see results of Leasi & Todaro 2008). Similar names imply a homology of these muscles, which may not always be the case. For example, it is likely that the musculi ventrolaterales of Paucitubulatina are not homologous to the ventrolateral muscle bands (musculi principales) in Macrodasyida and Neodasys. It is more likely that the somatic musculi laterales of Paucitubulatina are homologous to the ventrolateral muscle bands of Macrodasyida and Neodasys or at least to one or two of their fibers. In addition to the aforementioned five to six pairs of longitudinal muscles, there may be further pairs. In Xenotrichulinae, for instance, there is always a visceral pair of musculi ventromediales that spans between m. ventrales and m. ventrolaterales for most of its course (Hochberg & Litvaitis 2003a, Leasi & Todaro 2008, 2009). In Draculiciteria tesselata, there is another somatic (?) longitudinal muscle pair, the musculi paralaterales. This muscle spans between m. laterales and m. ventrolaterales (Hochberg & Litvaitis 2001d). The basal paucitubulatinan species Musellifer delamarei has additional somatic longitudinal muscles: There are two pairs of broad muscle bands running along the body wall in dorsal and ventral positions (Leasi & Todaro 2008). Further differences between species can be found in the specific course of certain longitudinal muscles. Especially the musculi

B

md

br

ph

 15

mv sc

Fig. 1.11: Muscular system (schematic) of the last common ancestor of Paucitubulatina. Myoanatomy in (A) dorsal and (B) ventral views. (C and D) Trunk cross sections at different levels (sectional plains indicated by bold lines). Note the reversal of the sequence of visceral circular and longitudinal muscles from pharyngeal (C) to intestinal (D) region. Abbreviations: br, brain; ddm, dorsodermal muscle; go, gonads; hm, helicoidal muscle; in, intestine; md, musculus dorsalis; ml, musculus lateralis; mv, musculus ventralis; mvl, musculus ventrolateralis; ph, pharynx; sc, somatic circular muscle; vc, visceral circular muscle.

 1 Gastrotricha

A

B

* mlv

C

D

50 µm

16 

mlv ph

vlm mcv mcv

vlm

vlm

hm pp

*

50 µm

E 100 µm

* ph

ml

om

50 µm

ddm

F

md

ph dvm

75 µm

ml

50 µm Fig. 1.12: Muscular system of Gastrotricha. Maximum projections of confocal image stacks. F-actin was stained with fluorescence-labeled phalloidin. (A) Turbanella ambronensis (Macrodasyida), horizontal view. (B–D) Turbanella hyalina, horizontal views. Note the stained filaments inside the adhesive tubes. (B) Anterior end. Note the sphincter-like circular muscles at the anterior and posterior end of the pharynx (asterisks). (C) Rear trunk end. (D) Whole specimen. (E) Dasydytes goniathrix (Paucitubulatina) with a highly derived somatic musculature consisting of oblique and segmented longitudinal muscles that are used for moving the long cuticular spines (asterisk, autofluorescence). Lateral view, specimen slightly tilted. (F) Xenotrichula velox, a rather primitive species of the Paucitubulatina that still has dorsoventral muscles in a visceral and somatic position, lateral view. Abbreviations: ddm, dorsodermal muscle; dvm, dorsoventral muscles; hm, helicoidal muscles; mcv, visceral circular muscles; md, musculus dorsalis; ml, musculus lateralis (of Paucitubulatina); mlv, visceral longitudinal muscles; om, oblique muscles; ph, myoepithelial pharynx; pp, pharyngeal pore; vlm, ventrolateral muscle blocks of Macrodasyida (musculi principales).

Dactylopodola baltica Neodasys cirritus Draculiciteria tesselata Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata Aspidiophorus marinus Chaetonotus sp. Halichaetonotus sp. 1/H. aculifer Lepidodermella squamata Musellifer delamarei Polymerurus nodicaudus

Dactylopodola baltica Neodasys cirritus Draculiciteria tesselata Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata Aspidiophorus marinus Chaetonotus sp. Halichaetonotus sp. 1/H. aculifer Lepidodermella squamata Musellifer delamarei Polymerurus nodicaudus Distant to mouth Close to mouthb Close to mouth Close to mouth Close to mouth Close to mouth Close to mouth Close to mouth Close to mouth Close to mouth Distant to mouth Close to mouth

1 1 1 1 1 1 0 0 0 0 1 0

0 0 1 1 1 1 0 0 0 0 0 0

Cross-striation Atypical cross-striation Atypical cross-striation Oblique striation Oblique striation Oblique striation Oblique striation Oblique striation Oblique striation Oblique striation Atypical cross-striation Oblique striation

Striation pattern

Posterior branching of musculus dorsalis with crossing

Dorsodermal muscle (branch of visceral musculus dorsalis)

0 0 1 1 1 1 1 1 1 1 1 1

6

5

– – 1 1 1 1 1 1 1 1 1 1

– – 0 1 1 1 0 0 0 0 0 0

Visceral musculi ventromediales

­

Longer than one third of intestine To one third of intestine Longer than one third of intestine Longer than one third of intestine Longer than one third of intestine To one third of intestine To one third of intestine To one third of intestine To one third of intestine To one third of intestine To the base of pharynx To one third of intestine

Extent of helicoidal muscles

7

Visceral musculi ventrolaterales

Visceral musculi paralaterales – – 1 0 0 0 0 0 0 0 0 0

­

­

4

Character-number according to Leasi & Todaro (2008)

>2 >2 1–2 1–2 1–2 1–2 1–2 1–2 1–2 1–2 >2 1–2

Anterior insertion Muscles in the of somatic longitu- furca/caudal lobes dinal musclesa

Number of myocytes (fibers) per somatic longitudinal musclea

3

2

1

Character-number according to Leasi & Todaro (2008)

Tab. 1.2: Characters related with musculature and muscle arrangement in Paucitubulatina.

– – 1 1 1 1 1 1 1 1 1 1

Visceral musculus dorsalis

­

(Continued)

Complete circular Complete circular Complete dorsoventral Complete dorsoventral Incomplete circular Incomplete dorsoventral Absent Absent Absent Absent Incomplete circular Complete dorsoventral

Visceral muscles in the intestinal region

8+9

– – 1 1 1 1 1 1 1 1 1 1

Visceral musculi ventrales

­

1.2 Morphology 

 17

11

Branched ­– ­– ­– –­ Not branched Branched

Xenotrichula punctata Aspidiophorus marinus Chaetonotus sp. Halichaetonotus sp. 1/H. aculifer Lepidodermella squamata Musellifer delamarei Polymerurus nodicaudus 0 0 0 0 0 1 0

1 0 0 0 0 Incomplete dorsoventral Absent Absent Absent Absent Incomplete dorso-ventralc Absent

Circular Circular Complete dorsoventral Complete dorsoventral Incomplete dorsoventral

Somatic muscles in the intestinal region

12 + 13

1 0 0 0 0 0 0

0 0 1 1 1

Somatic dorsoventral musclee pairs close to the pharyngeointestinal junction

14

Hochberg & Litvaitis (2001b) Hochberg (2005) Hochberg & Litvaitis (2001d) Leasi & Todaro (2008) Hochberg & Litvaitis (2003a), Leasi & Todaro (2008) Leasi & Todaro (2008) Hochberg & Litvaitis (2003a) Leasi & Todaro (2008) Hochberg & Litvaitis (2003a) Hochberg & Litvaitis (2003a) Leasi & Todaro (2008) Leasi et al. (2006)

Source

Modified and amended from Leasi & Todaro (2008). Dactylopodola baltica and Neodasys cirritus were used for the outgroup comparison. A question mark (?) indicates an unknown character state; dash (–), inapplicable character state; 0, absence; 1, presence. a The “somatic longitudinal muscles” are the ventrolateral muscle bands (musculus principalis) in Macrodasyida and Neodasys, called musculi laterales in Paucitubulatina according to Hochberg & Litvaitis (2003a). b Leasi & Todaro (2008) coded the longitudinal muscle insertion in N. cirritus distant to the mouth, although original data (Hochberg 2005) demonstrate a different pattern. c Leasi & Todaro (2008) coded the somatic muscles in the intestinal region of M. delamarei as “incomplete dorsoventral”, although they are later treated as “incomplete circular” (see, e.g., their figure 7).

–­ –­ Branched Branched Branched

Termination of circular or Semicircular muscle dorsoventral muscles in the band(s) on ventral intestinal region side of pharynx

10

Character-number according to Leasi & Todaro (2008)

Dactylopodola baltica Neodasys cirritus Draculiciteria tesselata Heteroxenotrichula squamosa Xenotrichula intermedia

Tab. 1.2: ( Continued)

18   1 Gastrotricha

1.2 Morphology 

laterales and m. ventrolaterales may run in close proximity to the pharynx in certain species, whereas in others, they follow the contours of the anterior body or continuously diverge toward the anterior end (compare reconstructions of Aspidiophorus marinus, Chaetonotus spp., Halichaetonotus spp., and Lepidodermella spp. in Hochberg & Litvaitis 2003a). In Xenotrichula intermedia, musculi ventrales are reported to cross over in the region of the anus (Hochberg & Litvaitis 2003a). However, such a crossover was not confirmed for X. intermedia and other members of the Xenotrichulinae (Leasi & Todaro 2008). Meanwhile, a peculiar x-shaped connection between the paired musculi dorsales in the posterior trunk region was detected in all species of Xenotrichulidae (Xenotrichulinae plus Draculiciteria tesselata) studied so far (Leasi & Todaro 2008, Tab. 1.2). In the pharyngeal region of Paucitubulatina, like in Macrodasyida and Neodasys, there are densely piled complete circular muscles that line the pharynx inward the visceral longitudinal muscle components. However, such muscle rings could not be detected with certainty in species of the derived paucitubulatinan taxon Dasydytidae (see Kieneke et al. 2008b, Kieneke & Ostmann 2012). In Musellifer delamarei, a semicircular muscle band on the ventral side of the pharynx was found, which is comparable to those muscles discovered in Dactylopodola baltica and other species of Macrodasyida (Leasi & Todaro 2008, compare Tab. 1.1 with Tab. 1.2). Even more structural diversity can be observed among the “circular” muscle components in the intestinal region of Paucitubulatina (Hochberg & Litvaitis 2001d, 2003a, Leasi & Todaro 2008, 2009, Tab. 1.2). In the assumed basal species M. delamarei, there are incomplete visceral circular muscles plus somatic dorsoventral muscles. “Incomplete” refers to the fact that these circular muscles do not represent closed rings but have a median gap dorsally and ventrally (Leasi & Todaro 2008). Some species of the Xenotrichulinae show a comparable muscle arrangement: The visceral component is represented by incomplete circular muscles, whereas the somatic component consists of dorsoventral muscle fibers that attach to the dorsal and ventral integument. In Draculiciteria tesselata and Heteroxenotrichula squamosa, the somatic and visceral components consist of such dorsoventral muscles. In the lineage that leads to the predominantly freshwater inhabiting Chaetonotidae (much likely this family does not represent a monophyletic group, see chapter Phylogeny), only assumed basal species such as Polymerurus nodicaudus possess comparable muscle components: This species still has dorsoventral muscles in a visceral position in its intestinal region, whereas any circular or dorsoventral components are absent in the somatic position (Leasi et al. 2006, Leasi & Todaro 2008, Tab. 1.2).

 19

Other taxa of the Chaetonotidae like Aspidiophorus, Chaetonotus, Halichaetonotus, and Lepidodermella completely lack circular or dorsoventral muscles apart from a tiny single pair close to the anus (Hochberg & Litvaitis 2003a, Leasi & Todaro 2008). Also, species of the exclusively freshwater-dwelling Dasydytidae lack any circular muscle components in the intestinal region (Kieneke et al. 2008b, Kieneke & Ostmann 2012), a result that partially revises the muscular description of Dasydytes ornatus by Remane (1936), who suspected the presence of incomplete circular muscles in that species. The Paucitubulatina are another example for muscular diversity: The muscle endings are simple in M. delamarei, whereas all investigated Xenotrichulidae and P. nodicaudus have branched muscle endings (Tab. 1.2). We can deduce from the evolutionary scenario of circular muscle evolution in Paucitubulatina developed by Leasi & Todaro (2008), that the last common ancestor of Paucitubulatina could have had a system of incomplete circular muscles in visceral and somatic positions in its intestinal region in addition to the longitudinal muscle pairs discussed above (Fig. 1.11). The presence of incomplete circular muscles in a somatic position, however, is purely hypothetical and has never been observed in any extant species of the Paucitubulatina (Leasi & Todaro 2008). Because a visceral helicoidal musculature is present in all investigated species of Paucitubulatina studied so far (Hochberg & Litvaitis 2001d, 2003a, Leasi et al. 2006, Kieneke et  al. 2008b, Kieneke & Ostmann 2012, Leasi & Todaro 2008, 2009), this muscular component is an ancestral paucitubulatinan character, too. The taxon Dasydytidae is a highly derived group of freshwater-dwelling planktonic gastrotrichs (almost all other species of Paucitubulatina have an endobenthic, epibenthic, or periphytic lifestyle, see chapter Ecology), which have paired groups of motile cuticular spines. These spines are actively movable and can be abducted and adducted serving either for supporting locomotion (species such as Haltidytes crassus may perform short “jumps” in the water column) or for performing defensive positions at which the animals take a strong ventral flexion of the trunk and abduct their spines to a maximum. Spine movement is brought about by a highly specialized musculature consisting of serially arranged somatic oblique muscles and segments of the partitioned musculi laterales (Kieneke et  al. 2008b, Kieneke & Ostmann 2012; Fig. 1.12 E). Although it was initially supposed that oblique and segmented longitudinal muscles represent elements of an antagonistically working system (Kieneke et  al. 2008b), it is more likely that both muscle components work synergistically to facilitate the spine abduction-adduction cycle (Kieneke & Ostmann 2012). It is not likely that the oblique musculature of Dasydytidae

20 

 1 Gastrotricha

is a derivate of ancestral somatic circular muscles because not a single investigated species of the Chaetonotidae, among them putative relatives of the Dasydytidae (see, e.g., Kånneby et  al. 2013), possesses a somatic circular musculature. Somatic circular (or dorsoventral) musculature, still present in Muselliferidae and Xenotrichulidae (see above), hence must have been lost within the lineage that leads to Chaetonotidae, Dasydytidae, and some other minor taxa of freshwater-dwelling gastrotrichs. Oblique musculature therefore represents a new evolutionary formation, an autapomorphy of the Dasydytidae (Kieneke & Ostmann 2012). Apart from the visceral and somatic musculature of Gastrotricha that is present as longitudinal, circular/dorsoventral, and helicoidal muscles (see above), there may be further muscle arrangements. Most prominent is a specialized musculature as part of the reproductive system. Taxa that possess a distinct caudal organ as a sperm-transferring device (see chapter Reproductive Organs) may have a strong circularly or slightly helically arranged musculature that surrounds this accessory reproductive organ as, for example, in Macrodasys sp. (Ruppert 1978a), Tetranchyroderma papii (Hochberg & Litvaitis 2001c), or in Lepidodasys ligni (Hochberg et al. 2013). Muscle contractions of the caudal organ are used to support the release of spermatozoa from the caudal organ lumen or, as in Macrodasys sp., to evert the copulatory tube (Ruppert 1978a, see also chapters Reproductive Organs and Reproductive Biology). However, there are also species that contain a caudal organ but obviously lack a specialized musculature like Crasiella fonseci (Hochberg 2014), although congeneric species, e.g., C. diplura, possess such a circular muscle sheath (Guidi et al. 2011). It is hypothesized that caudal organ musculature of C. fonseci may be formed rather late during an individual’s development (Hochberg 2014). A sheath of circular muscles also surrounds the distal parts of the vas deferens in species of Thaumastodermatidae (Ruppert 1978b). A comparable musculature on the distal section of the sperm ducts may be present in species of Turbanella (see figure 2c of Leasi et  al. 2006). A narrow, ring-shaped muscle at the level of the anus was reported from different species. This single circular muscle probably represents an anal sphincter (see, e.g., Leasi et al. 2006, Kieneke et al. 2008b). A sphincter oris surrounding the mouth opening is reported for different species of the Paucitubulatina (Remane 1936). A strong, sphincter-like circular muscle is also present around the mouth opening of the macrodasyid Crasiella fonseci (Hochberg 2014) and in species of Turbanella (Fig. 1.12 A–B). The ultrastructure of musculature and muscle cells was intensively studied in the marine gastrotrich Turbanella cornuta (Teuchert 1974). These findings were later complemented by ultrastructural data of several other

species from all of the three major gastrotrich subtaxa Macrodasyida, Neodasys (Multitubulatina), and Paucitubulatina (Ruppert 1991, see Tab. 1.3). Longitudinal muscle cells of T. cornuta are to 40 µm long, spindleshaped, and with an axially situated cytoplasmic compartment at one end that houses the nucleus (see Fig. 1.13 A for longitudinal muscles in Polymerurus). In cross section, such longitudinal muscle cells are somehow leaf-shaped with a coelomyarian to ribbon-like arrangement of contractile elements. The ventrolateral longitudinal muscle bands of T. cornuta (musculi principales, but see Teuchert 1974 for concerns about this terminology) consist each, in cross section, of nine fibers; each fiber is composed of 8–12 consecutive mononuclear cells (Teuchert 1974). Circular muscle cells of T. cornuta are as well spindle-shaped (approximately 10 µm long) but with an abaxially positioned nucleus (Teuchert 1974). An abaxial position of the nucleus, however, was also found in longitudinal muscle cells of Neodasys sp. (Ruppert 1991; Fig. 1.13 B–C). Variation among the ultrastructure and cytomorphology of gastrotrich muscle cells can be found, for instance, in the striation pattern, type of z-material, and the absence/presence of a t-system for the excitationcontraction-coupling (Ruppert 1991, Tab. 1.3). A peripheral coupling of the muscle cell membrane (sarcolemma) with the sarcoplasmic reticulum was confirmed for almost all species investigated so far. Interestingly, taxa that are generally regarded to occupy rather basal positions within the phylogenetic tree of the Gastrotricha such as Neodasys, Dactylopodola, Xenodasys, Chordodasiopsis, Draculiciteria, and Musellifer (see chapter Phylogeny) possess a cross-striated musculature, whereas most remaining taxa show an oblique striation pattern (Ruppert 1991, Tab. 1.3). Because Neodasys, Musellifer, and Draculiciteria have rods instead of dense bodies as z-material, their striation type is regarded as “atypical cross-striation”. If basal positions of the aforementioned taxa with crossstriated musculature will be supported (but see chapter Phylogeny for differing phylogenetic scenarios), this striation pattern would be part of the character pattern of the stem species of Gastrotricha as the most parsimonious reconstruction (Ruppert 1991). So far, Lepidodasys is the only known gastrotrich taxon with a smooth muscle organization (Ruppert 1991). Except Lepidodasys all gastrotrichs have radially arranged, cross-striated myofibrils in the myoepithelial pharynx (Fig. 1.13 D). Regarding the sarcomeres per contractile element of the pharynx, the number varies from one in Lepidodermella squamata up to 12 sarcomeres in Turbanella cornuta (Ruppert 1982). The mechanic coupling between neighboring muscle cells or a muscle cell and a non-muscle cell is by adhaerens

Circomyarian to polygonal

Circomyarian to polygonal Circomyarian to polygonal ? Coelomyarian/ribbon-like Coelomyarian/ribbon-like Coelomyarian/ribbon-like Coelomyarian/ribbon-like Coelomyarian/ribbon-like Coelomyarian/ribbon-like Polygonal Polygonal to coelomyarian Polygonal to coelomyarian ? Circomyarian to polygonal Circomyarian to polygonal Circomyarian to polygonal Circomyarian to polygonal Circomyarian to polygonal Polygonal to coelomyarian Coelomyarian/ribbon-like Circomyarian to coelomyarian Coelomyarian/ribbon-like

7 × 3 µm

6.0 µm 7.5 µm ? 3 × 1 µm 4.5 × 1.3 µm 3.4 × 1 µm 3.2 × 1.6 µm 12 × 1.7 µm 6 × 1 µm 4 × 2.5 µm 7 × 1.3 µm 5.3 × 1.5 µm ?

7 × 1.4 µm 1.4 µm 3.2 µm 3 × 1 µm 8.0 µm 3.0 µm 1.5 × 1.0 µm 4 × 1.3 µm 4.0 × 1.0 µm

0 1 1 1 1 0 0 0 0

1 1 0 0 0 0 0 0 0 0 0 0 0

1

1 0 0 0 0 1 1 1 1

0 0 1 1 1 1 1 1 1 1 1 1 1

0

1 0 0 0 0 0 0 0 0

0 0 0 0 0 0 0 0 0 0 0 0 0

0

? 2.5 µm ? ? ? ? ? ? ?

2.0 µm 1.0 µm ? ? ? ? ? ? 1.8 µm ? ? ? ?

0.8 µm

17–100 nm 32 nm 26 nm 21 nm ? ? ? 31 nm 23 nm

30 nm ? ? 33 nm 33 nm 28 nm 28 nm 27 nm 32 nm ? 39 nm ? ?

?

Cross-striated Oblique- Smooth Length of Diameter myofibrils striated myofibrils sarcomere of myosin myofibrils

0 0 1 ? ? 0 0 0 0

1 1 0 0 0 0 0 0 0 0 0 0 0

1

1 1 0 ? ? 1 1 1 1

0 0 1 1 1 1 1 1 1 1 1 1 1

0

1 1 1 1 1 1 1 1 1

1 1 1 1 1 1 1 1 1 (?) 1 1 1 1

?

0 0 1 1 1 0 0 0 0

1 1 0 0 0 0 0 0 0 0 0 0 0

1 (?)

Dense Rods as Peripheral coupT-system bodies as Z-material lings of sarcolemma Z-material and sarcoplasmic reticulum

Modified from Ruppert (1991). A question mark (?) refers to unknown data; 0, absence; 1, presence. a Ruppert (1991) presented muscular characters of an undescribed species of an undescribed genus referring to a drawing of his gastrotrich chapter in Ruppert (1988). Todaro et al. (2005) identified this animal as belonging to the taxon Diuronotus.

Chordodasiopsis antennatus (former Xenodasys antennatus) Xenodasys riedli Dactylopodola sp. Cephalodasys sp. Cephalodasys littoralis Crasiella diplura Macrodasys sp. Mesodasys sp. Paraturbanella sp. Turbanella cornuta Dolichodasys carolinensis Acanthodasys sp. Thaumastoderma sp. Oregodasys sp. (former Platydasys) Lepidodasys sp. Neodasys sp. Diuronotus sp.a Draculiciteria tesselata Musellifer sublittoralis Chaetonotus sp. Lepidodermella squamata Xenotrichula carolinensis Aspidiophorus sp.

Diameter of Arrangement of sarcomeres single within fiber (as seen when muscle fiber cross-sectioned)

Tab. 1.3: Characters related with the ultrastructure and cytomorphology of longitudinal muscle cells of various species of the Gastrotricha.

1.2 Morphology 

 21

22 

 1 Gastrotricha

A

B mi

sr

pl mi

nu

ml

mg

ecm

cc 1 µm

C

F

1 µm

D

cm

lm lnb

lm

phl

ac

mf 0.5 µm

junctions, most probably desmosomes. If a muscle inserts on the body wall, muscle cells do not attach directly to the cuticle, but always to an epidermal cell. Tension between muscle and cuticle is provided by microfilaments that span through the epidermis cell from adhaerens junctions between muscle and epidermal cell to hemidesmosomes between epidermal cell and cuticle (Ruppert 1991). A comparable situation is assumed for the mechanic coupling between the sections of partitioned musculi laterales/ oblique muscles and the movable spines in Dasydytidae (Kieneke et al. 2008b). This, however, has to be supported by ultrastructural studies in the future. Mechanic coupling of contractile elements (myofibrils) and the cuticle is slightly different in the pharynx (see also chapter Intestinal System for this issue). Here, the myofibrils of the myoepithelial cells are directly attached to the apical pharyngeal cuticle via specialized, plaque-like hemidesmosomes. Hemidesmosomes also attach the myofibrils basally to the basal lamina. There is quite a high ultrastructural diversity among hemidesmosomes of the pharynges of different species and groups of the Gastrotricha (see

1 µm

Fig. 1.13: Muscle ultrastructure (TEM-cross sections) of Gastrotricha. (A) Polymerurus nodicaudus (Paucitubulatina). Somatic longitudinal musculature in close proximity to the protonephridium. (TEM micrograph from Kieneke & Hochberg 2012, with permission by Wiley.) (B–D) Neodasys chaetonotoideus (Multitubulatina). (B) Cell body of a longitudinal muscle cell with nucleus. (C) Longitudinal muscle cells and nervous system in close proximity. (D) Detail of the pharynx showing myoepithelial cells and subpharyngeal visceral musculature. Abbreviations: ac, apical cell; cc, canal cell of the protonephridium; cm, circular muscles; ecm, extracellular matrix; lm, longitudinal muscles; lnb, longitudinal neurite bundle; mf, crossstriated myofilaments; mg, midgut; mi, mitochondria; ml, musculus lateralis; nu, nucleus; phl, pharyngeal lumen; pl, protonephridial lumen; sr, sarcoplasmic reticulum.

Ruppert 1982 for details). Some of the basal attachments of the pharyngeal myofibrils are furthermore mechanically coupled to the body cuticle via intracellular fibers that may span through different layers of cells before reaching the cuticle (Ruppert 1991). The ultrastructure of nerve-tomuscle connections for signal transduction was investigated in Turbanella cornuta by Teuchert (1977a). A peculiarity of these myoneural synapses is that the muscle cell itself forms one or few short processes that project into the neighboring nerve cell (Teuchert 1977a). Whether this represents a general pattern of the Gastrotricha, however, needs to be confirmed by further investigations (see Fig. 1.13 C for a close proximity between nervous and muscular cells). Somatic circular and longitudinal muscles in Gastrotricha are generally regarded as reciprocal antagonists (Ruppert 1991). Contractions of the longitudinal muscles cause a shortening of the body, whereas its diameter has to increase because of the unchanged volume of the animal and incompressibility of liquid (Fig. 1.14 A–E). Owing to the increasing diameter, the circular muscles will relax. If circular muscles contract, the diameter will decrease again that

1.2 Morphology 

leads to an elongation of the trunk and simultaneously to a relaxation of longitudinal muscles. Body movements of Gastrotricha that are brought about by muscle action include, for example, longitudinal elongation and shortening of the trunk (see above, Fig. 1.14 A–D), ventral and lateral flexion (Fig. 1.14 F), nodding and slightly turning the head, flexion of appendages (Fig. 1.14 G), or spreading of cuticular spines (e.g., Remane 1936, Ruppert 1991, Hochberg & Litvaitis 2001c, Kieneke et  al. 2008b). The main mode of locomotion in Gastrotricha is of course the cilia-mediated gliding or, in some taxa, swimming. Muscle action, however, severely aids ciliary gliding and swimming when, for instance, a lateral flexion of the whole trunk or lateral plus ventral/dorsal flexion of the head is used for controlling the direction of locomotion. Furthermore, appendages such as the caudal lobes or toes of many species of the Paucitubulatina or the already mentioned motile spines of Dasydytidae (see above) can actively be moved by muscle action and hence may be used as a “rudders” (Remane 1936, Kieneke & Ostmann 2012). Waving movements with the anterior trunk end are known from different species, e.g., Turbanella cornuta or Tetranchyroderma papii. These movements probably involve alternating contractions and relaxations of the ventrolateral muscle bands and the visceral longitudinal muscles (Remane 1936, Hochberg & Litvaitis 2001c).

A

B

100 µm

C

100 µm

E ph

100 µm

G *

mg

The waving of the head and anterior trunk is interpreted as a kind of “searching behavior”. Thus, the animal tries to optimize sensory perception (Remane 1936). A peculiar way of locomotion in many species of the Macrodasyida is shown during an escape behavior: Taxa such as Macrodasys, Turbanella, Paradasys, or Tetranchyroderma papii, just to mention some, may perform a leech-like or inchworm-like creeping, at which animals successively attach their posterior (or anterior) adhesive tubes to the substratum, then strongly flex or shorten their body, attach the adhesive tubes of the opposite body end (anterior or posterior tubes, respectively), and stretch the trunk again. Several quick repetitions of such actions allow a fast escape from a potential harmful stimulus, e.g., a collision with a predator or just with a scientist’s micropipette (Remane 1936, Hochberg & Litvaitis 2001c). There are interspecific differences in this special behavior. Macrodasys, for instance, escapes in a frontal direction, whereas taxa like Turbanella or Paradasys escape backward. Tetranchyroderma papii may engage in both, backward or forward directed inchworm-like creeping (Hochberg & Litvaitis 2001c). A similar escape behavior can be observed in Neodasys sp. However, these animals stay attached to a sand grain by means of their posterior adhesive organs and quickly contract the whole trunk accordion-like (Ruppert & Travis 1983).

D

100 µm

F

mg

an vlm

100 µm

200 µm

TbP

25 µm

yc

yc

 23

Fig. 1.14: Muscle-mediated movements of Gastrotricha. (A–D) Turbanella subterranea (Macrodasyida), dorsal views. The animal was consciously disturbed and showed a quick retraction of the trunk as a kind of escape response. (E) Fully retracted Turbanella subterranea with compressed organs such as the pharynx, midgut, and Y-cells. Due to incompressibility of liquids, the animal is much broader than usual. (F) A slipping Paradasys subterraneus (Macrodasyida) that takes a sharp right curve by contracting longitudinal muscles of its right side (asterisk). (G) Rear trunk of Dactylopodola deminuitubulata (Macrodasyida) with caudal pedicles. With the strong and cross-striated longitudinal musculature, species of Dactylopodola can perform rapidly repeated ventral flexions of the caudal pedicles and thereby escape with a “hopping” movement. Abbreviations: an, anus; mg, midgut; ph, pharynx; vlm, ventrolateral longitudinal muscle blocks; yc, Y-cells.

24 

 1 Gastrotricha

The escape response in Dactylopodola and Chordodasiopsis is different to the aforementioned mode. In these taxa, rapidly repeated ventral flexions of the caudal lobes (Fig. 1.14 G) cause a rearward “hopping”. A correlation between such fast movements and the occurrence of cross-striated body musculature in Dactylopodola and Chordodasiopsis is obvious and can be functionally explained because crossstriated muscles are better suited for short but quickly repeating contractions than is obliquely striated musculature. The latter, meanwhile, is hypothesized to be the optimal muscle type for soft-bodied, vermiform animals where hyperextensions are followed by strong contractions of the whole body (Ruppert 1991). A predominant occurrence of obliquely striated longitudinal musculature among Gastrotricha supports this hypothesis (see Tab. 1.3). In addition to the already mentioned defensive behavior of Dasydytidae (see above), taxa like Thaumastoderma, Lepidodasys martini, or Kijanebalola are able to partially retract their anterior body end as a defensive response (Remane 1936). We have observed a comparable behavior in some specimens of Turbanella cf. subterranea that where consciously disturbed. First, the animals partially retracted the head and/or the caudal lobes bearing the posterior adhesive tubes. Shortly after, an extreme contraction of the whole body may follow (see Fig. 1.14 A–E). The main role of the visceral circular and helicoidal muscles is regarded to antagonize dilations of the pharynx and the midgut (Ruppert 1982, 1991, Hochberg & Litvaitis 2001c). Further roles of these muscles in the intestinal region could be the allocation of propulsive force to shift big food items (e.g., diatoms) through the midgut and to stiffen the whole gut tube (Hochberg & Litvaitis 2001c). Egg deposition in Macrodasyida often involves strong contractions of the trunk of spawning animals (Teuchert 1968, see chapter Reproductive Biology). These contractions are obviously brought about by longitudinal muscles. In freshwater Chaetonotidae, ripe eggs leave the trunk on the ventral side, possibly through a still unknown pore, pushed by muscular contractions (Hummon & Hummon 1983a). One or two branches of musculi dorsales, the dorsodermal muscles, form an arch above the developing egg in paucitubulatinan gastrotrichs. These muscles are hypothesized to stabilize the position of the egg and during egg deposition they may lead it ventrally and out of the body (Hochberg & Litvaitis 2003a). The functional role of yet other muscle components like, e.g., the crossover muscle(s) in the posterior trunk of many species of the Macrodasyida is still not satisfactorily understood (Hochberg & Litvaitis 2001c).

1.2.4 Nervous system The nervous system of the stem species of Gastrotricha includes a dorsal to dorsolateral, bilateral symmetric brain

in the anterior part of the body (head region) and a pair of lateroventral longitudinal cords (neurite bundles) as main components (Rothe et  al. 2011a, Figs. 1.15 and 1.16 A–D). The brain has been reconstructed in different ways. Its basic structure has in principle already been described by Ludwig (1875) and Bütschli (1876), in more detail by Zelinka (1889) and Remane (1936). According to these authors, the brain consists of a bridge of neurons (the dorsal commissure) dorsal of the pharynx and cells (=somata) on both sides of this “bridge”. Ultrastructural investigations by Teuchert (1977a) and Wiedermann (1995) reconstructed the brain as circumpharyngeal, but with a stronger dorsal part. Both investigations found a different distribution of somata, which cover a broader area, either more or less homogeneous over the brain (Wiedermann 1995) or separated in an anterior and a posterior part (Teuchert 1977a). Immunohistochemical investigations on Dactylopodola baltica, Macrodasys caudatus and Dolichodasys elongatus (Hochberg & Litvaitis 2003b), on Neodasys cirritus, Xenodasys riedli, and Turbanella cf. hyalina (Hochberg 2007), on three Turbanella species (Rothe & Schmidt-Rhaesa 2008), two Dactylopodola species (Rothe & SchmidtRhaesa 2009), Oregodasys cirratus (Rothe & SchmidtRhaesa 2010), two Xenotrichula species (Rothe et al. 2011b), Lepidodasys worsaae (Hochberg & Atherton 2011), and Neodasys chaetonotoideus (Rothe et  al. 2011a) confirmed the description by Zelinka and Remane. There is a broad commissure composed only of neurites dorsal of the pharynx (Figs. 1.16 A–C and 1.17 A). This commissure is well shown with immunoreactivity (IR) against tubulin. IR against other neuronal components may show only subsets, as is shown exemplary in the IR against serotonin, histamine, and FMRFamides in Dactylopodola species, which all stain only some fibers within the dorsal commissure (Rothe & SchmidtRhaesa 2009). This observation also accounts for the immunohistochemical investigations in the other species. Nuclear staining shows a number of cells in the region lateral of the dorsal commissure (approximately 20 per side in Dactylopodola; Rothe & Schmidt-Rhaesa 2008), but the neural markers used (anti-serotonin, anti-FMRFamides, anti-histamine) all stain only subsets of the nerve cells of the brain (compare Fig. 1.16 C with 1.16 D, see also Fig. 1.17 B). Although there are at maximum few pairs of anti-serotonin IR cells (one to five pairs) in the studied species (e.g. Fig. 1.16 A–C), a count of anti-FMRF amide IR cells in the brain of Nesodasys chaetonotoideus and Xenodasys riedli yielded a mean number of 24 cells per hemisphere of the brain (Hochberg 2007, Rothe et al. 2011a). Therefore, it cannot be said with certainty how many neurons in total constitute the brain of Gastrotricha. All somata stained with any neuronal marker are positioned lateral of the dorsal commissure, a result that gets support by ultrastructural data (e.g., Rothe et al. 2011a).

1.2 Morphology 

The lateral ends of the dorsal commissure are the origin of the longitudinal nerve cords or neurite bundles (Fig. 1.16 A–D). Ultrastructural cross sections show one pair of longitudinal nerve cords in a lateroventral position in the animals (Teuchert 1977a, Ruppert 1991, Rothe et  al. 2011a; Figs. 1.13 C, 1.17 C–D). In species such as Turbanella cornuta, a basiepithelial and intraepidermal position of the nerve cords

sci avc dc

an scb

br

ph

vc

vln

sci

in

vln

pc

cg

Fig. 1.15: Nervous system (schematic) of the last common ancestor of Gastrotricha. Green color indicates general nervous patterns, blue color indicates serotonin expressing components. The orange ovals are serially arranged FMRF-immunoreactive cells alongside the ventrolateral neurite bundles that were possibly also present in the stem species of Gastrotricha. Abbreviations: an, anterior longitudinal neurite bundles; avc, anterior ventral commissure of the brain; br, brain; cg, caudal ganglion/anal ganglion; dc, dorsal commissure of the brain; in, intestine; pc, posterior commissure; ph, pharynx; scb serotonin-expressing nerve cells of the brain; sci, sensory cilia; vc, ventral commissure of the brain; vln, ventrolateral longitudinal neurite bundles. Drawing according to the reconstructed character pattern of Rothe et al. (2011a).

 25

is reported (e.g. Ruppert 1991). However, in putative basal taxa such as Neodasys, the nervous system is most likely subepidermal (Fig. 1.17 B–C). Immunohistochemical investigations show some additional aspects. The distance between the two longitudinal nerve cords is quite wide in most species, but in Xenotrichula species, the cords are closer together (Rothe et al. 2011b). In the broad species Oregodasys cirratus, the distance between the longitudinal cords is wide, but not as wide as possible. The cords run in a distance of about 50 µm from the lateral margin and fine neurites run from the cords into the lateral regions of the animal (Rothe & Schmidt-Rhaesa 2009). In this species, anti-serotonin IR reveals a second pair of longitudinal neurites or neurite bundles median of the longitudinal cords (Rothe & Schmidt-Rhaesa 2009), this structure is unknown from other species so far. Sometimes it appears that two longitudinal neurite bundles per body side are present in close proximity (see, e.g., for Dactylopodola; Rothe & SchmidtRhaesa 2009); in this case, it is likely that two fibers within the entire broader nerve cord were stained. In Turbanella species, the longitudinal cord first runs close to the pharynx and then turns laterally to proceed in a more lateral position. In the posterior end, both longitudinal cords merge in a loop. In Xenodasys riedli, there is a strong posterior commissure, from which a pair of additional neurites runs further posterior (Hochberg 2007). In the two Xenotrichula species and in Neodasys chaetonotoideus, a pair of anti-serotonin IR somata is present in the posterior part of the longitudinal nerve cords (Rothe et al. 2011a, b; Fig. 1.16 A, B). The presence of such an “anal ganglion” (sensu Remane 1936) is possibly also a character of the last common ancestor of all extant Gastrotricha (see Rothe et al. 2011a, Fig. 1.15). In some species, putative nerve cell somata where observed along the ventrolateral neurite bundles (Hochberg 2007, Rothe et al. 2011a). Very fine ventral commissures between the longitudinal nerve cords are present, but these are detected only by a particular IR. In Turbanella species, one serotonin-IR ventral commissure is present close to the level of the dorsal commissure (Rothe & Schmidt-Rhaesa 2008). In Dactylopodola species, four commissures are present in different positions along the body, all four are stained by anti-tubulin, two of them by anti-RF amide, and none by anti-serotonin or antihistamine (Rothe & Schmidt-Rhaesa 2009). Two ventral commissures are present in Neodasys chaetonotoideus, one is stained by anti-RF amide, and the other by anti-serotonin (Rothe et al. 2011b; Fig. 1.16 A). Several neurons are observed to run from the brain region into the anterior end, these neurons likely innervate the sensory structures in the head region. The anterior and posterior sensory organs can usually be seen in immunohistochemical investigations quite well. Some neurites also innervate the pharynx and its ciliated sensory cells (see chapters Intestinal System and Sensory Structures).

26 

 1 Gastrotricha

A

vc sbr

B

C

D dc

d

anb

br sbr

dc

pnb

dc lnb

v lnb

lnb lnb

ag

ag pc

pc

pc 100 µm

25 µm

50 µm

d

d

v

v pc 50 µm

Fig. 1.16: Nervous system of Gastrotricha. Maximum projections of confocal image stacks. Serotonin expressing cells (A–C) or neurons with immunoreactivity against anti-FMRF-amides (D) were stained with fluorescence-labeled antibodies. (A) Neodasys chaetonotoideus (Multitubulatina). Note that the specimen is slightly twisted in the rear trunk. (B) Chaetonotus maximus (Paucitubulatina). Note that both N. chaetonotoideus and C. maximus have a pair of neurons close to the posterior end (“anal ganglion”). (C and D) Dactylopodola typhle (Macrodasyida). Note that more neurons of the brain and an unpaired ventral neurite bundle of the pharynx are stained with anti-FMRFamide (D). Confocal datasets of A, C, and D: Birgen H. Rothe, Halle (Westf.). Abbreviations: ag, anal ganglion; anb, anterior neurite bundles; br, brain; d, dorsal; dc, dorsal commissure of the brain; lnb, longitudinal neurite bundle; pc, posterior commissure; pnb, pharyngeal neurite bundle; sbr, serotonin-expressing cells of the brain; v, ventral; vc, ventral commissure of the brain.

1.2.5 Sensory structures Long before the highly resolving electron microscopy was available for the study of microscopic animals, numerous different types of presumptive sensory organs were already known for the Gastrotricha. Among these are external sensory cilia (“tactile hairs” or “tactile bristles”), either positioned individually or associated with adhesive tubes (Fig. 1.18 A, G). Especially at the anterior end, those sensory cilia are mostly arranged in groups or tufts (Remane 1936). Further sensory structures are paired, tentacle-like palps on the anterior head region in several species (Fig. 1.18 A, B, D, F), sometimes combined with further structures like clusters of composed cilia (“cirri”) such as in species of the Xenotrichulidae. Palps in Gastrotricha may be rather short and cone-shaped protrusions on both sides of the head like in Turbanella cornuta (and other species of that genus) but

may also feature long and thin formations as in Dinodasys mirabilis (Remane 1936). Species of the taxon Thaumastoderma always have a combination of one pair of rod-shaped lateral tentacles plus one pair of spittle-like tentacles, a possible autapomorphy of that group (Kieneke 2010; Fig. 1.18 F). A further presumptive sensory device that is regularly described for different taxa of the Macrodasyida is the so-called “piston pit” or “pestle organ”. Such a structure is known from, e.g., species of Macrodasys, Urodasys, Paraturbanella, and certain members of the Thaumastodermatidae (Remane 1936). Each piston pit is a lateral, roundish depression in the head region that is provided with numerous sensory (?) cilia surrounding a central, knob-like elevation (see Remane 1936). Up to now, a full reconstruction of such an organ based on ultrastructural data has not been carried out. However, some scattered data and a presumptive chemoreceptive function for the pestle organs of Macrodasys

1.2 Morphology 

A

 27

B *

dc ph

br

br ph

br

* *

10 µm

C

5 µm

D mg lnb

lm

lnb

lm

ed

ed

cu 4 µm

1 µm

Fig. 1.17: Ultrastructure of the nervous system of Gastrotricha (TEM cross sections). (A–C) Neodasys chaetonotoideus (Multitubulatina). (A) Cross section of the brain (cerebral ganglion) at the level of the dorsal commissure. (B) Left hemisphere of an anterior portion of the brain with 3 anterior neurite bundles (asterisks). (C) Ventrolateral longitudinal neurite bundle in a subepithelial position. (D) Xenotrichula carolinensis (Paucitubulatina). Close-up of the longitudinal neurite bundle possibly close to a synapse with a ciliated epidermis cell. Note the high number of neurovesicles. Abbreviations: br, brain (cerebral ganglion); cu, cuticle; dc, dorsal commissure; ed, epidermis; lm, longitudinal musculature; lnb, longitudinal neurite bundle; mg, midgut; ph, pharynx.

(and the corresponding lateral organs of Neodasys) are reported by Gagné (1980a). According to Teuchert (1976a), the piston pit/pestle organ could be homologue to the posterior head sensory organ (described below) that is now known from different species. In addition to the aforementioned sensory structures, some species across diverse taxa possess pigment-bearing photoreceptors, for example, Dactylopodola baltica (Fig. 1.18 C) or Thaumastoderma heideri (Fig. 1.18 F) with their beautiful reddish

eye spots. In several species, the species name hints to the wealth of pigmented photoreceptors such as in Oregodasys ocellatus (Clausen 1965), Turbanella ocellata (Hummon 1974), or Macrodasys ommatus (Todaro & Leasi 2013; Fig. 1.18 E), just to mention some. Among the predominantly freshwater-inhabiting family Chaetonotidae (Paucitubulatina), there are few species that bear so-called “pseudocelli” such as Heterolepidoderma ocellatum. It is still questionable if these globular, light-refracting

28 

 1 Gastrotricha

structures really represent sensory organs or are parts of them (Schwank 1990). Species of the derived planktonic taxon Neogossea exhibit a pair of pigmented, kidneyshaped structures on the dorsal side of the anterior portion of the pharynx (e.g., Kieneke & Riemann 2007). These structures (originally called “Rückendrüsen”) are believed to be secretory structures (e.g., Schwank 1990). However, it is possible that these highly specialized animals benefit from

A

B

a light-perception sense and that both structures therefore represent pigmented photoreceptors. Only by means of ultrastructural investigations it will be possible to clarify the functional role of Neogossea’s “Rückendrüsen”. Investigations with the TEM have yielded great insights into microanatomy and putative function of, and possible homology between different sensory organs of the Gastrotricha. Until recently, full morphological

D

F

hy

50 µm

50 µm 40 µm

E

G

C 50 µm

50 µm

TbA

H

50 µm

50 µm

I

1 µm

cu

cu

ed

cp cp

rc

anb 2 µm

Fig. 1.18: Sensory structures of Gastrotricha. (A) Aspidiophorus tentaculatus (Paucitubulatina) with club-shaped tentacles and tactile cilia on the anterior end, ventral view. (B) Dactylopodola cornuta (Macrodasyida) with cone-shaped tentacles, ventral view. (C) Dactylopodola baltica with red-pigmented eyes. (D) Xenodasys riedli (Macrodasyida) with long cephalic tentacles, ventral view. (E) Macrodasys cf. ommatus (Macrodasyida) with eyes that obviously comprise pigmented cup cells. (F) Thaumastoderma heideri (Macrodasyida) with several paired sensory structures at its anterior end (pigmented eyes, spatulate tentacles, rod-shaped tentacles and sensory cilia). (G) Macrodasys caudatus with numerous sensory cilia at the anterior end, ventral view. (A, D–G) DIC micrographs. (B and C) BF micrographs. (H and I) Ultrastructure (TEM cross sections) of the anterior head sensory organ of Gastrotricha: (H) Dactylopodola baltica and (I) Neodasys chaetonotoideus (Multitubulatina). Abbreviations: anb, anterior neurite bundle; cp, microvilli-like ciliary processes; cu, cuticle; ed, epidermis; hy, hypostomion; rc, receptor cell; TbA, anterior adhesive tubes.

1.2 Morphology 

1980b). In different studied gastrotrich species such pores are lacking in the putative chemoreceptors (e.g., Teuchert 1976a, Gagné 1980b). However, a thinned body cuticle in the area of the chemoreceptor is assumed to be permeable to different substances (Gagné 1980b, see Fig. 1.21). All sensory organs of the Gastrotricha known so far consist of one single, few, or several ciliated receptor cells. The only hint of non-ciliated receptor cells in Gastrotricha is the unpublished observation of rhabdomeric photoreceptors in some gastrotrichs (see Gagné 1980b). Especially the anterior and posterior head sensory organs of many gastrotrich species (see below) additionally comprise a varying number and different types of supportive cells (see Tab. 1.4). Generally, the sensory cells are monociliar, bipolar, primary receptor cells with a nucleus-containing

cil mv

A

cut

cp de sj

ec

er

nu rc

2 µm

gly ec

mi ax

hd

B sj

cp

cil

ec

1 µm

reconstructions based on TEM investigations have been carried out for the cephalic tentacles and regular sensory processes of Chordodasiopsis antennatus (Rieger et  al. 1974), the anterior and posterior head sensory organs of Turbanella cornuta (Teuchert 1976a) and Dactylopodola baltica (Hochberg & Litvaitis 2003b, Liesenjohann et  al. 2006), diverse types of sensory cilia (“tactile hairs”) of T. cornuta (Teuchert 1976a), the sensory palps of Tetranchyroderma papii (Gagné 1980b), and the unique gravireceptor organs of Pleurodasys helgolandicus (Marotta et al. 2008). Additional ultrastructural data related with certain sensory organs exist for Cephalodasys maximus (Wiedermann 1995), Cephalodasys sp., Crasiella cf. diplura, and Xenotrichula carolinensis (Ruppert 1991), and for Lepidodermella squamata (Hochberg 2001). Predominantly based on morphological (ultrastructural) evidence, three functional types of receptor organs have been identified in Gastrotricha: mechanoreceptors (including the gravireceptors of P. helgolandicus), photoreceptors, and chemoreceptors (Teuchert 1976a, Ruppert 1991, Marotta et al. 2008). Although it is not possible to deduce the function of a sensory organ just by morphological and ultrastructural evidence (Laverack 1974, reviewed in Gagné 1980b), comparative morphology of organs of various taxa with a known function (e.g., proven by physiological experiments) facilitates the development of functional hypotheses for organs of taxa where, for example, physiological evidence is difficult to obtain as in Gastrotricha (see Gagné 1980b and references therein). An argument for a mechanoreceptive function is the common construction of the receptor cell among diverse taxa, which includes an external cilium that is surrounded by a collar of circumciliary microvilli (Teuchert 1976a, see Fig. 1.19 A, B). Photoreceptors frequently possess pigments that shield parts of the sensory cells or filter a certain spectrum of light. Different species of the Gastrotricha possess such “colored eyes” (see above). Another characteristic of photoreceptors is the presence of a significant membrane proliferation to house the visual pigments (Gagné 1980b), which frequently consists of stacked and highly ordered cell processes (Eakin 1972, see Fig. 1.20). Such structures are indicative for “non-pigmented eyes”. Additionally, light-refracting structures may be situated in close proximity to putative photoreceptors (Teuchert 1976a). A changing light sensitivity (negative phototaxis) during the reproductive cycle of Turbanella cornuta (Teuchert 1975a) is regarded as indirect evidence for a photoreceptive function for the anterior of both head sensory organ pairs in that species (Teuchert 1976a). To perceive chemical signals, the corresponding sensory devices must be able to communicate with the environment. Hence, one suspects certain openings/pores in such organs that enable dissolved substances to reach the actual sensory cells (Gagné

 29

ime mv

Fig. 1.19: Mechanoreceptive ciliated cell (schematic) of the epidermis of Turbanella cornuta. (A) Sagittal section. Note the single crossstriated ciliary rootlet and the numerous vesicles at the apical side of the cell. (B) Cross section at the level of the ciliary pit where the basal body of the cilium passes into the axoneme (level indicated by 2 black triangles in A). Note the 10 ridges of the ciliary pit (mv) that transform into the 10 circumciliary microvilli more distally. Abbreviations: ax, axon; cil, cilium; cp, ciliary pit; cut, cuticle; de, belt desmosome; ec, epidermis cell; er, endoplasmic reticulum; gly, glycogen; hd, hemi-desmosome; ime, inter-microvillar ECM; mi, mitochondrion; mv, microvilli; nu, nucleus. (A, Modified from figure 2A of Teuchert 1976a; B, according to TEM micrograph in figure 3A of Teuchert 1976a.)

30 

 1 Gastrotricha sc

mp

nu

cil (b) ol mv cil sj

sec

nu

dy rc

mi

sc nu

2 µm

er

ax

Fig. 1.20: Anterior head sensory organ (schematic) of Turbanella cornuta, sagittal section. Note the single cross-striated ciliary rootlet accompanied by numerous microtubules. Abbreviations: ax, axon; cil, modified cilium (dendritic section of receptor cell); cil, (b) modified cilium of a second receptor cell; dy, dyctiosome; er, endoplasmic reticulum; mi, mitochondria; mp, microvilli-like processes of the modified cilium (containing a central microtubule each); mv, microvilli; nu, nucleus; ol, organ lumen; rc, receptor cell; sc, sheath cell; sec, secretory cell with large secretory occlusions; sj, septate junction. (Modified from figure 7 of Teuchert 1976a.)

mv cu

ol

mv

ed

cil

sc

mt sc

nu rc

sj sc

sg nu mi

sc

sg

ax rc 15 µm

Fig. 1.21: Posterior head sensory organ (schematic) of Turbanella cornuta, horizontal section (anterior to the left). Note the single crossstriated ciliary rootlet per sensory cell and the secretion granules of the sheath cells with differing content. Endoplasmic reticulum and Golgi cisterns omitted for clarity. Abbreviations: ax, axon; cil, modified and irregularly branched cilium (dendritic section of receptor cell); cu, cuticle; ed, epidermis; mi, mitochondria; mt, microtubules; mv, microvilli; nu, nucleus; ol, organ lumen; rc, receptor cell; sc, sheath cell; sg, secretion granules; sj, septate junctions. (Modified from figure 4 of Teuchert 1976a.)

2  ≥ 1  ≥ 1 N.A. N.A. 1

1

1

1–2  ≥ 2 (?)  ≥ 1 N.A. N.A. 1

1

1

N.A.

 ≥ 4

Several

N.A.

 ≥ 4

Several

N.A. Yes

Yes

Yes

1 (cup-shaped, Yes with pigments)  ≥ 1 Yes

2–3 (as support Yesb cells)b N.A. N.A.

N.A.  ≥ 2

N.A.

 ≥ 2 (secretory)

Organ has diract contact with body cuticle

N.A.

Absent Present

Present

Present Present Present N.A. N.A. Present

Basal inflation of receptor cell(s)

N.A.

No

N.A.

Yesb

N.A. Yes

N.A.

Yes

Microvilli of receptor cells and/or sheath cells penetrate endocuticle

1 (secretory cell)  ≥ 1 (pigment cells) N.A. N.A. N.A. 1 (pigment cell)

Additional cells

Yes

N.A.

N.A.

Nob

N.A. N.A.

N.A.

Yes

Regular

N.A. Irregular

Irregular (?)

Irregular

Irregular

Irregular

N.A.

­–

Present Present Present N.A. N.A. Presenta

N.A.

N.A.

N.A.

–­

N.A. N.A.

N.A.

Present

Microtubules inside the processes

N.A.

N.A.

N.A.

Presentb

N.A. N.A.

N.A.

Present

Ciliary rootlet present

Present

Regular/parallel Present

Regular N.A. Regular/parallel N.A. N.A. Regular/parallel

Cilium branches Arrangement into numerous of (ciliar) microvillus-like processes processes

Yes

Yes

Yes N.A. Yes N.A. N.A. Yes

Photoreceptor Chemoreceptor

Chemoreceptor Chemoreceptor N.A. Chemoreceptor Chemoreceptor N.A.

General light sensivity Photoreceptor

Photoreceptor Photoreceptor Photoreceptor N.A. N.A. Photoreceptor

Presumed function

Liesenjohann et al. (2006); for presumed function: Ruppert (1991)

Hochberg & litvaitis (2003b) Liesenjohann et al. (2006)

Gangé (1980)

Ruppert (1991) Ruppert (1991)

Wiedermann (1995)

Teuchert (1976)

Presumed Reference function

N.A.

Absent

Present Present N.A. N.A. N.A. N.A.

Cilium branches into Arrangement of Central microtubule Ciliary rootlet numerous microvillus- ciliar processes inside the processes like processes

Modified and amended from Liesenjohann et al. (2006). A questionmark (?) indicates an uncertain character state, a dash (-) indicates an inapplicable character state, n.a. no data available. a) There is a single microfilament inside each microvillus-like process of the anterior head sensory organ of D. baltica according to Hochberg & Litvaitis (2003b). b) Gagné (1980) considers the sensory palps of T. papii to be homolog to the posterior hso of Turbanella cornuta

22–23b

N.A.  ≥ 1

N.A.  ≥ 1

22–23b

N.A.

N.A.

Cephalodasys maximus Cephalodasys sp. Crasiella cf. diplura

12–14

12–14

Number of Number of Number receptor receptor of sheath cell(s) cilia cell(s)

Turbanella cornuta

Tetranchyroderma papii Dactylopodola baltica (US Atlantic Coast) Dactylopodola baltica (North Sea) Stem species of Gastrotricha

1

1

 ≥ 2  ≥ 2 (?)  ≥ 1 N.A. N.A. 1

Posterior head sensory organ

Turbanella cornuta Cephalodasys maximus Cephalodasys sp. Crasiella cf. diplura Tetranchyroderma papii Dactylopodola baltica (US Atlantic Coast) Dactylopodola baltica (North Sea) Stem species of Gastrotricha

Number of Number of Number receptor receptor of sheath cell(s) cilia cell(s)

Anterior head sensory organ

Tab. 1.4: Ultrastructural characters related with the anterior and posterior head sensory organs in different species of the Macrodasyida.

1.2 Morphology 

 31

32 

 1 Gastrotricha

cell body (perikaryon), a proximal axon, and a distal ciliary (dendritic) segment (Teuchert 1976a, Ruppert 1991). Mechanoreceptors of Gastrotricha are sensory cells with mostly elongated and rather stiff external cilia (the cilium is, as all external cilia in Gastrotricha, enwrapped by the lamellar epicuticle). The central cilium arises from a ciliary pit and is surrounded by a collar of 10 circumciliary microvilli (“stereocilia”) arranged in a regular manner (Fig. 1.19 A, B). Microvilli arise from 10 longitudinal ridges that line the ciliary pit. The sensory cilium possesses a basal body and a well-developed, straight ciliary rootlet (Teuchert 1976a, Fig. 1.19 A). In contrast to this set of characters, ciliated locomotory epidermis cells of Gastrotricha almost always have a rostral plus a caudal ciliary rootlet, obliquely aligned within the cell and a number of eight circumciliary microvilli (Rieger 1976, Hochberg 2001, see also chapter Integument). The number of 10 circumciliary microvilli in mechanoreceptive sensory cells (Fig. 1.19 B) is so far only known from Gastrotricha and could therefore represent an autapomorphy of this phylum (Hochberg 2001). The microvilli are interconnected by a specialized ECM that displays a high degree of order (see figure 3A of Teuchert 1976a, Fig. 1.19 B). This “intermicrovillar” ECM resembles the one that can be observed between the eight circumciliary microvilli of the terminal cells of protonephridia of species like Turbanella cornuta (Teuchert 1973), Chaetonotus maximus (Kieneke et al. 2008b), or Polymerurus nodicaudus (Kieneke & Hochberg 2012). Possibly, this similarity hints to a common ectodermal origin of protonephridia and mechanoreceptors. Basally, mechanoreceptive sensory cells form an axon like all sensory cell types in Gastrotricha (Fig. 1.19 A). In T. cornuta, direct connections of these axons to the neuropil of the brain or with the ventrolateral longitudinal neurite bundles are reported (Teuchert 1976a). Numerous mechanoreceptive sensory cells (“sensory hairs”) are individually positioned along the body and are embedded between regular epidermal cells (Teuchert 1976a, Fig. 1.19 A, B). In species such as T. cornuta or Neodasys sp., a single mechanoreceptive sensory cell is furthermore associated with each adhesive tube/adhesive organ (Teuchert 1976a, Tyler et  al. 1980). In this case, the sensory cell body may be deeply submerged below the level of the epidermis as in T. cornuta (see Teuchert 1976a). The number of individual mechanoreceptive sensory cells on the trunk was dramatically reduced in the stem lineage of the Paucitubulatina: species of the basal marine taxon Xenotrichulidae possess only few pairs of elongated sensory cilia (up to seven) along their body and at the base of the caudal furca (see, e.g., Ruppert 1979). Members of Chaetonotidae, a probably not monophyletic group (see chapter Phylogeny) at maximum possess two pairs of stiff sensory cilia on their back (see, e.g., Schwank 1990). These so-called setolae (“sensory bristles”)

frequently originate from specialized cuticular scales that have a certain diagnostic value. Embedded in the myoepithelial wall of the pharynx of many species of Gastrotricha, there are three columns of individually positioned ciliated receptor cells that strongly resemble the mechanoreceptive sensory cells of the body surface (Ruppert 1982). The cilium penetrates the enducuticle but is enclosed by the epicuticle of the pharynx. It is surrounded by a collar of microvilli (10?) that arise from longitudinal ridges of the ciliary pit. A difference between external mechanoreceptors and the pharyngeal receptor cells is the presence of at least two ciliary rootlets in the latter (Ruppert 1982, Rothe et al. 2011a). At the anterior end of the pharynx in species such as Neodasys sp. and Halichaetonotus sp., there are three receptors consisting of two such ciliated sensory cells each (Ruppert 1982). Especially in the head region, there may be mechanoreceptors that consist of clusters of numerous mechanoreceptive sensory cells as, for example, close to the mouth opening of Turbanella cornuta. In this species, the receptors form small bulges that emerge above the level of the epidermis (Teuchert 1976a). Receptor cells are connected to each other via cellular junctions (apical belt desmosomes plus septate junctions, see figures 2C and 3B of Teuchert 1976a). A comparable cluster of three to four ciliated receptor cells is present anterior to the sensory palps of Tetranchyroderma papii (Gagné 1980b). At the posterior trunk end of species of the limnic-planktonic taxon Stylochaeta there is a pair of short, blunt processes, the so-called styli. Each stylus bears two to three cilia supposed to have sensory function (Schwank 1990). Comparable bulges with a single cilium, however, also occur along the body of species such as Dinodasys mirabilis (Remane 1936) or Oregodasys cirratus (Rothe & Schmidt-Rhaesa 2010). Further multicellular mechanoreceptors, the so-called sensory cirri, are present in the head region of species of the taxon Xenotrichulidae (e.g., Ruppert 1979, 1991, Rothe et al. 2011b). They are composed of several receptor cells, the externally projecting cilia of which are enclosed by a common lining of the epicuticle (see Ruppert 1991). Generally, an at least partly sensory function (besides the still locomotive duty) is assumed for most cilia on the head of Gastrotricha, frequently arranged as tufts or batches (Remane 1936). Hochberg (2001) discovered a special form of such sensory cilia on the head region of the freshwater paucitubulatinan species Lepidodermella squamata that display a wart-like surface on the lower ciliary shaft. Multicellular sensory organs with a presumed mechanoreceptive function but a considerably different construction are described for Chordodasiopsis antennatus. This species possesses peculiar, antenna-like, and ostensibly articulated processes on the head region, the cephalic tentacles, and along the whole trunk including the anterior end, viz. the regular sensory processes (Rieger et al. 1974).

1.2 Morphology 

Both types of sensory processes consist of numerous strongly elongated ciliated receptor cells that distally end with a rather short cilium surrounded by short microvilli. In the cephalic tentacles, at least two such external sensory cilia are positioned at each “annulus”, whereas there is a single cilium at the distal end of each “segment” of the regular sensory processes. At the distal tip of the latter, there are usually two cilia and three at the tip of each cephalic tentacle (Rieger et al. 1974). Proximal to the basal body, extremely elongated ciliary rootlets, accompanied by microtubules, extend deeply into each sensory cell. Owing to the successive ending of the receptor cells at the annuli of the sensory processes, each cell (and its ciliary rootlet) has a different total length. The main cell bodies with the nuclei are aligned within the epidermis below each process, in the case of the cephalic tentacles these cells are in close proximity to the brain of C. antennatus. In the regular sensory processes, the sensory cells proximally form long and thin appendages, probably axons that lead to the longitudinal neurite bundles. A conspicuous difference between both types of receptors in C. antennatus is the presence of several nerve fibers in the cephalic tentacles that originate in the brain (Rieger et al. 1974). Owing to the exclusive occurrence of unmodified ciliated receptor cells in the sensory processes and cephalic tentacles of C. antennatus, Gagné (1980b) rejects homology of these organs with the sensory palps of Tetranchyroderma papii and the posterior head sensory organs of Turbanella cornuta (see below). A quite unusual and among Gastrotricha unique sensory organ is the pair of supposed gravireceptor organs of Pleurodasys helgolandicus that are positioned on the dorsal surface in the anterior part of the animal (Marotta et al. 2008). Each organ consists of a single ciliated receptor cell with a similar cytomorphology as the common mechanoreceptive cells of Gastrotricha, i.e., they possess a long cilium that is basally surrounded by a collar of 10 short microvilli (see above). Hence, Marotta et al. (2008) suggest an evolutionary origin of the gravireceptor organs from simple mechanoreceptor cells. The peculiarity of the gravireceptor organs is the presence of a drumstick-shaped, external protuberance of the epicuticle into which the cilium of the receptor cell projects. The stalk and bulbous tip of the organ are formed by up to 100 densely piled layers of the epicuticle. Inside the globular part, there is a sphere that consists of numerous, electron-dense vesicles that obviously originate from the apical membrane of the receptor cilium. Because the drumstick-shaped external part of the organ has a jointed connection to the body cuticle and is freely movable and because the sphere is reminiscent to the otoliths of statocysts in other taxa, Marotta et al. (2008) conclude that the organs in P. helgolandicus must display gravireceptors. These organs provide P. helgolandicus with

 33

information about the animal’s position within the Earth’s gravitational field, a sense that might be quite important for an organism that lives in the three-dimensional space between sand grains. However, it is so far not understood why P. helgolandicus is the only known gastrotrich species with such sensory devices (Marotta et al. 2008). Based on detailed ultrastructural investigations of two different pairs of head sensory organs (or one of both, respectively) in the species Turbanella cornuta (Teuchert 1976a), Dactylopodola baltica (Hochberg & Litvaitis 2003b, Liesenjohann et al. 2006), Tetranchyroderma papii (Gagné 1980b), and some more fragmentary data of further species (Wiedermann 1995, Ruppert 1991), Liesenjohann et al. (2006) postulate the presence of two such pairs of sensory structures in the stem species of Gastrotricha, i.e., the anterior head sensory organ and the posterior head sensory organ (see Tab. 1.4, Figs. 1.20 and 1.21). As a result of their phylogenetic evaluation, the last common ancestor of Gastrotricha possessed one pair of anterior head sensory organs in the anterolateral region of the head (Liesenjohann et al. 2006). The anterior head sensory organ has no direct contact to the body cuticle. It is positioned below the epidermis that might, however, be thinned in the area of the receptor (e.g., Teuchert 1976a, Liesenjohann et  al. 2006). Each organ of the stem species is composed of one receptor cell with a single distal cilium that branches into numerous, regularly arranged, microvilli-like processes (Fig. 1.18 H, I). There is a single microtubule inside each ciliary process. Basally (proximal to the modified cilium), the receptor cell is inflated and is further proximally connected to the brain by a common nerve strand of anterior and posterior head sensory organs. In addition to the receptor cell, there is one sheath cell in the anterior head sensory organ of the stem species of Gastrotricha. This supportive cell enwraps the receptor cell with thin cytoplasmic lobes, hence forming an organ lumen inside which the ciliary processes of the receptor cell are densely piled. Based on the morphological arguments presented earlier, the anterior head sensory organ had a photoreceptive function. Because of ambiguous data, it is not clear if a ciliary rootlet, present in some but absent in other species, belongs to the ancestral construction of the anterior head sensory organ (see Tab. 1.4). The anterior head sensory organs have so far been reported and investigated in Turbanella cornuta (Teuchert 1976a, Fig. 1.20), Dactylopodola baltica from the North Sea (Liesenjohann et al. 2006), Dactylopodola baltica from the western Atlantic coast (Hochberg & Litvaitis 2003b), Cephalodasys maximus (Wiedermann 1995), and Cephalodasys sp. (Ruppert 1991). In addition to the aforementioned character pattern of the last common ancestor, extant species of Gastrotricha may show further supportive cells in their anterior head sensory organs. For instance, there

34 

 1 Gastrotricha

is at least one pigment cell in the anterior head sensory organ of Cephalodasys maximus (Wiedermann 1995) and one in Dactylopodola baltica from the western Atlantic coast (Hochberg & Litvaitis 2003b). Interestingly, not any of those additional supportive cells has been found in D. baltica from the North Sea (Liesenjohann et al. 2006). In Turbanella cornuta a secretory cell, quite comparable to the epidermal glands, is closely associated with the anterior head sensory organ (Teuchert 1976a, Fig. 1.20). Posterior to the anterior head sensory organ, the stem species of Gastrotricha possessed one pair of posterior head sensory organs (Liesenjohann et  al. 2006). Each organ is composed of several ciliated receptor cells probably with a single branching cilium (Fig. 1.21). As opposed to the anterior head sensory organ, the ciliary processes of the posterior organ are irregularly arranged and likely lack a central microtubule (microtubules inside the branching cilium were so far only discovered in Turbanella cornuta, see Teuchert 1976a, Tab. 1.4, and Fig. 1.21). The organ lumen is formed by at least one but probably more sheath cells and the whole organ has a direct contact to the body cuticle (Liesenjohann et  al. 2006). The posterior head sensory organ of the last common ancestor of Gastrotricha much likely had a chemoreceptive function (Ruppert 1991). Due to the paucity of data, it is not clear if ciliary rootlets inside the receptor cells belong to the ancestral construction of the posterior head sensory organ (see Tab. 1.4). Posterior head sensory organs have so far been reported and investigated in Turbanella cornuta (Teuchert 1976a, Fig. 1.21), Dactylopodola baltica from the North Sea (Liesenjohann et al. 2006), Cephalodasys maximus (Wiedermann 1995), and Crasiella cf. diplura (Ruppert 1991). The sensory palps of Tetranchyroderma papii differ considerably in their microanatomy from the posterior head sensory organs of, e.g., T. cornuta and D. baltica. However, they are considered to be homologous organs with a comparable chemoreceptive function (Gagné 1980b). The palps of T. papii consist of numerous (22–23) strongly elongated bipolar receptor cells with a modified, non-branching distal cilium. The cilium possesses an extremely short axoneme but is distally elongated into a dendritic process with an expanded diameter. Proximal to the basal body there is a long ciliary rootlet, and each receptor cell is drawn out into a thin axon that posteriorly leads to the brain of T. papii. Inside each palp, the receptor cells are accompanied by two to three elongated support cells that contain a bundle of densely packed and longitudinally arranged microtubules, bringing about mechanic support to the whole organ (Gagné 1980b). In the posterior head sensory organs of Turbanella cornuta, Tetranchyroderma papii, and Crasiella cf. diplura, the sheath cells/support cells form several short microvilli that penetrate the endocuticle (see Fig. 1.21), a character that is possibly related to the

assumed chemoreceptive function (Teuchert 1976a, Gagné 1980b, Ruppert 1991). Interestingly, such short microvilli have not been detected in the posterior head sensory organ of Dactylopodola baltica from the North Sea. It is supposed that the lack of those microvilli is related with a functional change of this organ in North Sea populations of D. baltica. As there is also a cup-shaped, pigment-bearing sheath cell, Liesenjohann et al. (2006) conclude that the posterior head sensory organ of their D. baltica rather sense directed light shielded by the pigment cell than being a chemoreceptor. The anterior head sensory organ of D. baltica from the North Sea seems to lack any pigment-bearing cell and could therefore represent a more general light sensitive organ.

1.2.6 Intestinal system The intestinal system of gastrotrichs starts with a terminal or slightly subterminal anterior mouth, followed by a buccal cavity, a large myoepithelial pharynx, a midgut, and it ends in a ventral anus in the posterior end (Fig. 1.22 A). The pharynx is constructed as a sucking pharynx and all gastrotrichs appear to suck in small food particles together with some water. Bacteria, diatoms, and probably detritus are the most common food particles (e.g., Todaro & Hummon 2008; Fig. 1.22 D, E, G, H). The mouth opening is in many cases (Paucitubulatina, several macrodasyids) a narrow, round opening (Figs. 1.22 B and 1.23 A, B). In other species, e.g., in Neodasys, it widens a bit to form a slight funnel, and in some species, especially those belonging to Thaumastodermatidae, the mouth opens with a wide funnel that occupies almost the entire frontal end of the animal (Figs. 1.22 C, D and 1.23 E). A narrow mouth opening appears well suited for a targeted picking up of food particles such as bacteria, whereas broad mouth openings appear better suited for an untargeted sweeping of the surrounding. In some species, e.g., in the genera Paraturbanella and Macrodasys, the mouth opening is a longitudinal slit rather than a round opening (Ruppert 1991). Especially among paucitubulatinans, the mouth opening is often surrounded by a strengthened cuticle, the mouth ring, from which further cuticular structures, e.g., spines, can originate and form more or less complex buccal structures often called “buccal cage” or “mouth basket” (see, e.g., Balsamo et  al. 2010a on Diuronotus aspetos as one example; Fig. 1.23 A). In Musellifer species, the anterior end of the head forms a snout-like part, the socalled muzzle, which is densely covered by cilia (e.g., Leasi & Todaro 2010). Some macrodasyids, for example, Prostobuccantia have similar elaborations of the mouth opening (Evans & Hummon 1991). Brought about by

1.2 Morphology 

A

B

mo

C

 35

D mo

pp

*

50 µm

50 µm

ph

E 100 µm

mg

20 µm

F

*

H

mg

20 µm

100 µm

pp

50 µm

*

an

G

ph

50 µm

oo

*

Fig. 1.22: Intestinal system of Gastrotricha. (A) Turbanella hyalina (Macrodasyida), horizontal view. Note the typical partition of the gut tube into mouth, buccal cavity, pharynx, midgut, and anus. (B) A marine Chaetonotus sp. (Paucitubulatina) with a subterminal mouth opening, ventral view. (C) Diplodasys cf. meloriae (Macrodasyida) with the characteristic funnel-shaped mouth opening of the Thaumastodermatidae, ventral view. (D) Tetranchyroderma sp. (Macrodasyida) with a rather big diatome inside the midgut (asterisk), horizontal view. (E) Midtrunk section (lateral view) of a Neodasys chaetonotoideus (Multitubulatina) that have had a menu of different diatomes. (F) Midtrunk section of Xenodasys riedli (Macrodasyida). Note the high density of motile cilia inside the posterior portion of the pharynx and in the anterior part of the midgut (asterisks). (G) Middle portion of the midgut of N. chaetonotoideus with 2 euglenoids inside. The algae were still moving when the micrograph has been taken. (H) Posterior part of the midgut of Xenotrichula carolinensis with a tiny diatome inside (asterisk). (A–C, E and F, H) DIC images. (D and G) BF images. Abbreviations: an, anus; mg, midgut; mo, mouth opening; oo, mature oocyte; ph, pharynx; pp, pharyngeal pore.

36 

 1 Gastrotricha

A

B ce

sci

sci

20 µm

5 µm

C

D

E

sci

an TbA

10 µm

4 µm

contractions of certain pharyngeal muscles, the cuticular elements of the buccal cage can be moved forward and the mouth opening may be extended in this way (e.g., Remane 1936, Schwank 1990). The mouth opening leads into a short buccal tube or buccal cavity. In some macrodasyid species, the pharynx appears to attach directly at the mouth opening, but in some macrodasyid species and in pauctitubulatinan chaetonotids, the buccal cavity forms a short cylindric compartment. Tooth-like structures are present in species of the macrodasyid genus Lepidodasys (Remane 1936, Ruppert 1991) and in several paucitubulatinans (Chaetonotus pawlowskii, some species of Aspidiophorus, Heterolepidoderma, and Arenotus (Schwank 1990). These are movable by musculature and are supposed to serve as scrapers (Schwank 1990, see also figures 82 and 83 in Ruppert 1991). The pharynx is a long cylinder with a triradiate lumen (Fig. 1.24 A–C). The musculature is oriented radially between the lumen and the surrounding ECM. Contracting radial myoepithelial cells rapidly extend the pharyngeal lumen and suck in food particles. Anteroposterior contraction waves over the pharynx lead food into the midgut (Remane 1936, Ruppert 1991). Most comparative information on the gastrotrich pharynx goes back to Ruppert (1982; updated in Ruppert 1991). The triradiate pharyngeal lumen has two different orientations. In species of Chaetonotida

TbA

15 µm

Fig. 1.23: Intestinal system of Gastrotricha. SEM micrographs of external openings of the gut tube. (A) Ventral view of head region of Chaetonotus sp. (Paucitubulatina) with the subterminal mouth opening surrounded by cuticular hook-like structures (mouth basket). (B) Anterior end of Macrodasys caudatus (Macrodasyida) with the terminal mouth opening surrounded by velum-like cuticular structures. (C) Pharyngeal pore of Mesodasys sp. (Macrodasyida). (D) Ventral anus of Tetranchyroderma sp. (Macrodasyida) close to the posterior adhesive tubes. (E) Funnellike mouth opening of Tetranchyroderma sp. Abbreviations: an, anus; ce, cephalion; sci, sensory cilia; TbA, anterior adhesive tubes.

(Paucitubulatina+Neodasys), it has the shape of a Y when cross-sectioned (Fig. 1.23 A, B), with paired dorsolateral branches and an unpaired ventral branch; in Macrodasyida, the orientation is an inverted Y (Fig. 1.24 C). The pharynx is ectodermal and therefore lined by cuticle. The pharyngeal cuticle is similar to the body cuticle and composed of endocuticle and epicuticle. The number of epicuticular layers is often larger than in the body cuticle (Ruppert 1991). The pharyngeal endocuticle often lacks the apical fibrillar layer that is present in the body cuticle and basally a number of spherical structures are observed (Ruppert 1991). The cells composing the pharynx are myoepithelial, nervous, sensory, and gland cells. The apical cells, i.e., those at the end of the luminal branches differ from the remaining (Figs. 1.13 D and 1.24 E), interapical cells in the abundant presence of tonofilaments, which connect the cuticle to the surrounding ECM and are therefore probably responsible for maintaining the triradiate shape of the lumen (Ruppert 1991). Most macrodasyid and larger chaetonotid species have multisarcomeral cells; some macrodasyids and most chaetonotids have monosarcomeric cells (Ruppert 1982, 1991). In general, three pharyngeal neurite bundles are present in Chaetonotida (Fig. 1.24 A) and four in Macrodasyida (Ruppert 1982). Each neurite bundle may be composed of about 15 neurites as in Dactylopodola baltica

1.2 Morphology 

A

 37

B *

*

*

2 µm

C

3 µm

D

* tm gl

10 µm

E F

3 µm lm cm ac

pl 1 µm

(Rothe & Schmidt-Rhaesa 2009). However, there are at maximum five neurites per pharyngeal neurite bundle in Neodasys chaetonotoideus (Rothe et al. 2011a). The position is basiepidermally, i.e., close to the outer ECM of the pharynx. The pharyngeal neurite bundles are associated with sensory cells, which are aligned in longitudinal rows (Ruppert 1982, Rothe & Schmidt-Rhaesa 2009, Rothe et al. 2011a). The sensory cells have a short cilium surrounded by microvilli. The cilium projects into the pharyngeal lumen, but remains covered by cuticle (Ruppert 1982, Rothe et al. 2011a, see also chapter Sensory Structures).

1 µm

Fig. 1.24: Ultrastructure (TEM cross sections) of the intestinal system of Gastrotricha. (A) Pharynx of Neodasys chaetonotoideus (Multitubulatina). Note the 3 pharyngeal neurite bundles (asterisks). (B) Pharynx of Xenotrichula carolinensis (Paucitubulatina). (C) Pharynx of Dactylopodola typhle (Macrodasyida). Note the ventral pharyngeal neurite bundle (asterisk). (D) Midgut of N. chaetonotoideus. (E) Detail of the midgut epithelium of N. chaetonotoideus. Note the dense fringe of microvilli. (F) Detail of the pharynx of N. chaetonotoideus with the multisarcomeric myofilaments and the subpharyngeal musculature. Abbreviations: ac, apical cell; cm, circular muscles; gl, gut lumen; lm, longitudinal muscles; pl, pharyngeal lumen; tm, transverse muscle.

With the exception of Lepidodasys, the pharynx in species of Macrodasyida opens to the external by a pair of pharyngeal pores (Fig. 1.23 C). In many species, these pores are in the posterior part, close to the transition to the midgut, but in some species, the pores are further anterior (e.g., in Macrodasys species). The pharyngeal pores have a sphincter that is derived from circumpharyngeal circular muscles. Although their actual function is not fully understood, it is assumed that excess water from food uptake is released through the pores (Ruppert 1991). Cilia and microvilli (apart from those of the pharyn-

38 

 1 Gastrotricha

geal sensory cells) in the pharynx are usually absent, but in few species, their occurrence is observed. Most conspicuous is the presence of long cilia in the region of the pharyngeal pores in species of Chordodasiopsis and Xenodasys (e.g., Schöpfer-Sterrer 1969, Rieger et al. 1974; Fig. 1.22 F) and Dendrodasys (Wilke 1954). Microvilli are less obvious, only in Lepidodasys and Neodasys do they occur in some abundance, but remain below the cuticle (Ruppet 1991). In Xenodasys, Dactylopodola, and Dolichodasys, however, microvilli penetrate the cuticle (Ruppert 1991). In several chaetonotids the pharynx forms one or more bulbous extensions. Schwank (1990) reports regions free of musculature in chaetonotids, which creates a characteristic banding pattern of the pharynx. The transition to the midgut is simple in macrodasyids, but more complex in chaetonotids. In Neodasys, the posterior part of the pharynx forms some folds before the lumen joins the midgut (Ruppert 1991). In species of Paucitubulatina, the posterior end of the pharynx extends into the anterior region of the midgut in the form of a plug (Ruppert 1991), which may function as a kind of weir or valve (Swank 1990). The midgut is straight, without any diverticles or attaching structures, and does not show any obvious compartmentalization. It is composed of a simple absorptive epithelium (Ruppert 1991). Microvilli are present (Fig. 1.24 D, E), cilia are rarely present (in Xenodasys and probably in Dendrodasys, see Ruppert 1991). Teuchert (1977a) reported putative sensory cells in the midgut of Turbanella cornuta. In some paucitubulatinans, a regionalization of the midgut is present and evident by a different coloration (Schwank 1990). Here the anterior part is called stomach. A comparable regionalization may also be found in some macrodasyidans such as Anandrodasys agadasys (Kieneke et al. 2013a). In Macrodasyida, no hindgut is present; only species of Paucitubulatina possess a short cuticularized hindgut (Ruppert 1991). The anus is a simple pore (Fig. 1.23 D), it is often inconspicuous, and in species of Urodasys, it appears to be lacking (e.g., Wilke 1954, Schöpfer-Sterrer 1974).

1.2.7 Body cavities and connective tissue Gastrotrichs are compact animals with comparably few connective tissue, but this condition has been interpreted in different ways. All tissues and cells attach closely to each other. Between epidermis and intestine, muscle and nerve cells, gonads, excretory cells, and few mesodermal cells called Y-cells are present. ECM surrounds the muscle cells, the Y-cells, the protonephridial and

gonadal tissue (Ruppert 1991). This ECM is often very thin and uniform, different layers cannot be distinguished (Ruppert 1991). Ruppert (1991) indicates tiny lacunar spaces in some places, but it is not clear whether such spaces constitute true lacunes of a primary body cavity or regions within the ECM. In figure 56 in Ruppert (1991), such “intercellular lacunae” have a gray staining, similar to the staining of ECM material and may therefore also be intercellular regions filled with ECM. Comparable widenings of the intercellular space filled with ECM are known from the region of the protonephridial terminal cells in different gastrotrich species (e.g., Kieneke et  al. 2007, 2008c, Kieneke & Hochberg 2012). Therefore, we interpret the body organization of gastrotrichs as acoelomate, i.e., with a primary body cavity in compact organization. Despite the compact organization of the gastrotrich body, they were occasionally interpreted as coelomate animals. This goes back to Remane (1936), who interpreted a muscular layer surrounding internal tissue as an equivalent of a mesodermal epithelium. In a case like gastrotrichs, this means consequently that the body cavity is filled with tissue, e.g., the gonads. This view was extended by Teuchert & Lappe (1980; see also Teuchert 1977b and Reisinger 1978). However, this interpretation has two flaws. First, body cavities are, as the name says, cavities and when a cavity becomes occupied by tissue, it is not a cavity any more. Then, as the muscle cells are completely surrounded by ECM, a cavity, if it was present, would be bordered by ECM and not by an epithelium. According to this, a cavity would be a primary body cavity (“pseudocoelom”) and not a coelom (see, e.g., SchmidtRhaesa 2007). A conspicuous cell type in a mesodermal position are the Y-cells, which have been best investigated in Turbanella (Teuchert 1977a, Travis 1983, Ruppert 1991). The Y-cells are arranged in a longitudinal row lateral of the pharynx and the intestine (Figs. 1.14 E; 1.25). Especially in the posterior part, they are in close relation to the longitudinal musculature. Each Y-cell has a central nucleus, little cytoplasm, and a large vacuole. It has been speculated that the row of Y-cells forms an antagonist to the musculature (Reisinger 1978), this would make sense in the flexing movements observed in Turbanella during aggregation and copulation (Teuchert 1968, 1978). Probably homologous cells occur in some other macrodasyids, such as species of Paraturbanella, Macrodasys, Lepidodasys, Acanthodasys, Oregodasys, Tetranchyroderma, and Chordodasiopsis (Travis 1983). Sometimes they occur in rows, sometimes as single cells. Most of these cells contain myofilaments and are therefore likely derived from muscle

1.2 Morphology 

cu

TbD

ed

TbDL

gl

yc

lm

yc

mg lnb

tes

cm

lc

lc

20 µm

Fig. 1.25: Position of the Y-cells of Turbanella cornuta, schematic cross section at the body region of the paired testes. There is no body cavity apart from the gut lumen and the lumina of the testes. Abbreviations: cm, circular muscles; cu, cuticle; ed, epidermis gl gut lumen; lc, locomotory cilia; lm, longitudinal muscles; lnb, longitudinal neurite bundles; mg, midgut; TbD, dorsal adhesive tubes; TbDL, dorsolateral adhesive tubes; tes, testis; yc, Y-cells. (Modified from figure 2E of Teuchert 1977.)

cells (Travis 1983, Ruppert 1991). Apart from the assumed antagonistic/skeletal role, their function is unknown. Neodasys sp. has large hemoglobin-containing cells in a comparable position (Kraus et al. 1981, Ruppert & Travis 1983), these are probably also homologous to the Y-cells, but their storage function for hemoglobin appears to be restricted to Neodasys (see chapter Physiology). However, reddish-colored Y-cells may also occur in further species such as in the recently described Oregodasys caymanensis (Hochberg et  al. 2014). Another strange organ that probably derived from muscle cells is the chordoid organ in species of Xenodasys and in Chordodasiopsis antennatus. It consists of anteroposteriorly piled disc-shaped cells posterior to the anus. Each cell contains myofibrils that transversally span through the cytoplasm (Rieger et  al. 1974). Although this organ in the mentioned gastrotrich species has a comparable ultrastructure like the notochord of the cephalochordate (Branchiostoma), it is not known if it likewise has a skeletal function.

1.2.8 Excretory system Protonephridia have been known for a long time to be present in freshwater gastrotrichs (e.g., Zelinka 1889). Initially, the whole group of Macrodasyida was supposed to lack protonephridia throughout (Remane 1936). Later on, serially arranged protonephridia were detected in several Macrodasyida species (Fig. 1.26) when using phase contrast optics (Wilke 1954, Teuchert 1967). The first electron microscopic survey was done on the protonephridial terminal organ of Chaetonotus sp. (Brandenburg 1962). This study supported the general

 39

ultrastructural composition of protonephridial terminal structures in bilaterian animals: they consist of a cytoplasmic hollow cylinder that is broken by pores or clefts, which are spanned by a filtering diaphragm, a system of motile cilia beating inside a protonepridial lumen (hence generating a negative pressure inside the proximal lumen), and microvilli surrounding the cilia. Subsequent studies on the protonephridial system of Turbanella cornuta (Teuchert 1973) showed an aberrant morphology of the protonephridia in that species consisting of up to five monociliary terminal cells, one large canal cell, and an adjacent putative nephridiopore cell. The composition of protonephridia in Dactylopodola baltica and Mesodasys laticaudatus is less complex as their protonephridia are composed of three cells: one terminal, one canal, and one nephridiopore cell (Neuhaus 1987, see also Bartolomaeus & Ax 1992). However, although quite similar in morphology and ultrastructure, slight differences were also detected: although most gastrotrich species, as far as we know now, lack a cilium in the nephridiopore cell, such an organelle was found in the lumen of the nephridiopore cell of Dactylopodola baltica (Neuhaus 1987). Recently, a series of studies on the excretory system of other gastrotrich species has been carried out (Kieneke et al. 2007, 2008c). The aim of these studies was to get a broader database of more species to infer the evolution of these organs as well as the character pattern of the stem species of Gastrotricha. Besides the systematically important species Neodasys chaetonotoideus (see chapter Phylogeny), two members of the derived taxon Paucitubulatina were studied. Data on Xenotrichula carolinensis and Chaetonotus maximus proved that the protonephridial system of the stem species of Paucitubulatina, whose monophyly is supported in almost all phylogenetic analyses (see chapter Phylogeny), consists of a single pair of protonephridia, each composed of a bicellular terminal organ with a so-called composite filter, a voluminous and aciliar canal cell with a convoluted lumen, and a nephridiopore cell (Kieneke et  al. 2008c). Such a basic protonephridial anatomy is also much likely in the freshwater paucitubulatinan species Polymerurus nodicaudus (Kieneke & Hochberg 2012). The excretory system in Neodasys chaetonotoideus consists, as well as in macrodasyids, of serial pairs (three) of tricellular protonephridia. Although the terminal and canal cell each have one cilium, it is lacking in the nephridiopore cell (Kieneke et  al. 2007). Unique character states, possibly apomorphic for the genus Neodasys, are the number of seven instead of the usually found eight circumciliary microvilli in the terminal cell and a bundle of numerous long ciliary rootlets in the canal cell.

 1 Gastrotricha

C

B

A

D

pn

50 µm

50 µm

50 µm

40 

F E

50 µm

50 µm

50 µm

Taking data from the published ultrastructure-based reconstructions of gastrotrich protonephridia (plus scattered results provided in Ruppert 1991) and using two phylogenetic hypotheses for character optimization (i.e., Hochberg & Litvaitis 2000, Todaro et  al. 2006a; see Fig. 1.53 A and F), a stable hypothesis for the protonephridial system of the last common ancestor, the stem species of Gastrotricha, was reconstructed (Kieneke et  al. 2007). Owing to several highly conserved characters shared by Neodasys chaetonotoideus, Dactylopodola baltica, and Mesodasys laticaudatus and to a probably derived position of Turbanella cornuta as well as the whole Paucitubulatina, a reconstruction of the gastrotrich ground pattern on the basis of both used phylogenetic scenarios yielded almost equal results. This is also true for most other tree topologies because Paucitubulatina mostly occupy a derived position (see chapter Phylogeny). In the

Fig. 1.26: Different representatives of Gastrotricha (schematic) showing position and numbers of protonephridia (in black). (A) Neodasys chaetonotoideus, dorsal view. (B) Redudasys fornerise, ventral view. (C) Turbanella cornuta, ventral view. (D) Mesodasys laticaudatus, dorsal view. (E) Xenotrichula carolinensis, ventral view. (F) Dactylopodola baltica, dorsal view. Abbreviation: pn, protonephridium. (A and E, modified from Kieneke et al. 2007; C, modified from Teuchert 1967; B, modified from Kisielewski 1987; D and F, modified from Neuhaus 1987.)

following, the protonephridial system as it probably existed in the common ancestor of Gastrotricha is described. Structural novelties in recent taxa with regard to the ground pattern will be presented later. The stem species of Gastrotricha has serial pairs of three-cellular protonephridia that are not connected to each other by a common duct (Fig. 1.27). The exact number of pairs remains unclear because this pattern varies considerably between the different species (see, e.g., Fig. 1.26). Dactylopodola baltica, for instance, has 2 pairs of protonephridia, and in Mesodasys laticaudatus, 11 pairs have been counted (Neuhaus 1987). Each protonephridium is situated in a lateral compartment of the trunk where it is adjacent to other tissues as, for example, the epidermis, musculature, and gut epithelium (Fig. 1.30 F). The nephridiopore cell, or at least a distal bulge of it, is embedded within the ventral epidermis via cellular junctions. A thin layer of ECM (perinephridial

1.2 Morphology 

A

tc

B

bb

ime

E

ci cj

elf mv

ci

mv

elf

ime di

cj

C

pl

cr

ci

bb

cc

D

ci

mv

cj mi ep

mi mv

ac

ci

ep

nc

cj np

cj

cut

ECM) surrounds the whole organ. This matrix can slightly be thickened in the filter region of the terminal cell (see, e.g., Kieneke et al. 2007 for Neodasys chaetonotoideus, Fig. 1.30 B–C). The three protonephridial cells are the proximal terminal cell, the canal cell, and the distal nephridiopore cell (Fig. 1.27). All cells are connected to the adjacent one by cellular junctions, a combination of belt desmosomes (probably for the histomechanic connection), and septate junctions that seal the intercellular gap against diffusion processes. Each of the cells forms a lumen by curling around itself and sealing the resulting cleft with an autodesmosome (term sensu Kristensen & Hay-Schmidt 1989). This condition was termed enfolded lumen (Kieneke et  al. 2008c). Linked to each other, the protonephridial lumen forms a continuous duct from the proximal filter region to the distal nephridiopore (Fig. 1.27). In Turbanella cornuta, Dactylopodola baltica, and Mesodasys laticaudatus, the terminal cell is described as a cytoplasmic hollow cylinder broken by pores and short slits (Teuchert 1973, Neuhaus 1987), hence representing a simple filter according to the terminology of Kieneke et al. (2008c). However, Kieneke et al. (2007) suggest an enfolded lumen filter in the ground pattern of Gastrotricha is quite likely (Fig. 1.27 B, C, E). In Neodasys chaetonotoideus, such an enfolded lumen filter is formed by a meandering cleft of the single terminal cell sealed by a diaphragm (Fig. 1.30 C). The same cleft is further distally sealed off by an autodesmosome. Ultrastructural data presented by Teuchert

pl 2 µm

 41

Fig. 1.27: Protonephridium (scheme) of the last common ancestor of Gastrotricha. (A) Longitudinal section through the whole organ showing all 3 cells. Levels of sectional plane for B, C, and D are indicated by bold lines. (B) Cross section of the terminal cell (region of the nucleus). (C) Cross section of the terminal cell (filter region). (D) Cross section of the canal cell. (E) Exterior view of the terminal cell with the enfolded lumen filter. Abbreviations: ac, accessory centriole; bb, basal body; cc, canal cell; ci, cilium; cj, cellular junction (septate junction plus belt desmosome); cr, ciliary rootlet; cut, body cuticle; di, filter diaphragm spanning the cleft; elf, enfolded lumen filter; ep, epidermis; ime, inter-microvillar ECM; mi, mitochondria; mv, microvilli; nc, nephridiopore cell; np, nephridiopore; pl, protonephridial lumen; tc, terminal cell. Black triangles indicate the seam of the enfolded lumen, the broken black line represents the ECM surrounding the whole organ.

(1973) seem to support such filter architecture at least in the protonephridium of T. cornuta because cross sections of the distal part of the terminal cell indicate an enfolded lumen because of the presence of a septate junction in this region of the cell (see Fig. 1.28 C). Such a construction of the terminal cell lumen in T. cornuta was already supposed by Rieger et al. (1974) and supported by their own unpublished results on Turbanella ocellata. The terminal cell of the stem species of Gastrotricha possesses a single cilium projecting into the lumen and is encircled by a column of eight long microvilli. Each of them is linked to the adjacent one by a flocculent ECM (Fig. 1.27 B–C). In some species, however, this inter-microvillar ECM shows a remarkable ultrastructure: in Turbanella cornuta and Chaetonotus maximus, it consists of numerous alternating pores when sectioned tangentially and of alternating electron-lucent and electron-dense bands in cross sections (Teuchert 1973, Kieneke et al. 2008c; Fig. 1.30 E). Interestingly, the fine structure of this inter-microvillar ECM is quite similar to material present between the 10 circumciliary microvilli of external mechanoreceptive sensory cells (“tactile hairs”) of Turbanella cornuta (Teuchert 1976a, see chapter Sensory Structures, Fig. 1.19 B). Such a similarity between terminal cells and epidermal sensory cells could hint to the assumed ectodermal origin of protonephridia. The cilium of the terminal cell of the stem species of Gastrotricha has a basal body but an accessory centriole and rootlet structures are lacking. Of course, there are species

42 

 1 Gastrotricha

di

B

ime

tl

ci

tc sf

ime

bb

mi

tc

C cj

sf

mv

cj

cj cl

cj

bb

A tc

tl

cl

cc

mv

tc

cl nl mv

cj

np

nc

cj

sf 5 µm

Fig. 1.28: Scheme of the derived protonephridium of Turbanella cornuta (Turbanellidae). (A) Schematic longitudinal section, partially exterior view (modified from Teuchert 1973). (B) Cross section of a terminal cell at the filter region. (C) Cross section of a terminal cell at slightly more proximal region than in B. Levels of sectional plane for B and C are indicated by bold lines, cross sections slightly enlarged. Abbreviations: bb, basal body; cc, canal cell; ci, cilium; cj, cellular junction (septate junction plus belt desmosome); cl, canal cell lumen; di, filter diaphragm spanning the pores; ime, inter-microvillar ECM; mi, mitochondria; mv, microvilli; nc, nephridiopore cell; nl, nephridiopore cell lumen; np, nephridiopore; sf, simple filter; tc, terminal cell; tl, terminal cell lumen. The black triangle indicates the seam of the enfolded lumen of the terminal cell, the broken black line in B and C represents the ECM surrounding the whole organ. (Modified from Teuchert 1973.)

that do possess an accessory centriole and ciliary rootlet structures in its protonephridia terminal cells as, for example, Dactylopodola baltica (Neuhaus 1987). The canal cell is interlinked between the proximally adjacent terminal cell and the distally adjacent nephridiopore cell. It constitutes the major part of the protonephridial lumen. The single cilium and a bundle of numerous microvilli (a “brush border”) project into this lumen (enfolded lumen, see above; Figs. 1.27 D, 1.30 G). The large number of microvilli in this cell probably constitutes an extended surface of the luminal cell membrane and hence increases the rate of exchange processes through this membrane. The canal cell cilium of the common ancestor of Gastrotricha has a basal body and a ciliary rootlet, an accessory centriole is lacking (Fig. 1.27 A). The nephridiopore cell constitutes the permanent and cuticle-lined, ventral nephridiopore (Fig. 1.30 H–J). Owing to its ability to secrete a cuticle, the nephridiopore cell, if not the whole protonephridium, is regarded to be of ectodermal origin (see Neuhaus 1987 and above). Due to to equivocal data – some species possess nephridiopore cells with an enclosed lumen, others with an enfolded lumen – is yet not sure which type was present in the common ancestor of Gastrotricha. Although there are

species that have a monociliated nephridiopore cell (i.e., Dactylopodola baltica, Neuhaus 1987), such an organelle is missing in the stem species (Fig. 1.27). However, there is a basal body and possibly an accessory centriole as well (the latter character stayed equivocal, see table 2 in Kieneke et al. 2007). A characteristic set of organelles can be found in each of the protonephridial cells of the studied species (i.e., an active nucleus, mitochondria, rough endoplasmic reticulum, dictyosomes). Mitochondria are relatively abundant in the protonephridial cells to provide energy (or ATP, respectively) for the beating actions of the protonephridial cilia and other energy-dependant transport processes. Especially in the canal cell one can find a high abundance of small (50–100  nm in diameter) vesicles close to the luminal membrane, sometimes coalesced with it. In addition, there are coated pits and coated vesicles (Ruppert 1991). This can hint to transport processes in both directions: excretion as well as re-adsorption via vesicular transport to or from the protonephridial lumen is possible. Species like Xenotrichula carolinensis have large vacuoles in the canal cell that could be used for storing certain degradation products (Kieneke et  al. 2008c).

1.2 Morphology 

tl

B

cc di

ime

ci cf

cc

mi

cj

pcl

cj

nc

to

A

cj

al

pcl

C

to

mv

 43

nc 2 µm

dcl ep

nl cu

np

Fig. 1.29: Scheme of the derived protonephridium of Paucitubulatina as exemplified by Chaetonotus maximus. (A) Schematic longitudinal section, composite filter of the terminal organ as exterior view. (B) Cross section of the terminal organ at the filter region. (C) Cross section of the canal and the nephridiopore cell at the level of the canal cell nucleus. Levels of sectional plane for B and C are indicated as bold lines. B and C slightly enlarged. Abbreviations: al, anterior loop of the canal cell lumen, cc canal cell; cf, composite filter; ci, cilium; cj, cellular junction (septate junction plus belt desmosome); cu, body cuticle; dcl, distal canal cell lumen; di, filter diaphragm spanning the clefts; ep, epidermis; ime, inter-microvillar ECM; mi, mitochondria; mv, microvilli; nc, nephridiopore cell; nl, nephridiopore cell lumen; np, nephridiopore; pcl, proximal canal cell lumen; tl, terminal organ lumen; to, terminal organ (consists of 2 adjacent terminal cells). The black triangles indicate the seam of the enfolded lumen of the canal cell and the nephridiopore cell, the broken black line in B represents the ECM surrounding the whole organ. (Modified from Kieneke et al. 2008c.)

Many evolutionary changes from the above developed character states of the common ancestor of Gastrotricha have at least (as we know today) occurred within the stem lineages of the genus Turbanella and within that of the taxon Paucitubulatina, some of which have already been mentioned earlier. The protonephridial system of Turbanella cornuta, T. ambronensis, and T. subterranea consists of four pairs of protonephridia situated in the middle pharynx region as well as in the anterior, middle, and posterior intestine region (Teuchert 1967, see also Fig. 1.26 C). Each protonephridium of Turbanella cornuta consists of two to four monociliary terminal cells, one voluminous, monociliary canal cell and one smaller, non-ciliated nephridiopore cell that is embedded within the ventral epidermis (Teuchert 1973). The multiple terminal cells neither form a common terminal organ nor an aggregation of cells but are singular cells attached to the canal cell at different sites (Fig. 1.28). According to Teuchert (1973), there is no continuous lumen in the protonephridia of Turbanella cornuta

(i.e., a connection from the terminal cell lumina via the canal cell lumen to the nephridiopore cell lumen). Within the stem lineage of Paucitubulatina, different evolutionary transformations have happened. The number of protonephridia was reduced to a single pair (see Figs. 1.26 E and 1.30 A). Each protonephridium consists of a bicellular terminal organ formed by two monociliary terminal cells that lie close to each other (Fig. 1.30 B), one huge canal cell and one small nephridiopore cell (Fig. 1.29). Both terminal cells form a so-called composite filter by providing reciprocally indented, finger-like processes thus enclosing cilia and microvilli. The cleft between those podocyte-like processes is spanned by a filter diaphragm (Kieneke et al. 2008c, Kieneke & Hochberg 2012). The huge canal cell constitutes the major part of the protonephridial lumen in species of the Paucitubulatina. As in Neodasys and macrodasyidan species, it is formed by an invagination of the cell membrane (enfolded lumen) of the canal and the nephridiopore cell. However, the canal and nephridiopore cell of

44 

 1 Gastrotricha

A

B

G

mv

bb

ecm

*

mg 20 µm

C

cc

400 nm

D va

mv

tc

cc

*

to *

1 µm

* *

*

2 µm

E

F mg

mv

0.5 µm

H

*

*

npc ed

te 5 µm

cir

1 µm

I

J npc 2 µm

*

1 µm

1 µm

Fig. 1.30: Ultrastructure of the protonephridia of Gastrotricha. (A) Chaetonotus sp. (Paucitubulatina) with 1 pair of protonephridia (triangles) already visible under the light microscope (DIC image). (B) Cross section of the terminal organ of Chaetonotus maximus (Paucitubulatina) consisting of 2 monociliary terminal cells. (C) Cross section of the protonephridial terminal cell of Neodasys chaetonotoideus (Multitubulatina). Note the widened extracellular space around the cell. (D) Terminal organ and canal cell of Xenotrichula carolinensis (Paucitubulatina). Note the convoluted protonephridial lumen (asterisks). (E) Oblique longitudinal section of the terminal organ of C. maximus. Note the reticulate material between the circumciliary microvilli (triangle). (F) Position of the protonephridium of X. carolinensis (asterisk) between midgut, muscles and testes. (G) Longitudinal section of the canal cell and nephridiopore cell of N. chaetonotoideus. The protonephridial lumen is densely filled with microvilli (asterisks). (H) Lumen of the nephridiopore cell of C. maximus. Note the autodesmosome of the enfolded lumen (triangle). (I) Nephridiopore cell of N. chaetonotoideus with the distal lumen close to the nephridiopore (triangle). (J) Cuticular tube (asterisk) that continues the nephridiopore of C. maximus. (A) DIC image. (B–J) TEM micrographs. Abbreviations: bb, basal body; cc, canal cell; cir, cirri; ecm, extracellular matrix; ed, epidermis; fi, filter; mg, midgut; mv, circumciliary microvilli of the terminal cell; npc, nephridiopore cell; tc, terminal cell; te, testis; to, terminal organ; va, vacuole.

1.2 Morphology 

paucitubulatinan species lack cilia and microvilli altogether. A remarkable evolutionary novelty is the strong elongation of the canal cell lumen: in all species studied so far, Xenotrichula carolinensis, Chaetonotus maximus, and Polymerurus nodicaudus, the narrow lumen runs strongly convoluted through the canal cell (Kieneke et  al. 2008c, Kieneke & Hochberg 2012; Figs. 1.29, 1.30 D). Although not studied in detail, such a meandering course of an elongated canal cell lumen is also present in Aspidiophorus sp. (figure 16 of Ruppert 1991) and other (all?) members of the Paucitubulatina (see, e.g., light microscopic image 3 of Chaetonotus bisacer and image 38 of Heterolepidoderma sp. in Weiss 2001 and figure 3 on page 235 of Chaetonotus robustus in Kreuz & Foissner 2006). In freshwater species such as C. maximus and P. nodicaudus, the canal cell lumen even forms a prominent anterior loop (see Fig. 1.29), possibly used to utilize the counter current flow effect for concentrating salts, which shall be saved for the organism via re-adsorption (Kieneke & Hochberg 2012). A comparable construction with highly elongated and convoluted protonephridial lumen that forms distinct loops is probably also present in other freshwater-dwelling taxa such as the Neogosseidae (see figure 2S in Todaro et al. 2013). Open questions addressing the protonephridial system of Gastrotricha concern of course the morphology of the filter in macrodasyidan gastrotrichs (i.e., simple filter or enfolded lumen filter), which is yet not satisfyingly known (see Kieneke et al. 2007). Furthermore, protonephridia morphology and ultrastructure is not known for the majority of genera of the Macrodasyida. Among them are taxa like Tetranchyroderma, Thaumastoderma, Pseudostomella, Oregodasys, Acanthodasys, and Diplodasys, all of the family Thaumastodermatidae. As Thaumastodermatidae could be the sister group of Paucitubulatina according to one of the DNA sequence-based cladistic analyses (Todaro et al. 2006a), it would be really interesting to find out whether there are similarities (putative synapomorphies) between the protonephridia of Thaumastodermatidae and Paucitubulatina. To our knowledge, there is just a single published TEM micrograph of a cross section of the protonephridium of a species of Thaumastodermatidae (Diplodasys sp.: see figure 9f of Rieger et  al. 1974). Apart from this special focus on the Thaumastodermatidae, protonephridia ultrastructure has proven its general value for phylogenetic questions, not only in Gastrotricha. Hence, it is of valid interest to increase our knowledge about protonephridia morphology and ultrastructure among the diverse lineages within this phylum. Probably, this will provide new input to the still unresolved internal phylogeny of Gastrotricha (see chapter Phylogeny). Some cross sections through a protonephridium of Chordodasiopsis antennatus, for instance, indicate an aberrant morphology with numerous microvilli inside the terminal cell lumen

 45

that surround the circle of eight circumciliary microvilli of the terminal cell (see figure 9c of Rieger et al. 1974).

1.2.9 Reproductive organs Almost all marine gastrotrich species (the entire Macrodasyida and the basal chaetonotidan groups Neodasys, Xenotrichulidae and Muselliferidae) are hermaphroditic animals with male and female gonads and a set of two accessory structures (at least in Neodasys and Macrodasyida), one of which, the frontal organ, serves as a “female” bursa and sperm-storing device, and the other, the caudal organ, serves as a “male” penis structure (Figs. 1.31 and 1.36 E). This fundamental function of both these structures, at least for the genera Urodasys and Macrodasys, was for the first time correctly interpreted by Schoepfer-Sterrer (1974). Many species additionally possess further structures like certain ducts (e.g., vasa deferentia), glandular tissues like the so-called rosette organ, or the caudal gland cells that can be found in different species of the marine taxon Thaumastodermatidae. Within the Paucitubulatina exclusive of Xenotrichulidae and Muselliferidae, the reproductive system is arranged differently, certainly correlated with a different reproductive biology. Until the first half of the 20st century, only few fresh or brackish water Paucitubulatina were known to develop spermatozoa beside the eggs (e.g., Chaetonotus hermaphroditus), so that it was a common thought that the freshwater inhabiting Paucitubulatina was a group of almost exclusively parthenogenetic organisms (e.g., Remane 1936). However, in the second half of that century more and more reports of freshwater species producing hermaphroditic specimens accumulated and development of spermatozoa in Lepidodermella squamata was studied intensely at the cytological level (Hummon 1984a). Weiss (2001) demonstrated in a comprehensive study the occurrence of such sperm-bearing individuals in selected species of the majority of freshwater-inhabiting genera of the Gastrotricha-Paucitubulatina. Hence, it was concluded that an assumed life cycle of parthenogenetic reproduction followed by the development of hermaphroditic individuals that possibly engage in cross fertilization is widespread among the freshwater taxa (Weiss 2001). The morphology of the reproductive organs (male and female gonads, accessory structures such as the frontal and caudal organs, outlet ducts, gland tissues) were studied in quite a lot species of Gastrotricha at the ultrastructural level (e.g., Ruppert & Shaw 1977, Ruppert 1978a, b, 1991, Hummon 1984a–c, Kieneke et al. 2008d, 2009, Guidi et al. 2011, Todaro et al. 2012a, Guidi et al. 2014). The study of reproductive anatomy based on conventional light microscopy can also provide detailed results (e.g., Ferraguti &

46 

 1 Gastrotricha

B

A

mg

sp

te ao

ph pn ao

lm

lc pn

ut

ep

C

mo pn

te

in

te

mg

hm

D

fo

ut

cu

mp

ov

ip ov

sp

lnb fop fo

fop

cop

mt

ms

E

co ms

mg

lm ao

cop

Balsamo 1994, Fregni et  al. 1999, Balsamo et  al. 2002), but in many cases, especially when regarding the accessory organs, findings are ambiguous. Furthermore, light optic observations do rarely provide information on, for instance, lining epithelia, extracellular material, or inconspicuous genital pores. Comprehensive ultrastructural data of the reproductive organs in putative basal species were missing for a long time. We here briefly review the reproductive morphology of two putative basal taxa, Dactylopodola typhle and Neodasys chaetonotoideus (Kieneke et al. 2008d, Kieneke et al. 2009) because they have been very important for the purpose of reconstructing the character pattern of the common ancestor of Gastrotricha (see Kieneke et al. 2009). These ground pattern features of the reproductive system of Gastrotricha are later presented followed by descriptions of the reproductive organs in further taxa that will highlight some major evolutionary changes.

50 µm

100 µm

co

lnb

Fig. 1.31: Reproductive system (schematic) of the last common ancestor of Gastrotricha. (A) Reproductive anatomy, dorsal view. (B–E) Trunk cross sections at different levels (indicated by bold lines in A). Abbreviations: ao, adhesive organ; co, caudal organ; cop, caudal organ pore; cu, cuticle; ep, epidermis; fo, frontal organ; fop, frontal organ pore; hm, helicoidal muscle; ip, internal pore of the frontal organ; lc, locomotor cilia; lm, longitudinal muscle block; lnb, longitudinal neurite bundle; mg, midgut; mo, mature egg; mp, male genital pore; ms, muscular sheath; mt, mesodermal tissue; ov, ovary; ph, pharynx; pn, protonephridium; sp, spermatozoa; te, testis; ut, uterus.

The study of Dactylopodola typhle revealed a reproductive system generally comparable to that of other species of the Macrodasyida (Kieneke et  al. 2008d). However, there are considerable differences as well. Beside the paired male and female gonads (ovaries mature frontally and fuse to form a common uterus region), there is only one voluminous accessory sex organ, the caudal organ, that is a secretory structure with a complicated branched lumen. Furthermore, there is a cellular duct, the cervix, which communicates with the uterus lumen and possibly has a dual function for (1) the uptake of the foreign spermatophore and (2) for oviposition. The cervix opens dorsolaterally in a single pore that is lined by a rosette of secretory epithelial cells (Fig. 1.32 B). Early oocytes are lined by ECM only but there is an epithelial wall at the uterus region continued by the epithelium of the cervix. The wall of the paired testes is made up of parts of the early germ cells (spermatogonia or spermatocytes) and therefore

1.2 Morphology 

represents a germinal epithelium. On the basis of the morphological reconstruction, a hypothesis for mating and spermatophore transfer in D. typhle has been proposed that suggests a reciprocally transfer of the spermatophore from the central lumen of the glandular caudal organ into the uterus lumen of the mating partner via the cervix. According to that hypothesis, the trunk musculature provides the pressure for draining the caudal organ because it lacks in an own muscular sheath (Kieneke et al. 2008d). For Neodasys chaetonotoideus, the gross anatomy of the reproductive system already given by Remane (1936) and for Neodasys sp. by Ruppert (1991) could generally be confirmed by Kieneke et al. (2009). However, many histological and ultrastructural data now complement our knowledge of the reproductive system of this crucial and probably most basal gastrotrich taxon (Hochberg 2005, Kieneke et al. 2008a, see also chapter Phylogeny). In general, the reproductive system of N. chaetonotoideus consists, like in many macrodasyidan species, of paired lateral testes, an unpaired dorsal and caudally maturing ovary and a set of two accessory reproductive organs, the frontal and caudal organ (Figs. 1.32 A, 1.36 E, and 1.37 A). However, both accessory reproductive structures of N. chaetonotoideus form a common organ (the frontocaudal organ; Fig. 1.37 B) in immature, not inseminated specimens (but with clearly separated lumina), whereas both components seem to be more separated but still in close conjunction in mature and inseminated animals (Fig. 1.37 D). Such a close association or even cellular continuity of the frontal and caudal organs was already highlighted by Ruppert & Shaw (1977) and Ruppert (1978b) for the taxa Dolichodasys and parts of the Thaumastodermatidae (see Figs. 1.34 B and 1.35 C). The testicular wall is, as in D. typhle and other gastrotrich species, a germinal epithelium, whereas distinct sperm ducts are lacking in N. chaetonotoideus. The testes are probably discharged via simple ventral pores. The ovary region is not provided with a wall epithelium. Instead, mesodermal cushion cells, possibly Y-cells, adjoin the early oocytes. More distally, there is a special oviduct epithelium that is obviously engaged in yolk production: The passing egg inside this vitellogenic oviduct endocytes yolk material that is provided by exocytosis of the oviduct cells (Kieneke et al. 2009). Distally, the vitellogenic oviduct is continued by the uterus wall (Fig. 1.32 A). The mature egg inside the uterus undergoes further vitellogenesis by auto synthesis. The uterus lumen directly passes to the lumen of the frontal organ (dorsofrontal lumen of the frontocaudal organ) by a narrow internal pore. In inseminated individuals, a foreign spermatophore is stored in that part of the frontocaudal organ (Fig. 1.37 D). Hence, fertilization via the internal pore is warranted. The frontal organ possesses a laterally directed external pore

 47

for which a dual function is suggested, (1) the uptake of foreign spermatophores and (2) the release of the fully matured and fertilized eggs. The caudal organ of N. chaetonotoideus is hypothesized to engage in external formation of the spermatophore by providing secretions that are probably released through an external pore. There is some evidence that mature specimens of N. chaetonotoideus carry their own spermatophores attached to the caudal adhesive organs (the caudal “feet”) for a while before sticking it to a mating partner (Kieneke et al. 2009). Morphological data of the most satisfyingly investigated gastrotrich species, most of which studied using a multitude of techniques (in vivo light microscopy, histology, TEM, SEM), have been used to reconstruct the character pattern of the reproductive system of the stem species of Gastrotricha (see Kieneke et al. 2009: Table 1). To make this inference, three phylogenetic hypotheses on the internal relationships of Gastrotricha (Hochberg & Litvaitis 2000, Todaro et al. 2006a, Kieneke et al. 2008a; see also chapter Phylogeny) served as the basis for a parsimonious character optimization. Hence, the three resulting ground pattern hypotheses varied in some character states due to topological differences between the three phylogenetic hypotheses. To obtain consistent information of the reproductive system of the stem species of Gastrotricha, a consensus of all three ground pattern hypotheses has been formed (Kieneke et  al. 2009). Based on this procedure, the reproductive system of the common ancestor of Gastrotricha was probably composed of the components listed in the following. It has to be stressed that the most recent studies on the anatomy and ultrastructure of the reproductive tract of Crasiella diplura (Guidi et  al. 2011), Dinodasys mirabilis (Todaro et al. 2012a), and Megadasys sterreri (Guidi et al. 2014) were not considered at that time. The stem species was a hermaphroditic organism with a female gonad that is separated into a proximal ovary region and a distal uterus region that carries one or few mature oocytes and is situated dorsally above the intestine (Fig. 1.31 A, C). Based on the outgroup comparison, the oocytes much likely matured frontocaudally (see discussion on this issue in Kieneke et al. 2009). The early oocytes at the ovary region were not covered with a cellular wall (Fig. 1.31 A, D). Instead, they were surrounded by a thin layer of extracellular material separating them from other adjacent tissues. However, the mature oocyte(s) at the uterus region was/were completely surrounded by a flattened uterus wall epithelium (Fig. 1.31 A, C). It is equivocal if the stem species of Gastrotricha had a paired or an unpaired female gonad; the situation in putative sister taxa of gastrotrichs such as the Cycloneuralia, Gnathifera, or the Plathelminthes is ambiguous (see discussion in Kieneke et  al. 2009). Furthermore, the stem species

48 

 1 Gastrotricha

A

B

in

fp (ro)

100 µm

ph

ph

ce

100 µm

te

in

ut

ov te

mp cop

in

co

ut ip fop

ph

ov

vo

mp fo spp

D

spp

te

ph in

mo

50 µm

co

50 µm

te

sd

ov xo

C

had one pair of long and tube-shaped testes in a lateral body position alongside the intestine (Figs. 1.31 A–C and 1.36 B). The testicular wall consisted of early germ cell stages thus forming a germinal epithelium (Fig. 1.31 B, C). There were no distinct seminal ducts but the testicular lumen probably opened directly to the exterior via a pair of simple ventral pores. The general mode of spermatogenesis was not considered in the reconstruction of the ancestral character pattern. However, in most species of the Macrodasyida studied so far, development of spermatozoa via spermatogonia, spermatocytes, and spermatids proceeds from the periphery of the testes toward the central lumen (centripetal), and from posterior to anterior (see Guidi et al. 2011 and studies referenced therein, see also chapter Spermatogenesis and Spermiogenesis). We therefore assume that the last common ancestor of Gastrotricha also showed this general modality of spermatogenesis.

ov xo

Fig. 1.32: Reproductive system (schematic) of different taxa of Gastrotricha. (A) Neodasys chaetonotoideus (Multitubulatina). (B) Dactylopodola typhle (Macrodasyida, Dactylopodolidae). (C) Lepidodermella squamata (Paucitubulatina, Chaetonotidae). (D) Xenotrichula intermedia (Paucitubulatina, Xenotrichulidae). Abbreviations: ce, cervix; co, caudal organ; cop, caudal organ pore; fo, frontal organ; fop, frontal organ pore; fp, female genital pore; in, intestine; ip, internal pore of the frontal organ; mo, mature egg; mp, male genital pore; ov, ovary; ph, pharynx; ro, rosette organ; sd, sperm duct; spp, spermatophore; te, testis; ut, uterus; vo, vitellogenic oviduct; xo, x-organ.

A cellular (epithelial) frontal organ with a simple lumen opened to the exterior via a lateral to ventral pore (Fig. 1.31 A, D). The frontal organ was situated caudal to the uterus region with the mature oocyte(s). There was a subepithelial ECM surrounding the frontal organ but no specialized musculature. Secretion vesicles were present inside the frontal organ wall epithelium that was probably continuous with the uterus wall epithelium. The frontal organ contained an internal pore that is directed frontally toward the mature oocyte. Foreign spermatozoa or allosperms were kept inside the frontal organ lumen, and fertilization was enabled through the internal pore (Fig. 1.31 A). The stem species of Gastrotricha also possessed a cellular caudal organ provided with a simple, not compartmentalized lumen that opened to the exterior via a ventral pore (Fig. 1.31 A, E). Luminal microvilli and secretory vesicles inside the caudal organ wall cells

1.2 Morphology 

were present. The caudal organ was provided with a thick subepithelial ECM and a sheath of subepithelial circular and longitudinal muscles (Fig. 1.31 E). This organ served for processing the own spermatozoa (autosperms) and for transferring them to the mating partner by means of muscle action. A connection of the frontal and caudal organ was present. The exact structure and shape of this connection, however, could not be reconstructed. There was possibly just a compact column of cubic cells like in Dolichodasys carolinensis (Ruppert & Shaw 1977, see Fig. 1.34 B) that may represent a remnant of a common ontogenetic anlage of both accessory reproductive organs. A direct connection of both accessory organs with communicating lumina as it is present in different species of the derived taxon Thaumastodermatidae (Ruppert 1978b, see Fig. 1.35 C) obviously reflects the derived condition. Comparing the aforementioned ground pattern features with the reproductive system of Macrodasys sp., a species of which the reproductive morphology (see Fig. 1.33 C) as well as the whole mating process and mode of spermatozoa transfer is well-known (Ruppert 1978a, see chapter Reproductive Biology), one can assume that the stem species of Gastrotricha might have engaged in a comparable mode of mating: the own spermatozoa are released through the testicular pores and then transported externally into the caudal organ lumen where they are provided with certain substances such as mucus or nutrients. Two reproductive partners meet and curl around each other, thus bringing the caudal organ pore of one specimen in contact with the frontal organ pore of the second specimen and vice versa. Owing to muscular contraction of the caudal organ, spermatozoa are pressed into the frontal organ lumen of the other specimen, respectively. Given the anatomic conditions reconstructed for the stem species of Gastrotricha, a release of the fertilized eggs through the frontal organ pores (first: passage from the uterus to the frontal organ via the internal pore; second, release of the egg via the external pore of the frontal organ) rather than body wall rupture for oviposition is possible (see discussion in Kieneke et al. 2009). Such a multirole functioning of the accessory reproductive organs was already suggested by Remane (1936). However, there are indeed observations of a body wall rupture during egg deposition in some species, too (Teuchert 1968, see chapter Reproductive Biology; Fig. 1.50 A). Several variations from the character pattern of the stem species can be observed in present-day species and will be reviewed in the following. Most of these can be interpreted as evolutionary transformations of certain internal lineages of the Gastrotricha. However, there still is little knowledge of the anatomy and ultrastructure of these tiny animals, which results in an incomplete

 49

understanding of the structure, the function and evolution of their reproductive system.

1.2.9.1 Female gonad Oocytes in the reconstructed ancestral ovary mature in an anteroposterior direction, a mode that can be found in genera such as Dolichodasys and Neodasys (Ruppert & Shaw 1977, Kieneke et al. 2009, see Figs. 1.32 A and 1.34 B) or in the species Urodasys viviparus, the only viviparous species known so far (see Wilke 1954, Schoepfer-Sterrer 1974; Fig. 1.50 B). In contrast, most other extant gastrotrichs show the opposite pattern. In taxa like, just to give some examples, Macrodasyidae exclusive of Urodasys viviparus (Fig. 1.33 C), Turbanellidae, Dactylopodolidae, Thaumastodermatidae, and Gastrotricha-Paucitubulatina, the eggs mature from posterior to anterior (see Figs. 1.32 C, D, 1.33 A–C, 1.34 A, C, and 1.35 A–C). Flattened epithelial linings of the entire female gonad (early oocytes plus uterus region) are described in different species that have been studied by means of TEM, e.g., Macrodasys sp. 1 and 2 (Ruppert 1978a, see Fig. 1.33 C), Oregodasys (=former Platydasys) cf. ocellatus, Thaumastoderma heideri, Acanthodasys sp., and Diplodasys ankeli (Ruppert 1978b, see Fig. 1.35 A, B). In other species studied by TEM, such thin wall epithelia have been detected in the uterus region lining the mature egg(s). Meanwhile, they could not be found in the ovary region (e.g., Tetranchyroderma bunti: Ruppert 1978b, see Fig. 1.35 C, Dactylopodola typhle: Kieneke et al. 2008d, see Fig. 1.32 B, Neodasys chaetonotoideus: Kieneke et al. 2009, see Figs. 1.32 A and 1.37 A, C) or were proved to be absent at all (e.g., Paraturbanella teissieri: Balsamo et al. 2002, see Fig. 1.34 A, Crasiella diplura: Guidi et  al. 2011, see Fig. 1.33 A, Dinodasys mirabilis: Todaro et al. 2012a). In the latter two cases, the germ cells themselves constitute the gonad wall of the ovaries, i.e., they represent a germinal epithelium (see Todaro et  al. in 2012a), as it is the case in the testes of most Gastrotricha (see below). In Turbanella cornuta, however, there is an incomplete cellular wall of the ovary region (Teuchert 1977a). It was reconstructed unambiguously that a uterus wall epithelium was present in the stem species of Gastrotricha (Kieneke et al. 2009). Although present in many investigated gastrotrich species (see above), an epithelium enclosing the entire female gonad (ovary plus uterus region) is unlikely for the common ancestor of Gastrotricha since putative basal taxa such as Neodasys and Dactylopodola lack a cellular envelope of the ovaries. It is a scenario worth considering that a uterine wall was initially developed as an extension from the wall epithelium

50 

 1 Gastrotricha

of the frontal organ. Later in evolution, this wall may have extended to line the whole female gonad.

1.2.9.2 Male gonad Although reconstructed to be absent in the last common ancestor (this again depends on the condition in the basal taxa Neodasys and Dactylopodola), most contemporary hermaphroditic gastrotrich species possess distinct seminal ducts (vasa deferentia). These are directed caudally where they open into two ventral male gonopores (e.g., in Macrodasys sp.: Ruppert 1978a, see Fig. 1.33 C, Dolichodasys carolinensis: Ruppert & Shaw 1977, see Fig. 1.24 B, also probable in Chordodasiopsis antennatus: Rieger et al. 1974) or in a common unpaired midventral gonopore (e.g., in Crasiella: Schmidt 1974, Guidi et al. 2011, Lee & Chang 2012, see Fig.

1.33 A, also probable in Megadasys: Schmidt 1974, Guidi et al. 2014). However, they project caudally and then sharply bend frontally in species of the taxon Turbanellidae (e.g., Turbanella cornuta: Teuchert 1976b, Paraturbanella teissieri: Balsamo et  al. 2002, see Fig. 1.34 A, Dinodasys mirabilis: Todaro et al. 2012a). This character is hypothesized to be an important autapomorphy of that family (Todaro et  al. 2012a). Also, the paucitubulatinan group Xenotrichulidae has sperm ducts that run frontally (e.g., Ruppert 1979, see Fig. 1.32 D). In both taxa, the paired sperm ducts fuse and open into a common unpaired midventral gonopore. As Xenotrichulidae and Turbanellidae are not close relatives, the anteriorly projecting and fusing sperm ducts must have been developed twice independently in both groups. In Thaumastodermatinae (a subfamily of Thaumastodermatidae, e.g., Tetranchyroderma bunti, Thaumastoderma heideri, Oregodasys cf.

B A

C

ph

ph

ph

in

in mo

te

te

in

mo

ut

ip

sd

sd ov

te ip

mp sp

mp ov

sd

fo fop

mo

ov

fop fo co

ms

ms

sp

co an

cop 100 µm

50 µm

cop

cop 300 µm

co

Fig. 1.33: Reproductive system (schematic) of different taxa of Gastrotricha. (A) Crasiella diplura (Macrodasyida, Planodasyidae). (B) Mesodasys laticaudatus (Macrodasyida, Cephalodasyidae). (C) Macrodasys sp. (Macrodasyida, Macrodasyidae). Abbreviations: an, antrum feminimum; co, caudal organ; cop, caudal organ pore; fo, frontal organ; fop, frontal organ pore (external); in, intestine; ip, internal pore of the frontal organ; mo, mature egg; mp, male genital pore; ms, muscular sheath; ov, ovary; ph, pharynx; sd, sperm duct; sp, spermatozoa; te, testis; ut, uterus.

1.2 Morphology 

ocellatus: Ruppert 1978b), the seminal duct of the single testis distally forms a glandular section and opens into a ventral pore close to the opening of the caudal organ (Fig. 1.35 C). External ventral male gonopores are hence widespread among Gastrotricha and represent the ancestral ­condition. They are not always permanent and well-differentiated openings but rather preformed areas in the body wall that become pores when the testes are fully matured and sperm release is immediately imminent. Such conditions have been reported, e.g., for Dolichodasys carolinensis (Ruppert & Shaw 1977) or Dinodasys mirabilis (Todaro et al. in 2012a), and are also probable in species such as Neodasys chaetonotoideus and Dactylopodola typhle (Kieneke et al. 2008d, 2009). Within species of the taxon Diplodasyinae (the other subfamily of Thaumastodermatidae, e.g., Acanthodasys thrinax and Diplodasys ankeli: Ruppert 1978b) and in Mesodasys (e.g., Mesodasys laticaudatus: Ferraguti & Balsamo 1994, M. adenotubulatus: Fregni et al. 1999), the seminal ducts directly discharge into the caudal organ (Figs. 1.33 B and 1.35 A, B). Hence, the male “gonopores” in these taxa are situated within the wall epithelium of this accessory structure. The functional male gonopore in these species, however, is the unpaired external opening of the caudal organ. As already mentioned, one can observe a further evolutionary transformation related with the male gonads in species of the subfamily Thaumastodermatinae: they possess a well-developed right testis, whereas the left one is completely absent (Ruppert 1978b, see Figs. 1.35 C and 1.36 C). A convergent situation was evolved in another group: in different species of the genus Urodasys (Macrodasyidae), the right testis is reduced or even fully lost, whereas the left one is well developed (see SchoepferSterrer 1974). Although a bounding testicular germinal epithelium is reported from most species studied at the ultrastructural level, there may also be species were the male germ cells are only partly enclosed, e.g., by lateral somatic circular muscles such as in Chordodasiopsis antennatus (Rieger et al. 1974).

1.2.9.3 Frontal and caudal organ Whereas a sac-shaped, hollow frontal organ serving as a seminal receptacle for storing spermatozoa received from the mating partner unambiguously is a component of the reproductive system of the common ancestor, this structure is not present in some gastrotrich species such as Acanthodasys thrinax, Diplodasys ankeli (Ruppert 1978b, see Fig. 1.35 A, B) or Dactylopodola typhle (Kieneke et al. 2008d, see Fig. 1.32 B). In those species, another structure takes the role of storing the foreign sperms: the frontal sac that is a pouch-like differentiation of the uterus or of

 51

the proximal end of the cervix, respectively (Figs. 1.32 B and 1.35 A, B). The latter structure is an epithelial duct putatively serving for egg deposition and picking up the allosperms via a dorsolateral pore (called the “rosette organ” in Acanthodasys thrinax and Diplodasys ankeli, see Ruppert 1978b). The duo-functional cervix, possibly a continuation of the uterus wall epithelium, is so far only described for Dactylopodola typhle (Kieneke et al. 2008d) but is interpreted to be present in even more species (see Kieneke et al. 2009). Like the common ancestor of Gastrotricha, most species of Macrodasyida and Neodasys possess a caudal organ as a copulatory structure (or as a device included in spermatophore formation) that has to be charged with the own sperms prior to copulation. As the male gonopores in many species lie apart from the pore of the caudal organ, spermatozoa have to be transported to the lumen of the caudal organ externally. This procedure has been studied in detail in two species of Macrodasys (Ruppert 1978a, see chapter Reproductive Biology). In taxa such as Mesodasys, Diplodasys, or Acanthodasys, which possess a direct connection of the sperm ducts to the caudal organ (see above), a complicated external transport of spermatozoa is avoided. In Turbanellidae, a real caudal organ is probably lacking (see, e.g., Fig. 1.34 A). Observations of the mating behavior of Turbanella cornuta suggest that these organisms transfer their spermatozoa to a putative frontal organ of the mating partner directly from the unpaired ventral pore of the fused sperm ducts (Teuchert 1968), which makes a caudal organ for sperm transfer redundant (but see for further interpretations of sperm transfer in T. cornuta in the chapter Reproductive Biology). Such kind of sperm transfer modalities presumably represent the general mode in Turbanellidae, whereas structures previously mistaken for caudal organs in different species of that family apparently have a different function, not necessarily related to the reproductive system (Balsamo et al. 2002). However, the recently investigated turbanellid Dinodasys mirabilis possesses a huge glandular and hollow structure, the posterior gland organ, at a comparable position like the caudal organ in many other Macrodasyida. However, the authors suggest that this gland organ of D. mirabilis is not homologous to the “true” copulatory caudal organ of many other taxa (Todaro et al. 2012a). Until now, we have just reported about the presence or absence and the general role of the accessory reproductive organs in Gastrotricha. It has to be stressed that the accessory reproductive organs, especially the caudal organ, belong to the most complex structures in gastrotrichs and show a high morphological diversity among the different taxa (Hummon & Hummon 1988, Balsamo et al. 1999). Although the frontal organ in many cases is

52 

 1 Gastrotricha

B

A

hg

C ov ph te ph

in

te

mo in

mp ip mp

sd

mo

mo sp

fo fop

fop in

ov

50 µm

100 µm

co

sp gt

ov

ms cop 100 µm

fo

tc

a rounded to pear-shaped hollow epithelial pouch (e.g., in species of Urodasys: Schoepfer-Sterrer 1974, Dolichodasys carolinensis: Ruppert & Shaw 1977, see Fig. 1.34 B, species of Thaumastodermatinae: Ruppert 1978b, see Fig. 1.35 C, species of Turbanellidae: Todaro et  al. 2012a and references therein, see Fig. 1.34 A), it can also be a rather tube-shaped structure like in Chordodasiopsis antennatus (Rieger et al. 1974). Species of the genus Macrodasys show a high degree of interspecific structural variability of their frontal organs as well as a complex compartmentalization of this accessory sex organ into a seminal receptacle and a spermatheca (see, e.g., Ruppert 1978a, Evans 1994, Fig. 1.33 C). Further variation of this organ can be found in the position of the external pore: while the pore is situated on the ventral body surface in many taxa (e.g., Dolichodasys carolinensis: Ruppert & Shaw 1977, species of Macrodasys: e.g., Ruppert 1978a, Evans 1994, species of Urodasys:

Fig. 1.34: Reproductive system (schematic) of different taxa of Gastrotricha. (A) Paraturbanella teissieri (Macrodasyida, Turbanellidae). (B) Dolichodasys carolinensis (Macrodasyida, Cephalodasyidae), posterior part of the trunk. (C) Redudasys fornerise (Macrodasyida, Redudasyidae). Abbreviations: co, caudal organ; cop, caudal organ pore; fo, frontal organ; fop, frontal organ pore (external); gt, glandular tissue; hg, hermaphroditic gonad; in, intestine; ip, internal pore of the frontal organ; mo, mature egg; mp, male genital pore; ms, muscular sheath; ov, ovary; ph, pharynx; sd, sperm duct; sp, spermatozoa; tc, tissue connection; te, testis; ut, uterus.

Schoepfer-Sterrer 1974), it opens dorsally to dorsolaterally in some species of Urodasys (Schoepfer-Sterrer 1974) and in Turbanellidae (e.g., Balsamo et al. 2002, Todaro et al. 2012a) or even laterally as in species of Neodasys (Ruppert 1991, Kieneke et al. 2009, see Fig. 1.32 A). In some species of Macrodasys and Urodasys, the internal pore of the frontal organ is provided with cuticularized hard parts, structures that are interpreted to have a valve-like function to release the sperms from the frontal organ lumen to the matured eggs one by one (Schoepfer-Sterrer 1974). The caudal organ lumen can be differentiated as a narrow, branched or non-branched duct (e.g., in Macrodasys sp. 1 and 2: Ruppert 1978a, see Fig. 1.33 C), a highly branched lumen (e.g., Dactylopodola typhle: Kieneke et al. 2008d, see Fig. 1.32 B), or a simple spheric chamber (e.g., species of Thaumastodermatidae, Crasiella, and Mesodasys: Ruppert 1978b, Ferraguti & Balsamo 1994, Fregni et al.

1.2 Morphology 

B

ro

fp

te

in

A

C

ce

ph

 53

fs ip

ph

sp mo

fp ro

in

fs

ce

te

fp?

in ov

sp ip ms

ce

sd mo

co

te

mo ip

gt ms

co cop

fo

sd 100 µm

100 µm

ov

ov

cop 100 µm

sd

ms co mp

cop

1999, Guidi et  al. 2011, see Figs. 1.33 A, B and 1.35 A–C). In species of Macrodasys, the caudal organ is further compartmentalized into a glandulomuscular portion and an antrum feminimum (Ruppert 1978a, Evans 1994, see Fig. 1.33 C). There are species whose caudal organ contains protrudable copulatory tubes with cuticularized teeth-like structures (e.g., Macrodasys sp. 1 and 2: Ruppert 1978a), contractible filaments (e.g., Dolichodasys carolinensis and D. delicatus: Ruppert & Shaw 1977, see Fig. 1.34 B), or even cuticularized stylets or canals (several species of Urodasys: Schoepfer-Sterrer 1974, Oregodasys styliferus: Boaden 1965, Tetranchyroderma bronchostylus: Atherton & Hochberg 2012) (Fig. 1.36 G, H). From an evolutionary point of view, such structures and modifications surely improved the exchange of spermatozoa by means of internal insemination whereas the exact function is hardly known in most cases. In Urodasys spirostylis for instance, the striking resemblance of the corkscrew-shaped stylet with the vagina mouthpiece, the strongly cuticularized external opening of the frontal organ, suggests that the frontal and caudal organs represent a key-lock system (Schoepfer-

gt

Fig. 1.35: Reproductive system (schematic) of different taxa of Gastrotricha. (A) Diplodasys ankeli (Macrodasyida, Thaumastodermatidae, Diplodasyinae). (B) Acanthodasys sp. (Macrodasyida, Thaumastodermatidae, Diplodasyinae). Posterior part of the trunk. (C) Tetranchyroderma bunti (Macrodasyida, Thaumastodermatidae, Thaumastodermatinae). Abbreviations: ce, cervix; co, caudal organ; cop, caudal organ pore; fo, frontal organ; fop, frontal organ pore (external), female genital pore; fs, frontal sac; gt, glandular tissue; in, intestine; ip, internal pore of the frontal organ/frontal sac; mo, mature egg; mp, male genital pore; ms, muscular sheath; ov, ovary; ph, pharynx; ro, rosette organ; sd, sperm duct; sp, spermatozoa; te, testis; ut, uterus.

Sterrer 1974). Another strategy for efficient sperm transfer is the formation and exchange of spermatophores. This is reported at least for some species of the genus Dactylopodola (Teuchert 1968, Kieneke et  al. 2008d) and for Neodasys sp./N. chaetonotoideus (Ruppert 1991, Kieneke et al. 2009). It is hypothesized that the frontal organ or the caudal organ of those species are involved in the production of the spermatophore (Ruppert 1991, Kieneke et  al. 2008d, 2009, see also chapter Reproductive Biology). As already mentioned there are different possible types of connectivity of both accessory reproductive organs (frontal and caudal organ completely apart, simple tissue connection, directly communicating lumina). The situation in Thaumastodermatinae is of special interest because the directly connected and communicating accessory organs represent a challenging anatomic condition. Ruppert’s (1978b) explanation is a duo-role function (male and female) of the caudal organ in Thaumastodermatinae. His hypothesis is that the part of the caudal organ connected with the vas deferens is still male in function but the portion that communicates with the frontal organ has

54 

 1 Gastrotricha

A

B

C *

* mo

50 µm

50 µm

mg

E

D sph

*

mg

* F

*

mo

G H co

50 µm

100 µm

50 µm

50 µm

sp

mg

fo co

fo

ip fo sp

mo

nu spf

50 µm

mg

co

co

25 µm

Fig. 1.36: Reproductive system of Gastrotricha. (A) Xenotrichula intermedia (Paucitubulatina) with a mature egg. Note the size ratio between gamete and organism. (B) Xenodasys riedli (Macrodasyida) with paired testes. Note the spiraled sperm heads (asterisks). (C) Tetranchyroderma sp. (Macrodasyida) in ventral view with a single right testis (asterisk). (D) Middle portion of Oregodasys cf. phacellatus (Macrodasyida) with mature egg and single testis. (E) Posterior trunk portion (horizontal view) of Macrodasys sp. (Macrodasyida) with testes (asterisks), ovary and accessory reproductive organs. (F) Close-up of the accessory reproductive organs of Macrodasys sp. Note the cuticularized internal pore of the frontal organ and the foreign spermatozoa within its lumen. (G) Frontal and caudal organ of Urodasys spirostylis (Macrodasyida). (H) Close-up of the spiraled stylet inside the caudal organ of U. spirostylis. (A) BF image and (B–H) DIC images. Abbreviations: co, caudal organ; fo, frontal organ; ip, internal pore of the frontal organ; mg, midgut; mo, mature egg; nu, nucleus; sp, spermatozoa; spf, flagella of spermatozoa; sph, spiraled heads of spermatozoa.

1.2 Morphology 

A

B

tes tes

fo

ov

mg *

mg co

10 µm

C

E nu

ed

fo

*

mo

*

1 µm

sp

10 µm

F

* 10 µm

D

 55

3 µm

G

tes

mo

* fo

lm

10 µm

2 µm

co

Fig. 1.37: Ultrastructure (TEM micrographs) of the reproductive system of Gastrotricha. (A–D) Neodasys chaetonotoideus (Multitubulatina). (A) Cross section showing paired lateral testes and unpaired dorsal ovary. (B) Cross section through the frontocaudal organ. Note the lumen of the frontal organ (asterisk). (C) Median section showing a mature oocyte within the uterus region. (D) More sagittal section showing a testis with developing spermatozoa, the frontal organ, and the caudal organ. Note the spermatophore inside the frontal organ (asterisk). (E and F) Cross sections of Dactylopodola typhle (Macrodasyida). (E) Right testis with spermatogonia or spermatids in the dorsal part (asterisks) and maturing sperm cells below. (F) Synaptonemal complexes within the testicular germ cells of D. typhle are indicative for meiosis. (G) Cross section through 1 testis of Xenotrichula carolinensis (Paucitubulatina). Spermatozoa inside the lumen are sectioned at different regions (nuclear, mitochondrial, flagellar). Abbreviations: co, caudal organ; ed, epidermis; fo, frontal organ; lm, longitudinal muscles; mg, midgut; mo, mature egg; nu, nucleus of the mature egg; ov, ovary; sp, sperm cells; tes, testis.

been modified to a female accessory structure (see also discussion of Atherton & Hochberg 2012). In this regard, another functional scenario is quite interesting: the distal part of the vas deferens (glandular and/or provided with a muscular sheath) in Thaumastodermatinae evolved as a

new structure. The caudal organ is still male in function (uptake and transfer of spermatozoa), the frontal organ still female (receive and storage of spermatozoa). According to this scenario, the direct connection of both accessory organs would imply that species of Thaumastodermatinae

56 

 1 Gastrotricha

are able to engage in self-fertilization (see Ruppert 1977). However, even if anatomically possible it does not seem that self-fertilization occurs (Balsamo 1992).

1.2.9.4 Reproductive system of the Paucitubulatina The whole reproductive system, together with the whole life history, has strongly been modified in the gastrotrich subtaxon Paucitubulatina. Putative basal paucitubulatinan groups such as the marine Xenotrichulidae (Figs. 1.32 D and 1.37 G) and Muselliferidae are still hermaphrodites that develop full male and female gonads during their life history (e.g., Ferraguti et  al. 1995, Guidi et  al. 2003a, Balsamo et al. 2010a). Species of the genus Musellifer might be the only members of Paucitubulatina that possibly possess an accessory reproductive organ in addition to the gonads (Hummon 1969). However, the presence of any accessory organ could not be supported for at least Musellifer profundus (Leasi & Todaro 2010). The more derived and predominantly freshwater taxa (traditional families Proichthydidae, Chaetonotidae, Dasydytidae, Neogosseidae) show a different morphological and temporal arrangement of reproductive structures. As a model organism, Lepidodermella squamata was studied intensely to reveal gametogenesis and reproductive biology in a freshwater gastrotrich (Hummon 1984a–c, 1986). In this species, populations consist of individuals that possess a paired female gonad producing subitanuous eggs (tachyblastic eggs) by parthenogenesis. These animals can also produce parthenogenetic resting eggs (opsiblastic eggs) with a resistant eggshell. Specimens, possibly each individual of a population (see discussion in Weiss 2001) enter a postparthenogenetic stage and cysts with simple rod-shaped, filiform, spindle-shaped, or oval male germ cells (simplified spermatozoa) are formed (Figs. 1.32 C and 1.41 B–F). There are no proper testes with wall epithelia or outlet ducts; however, a vesiculated region in the ventral epidermis of Lepidodermella squammata was observed close to each sperm packet and interpreted as male genital pores (Hummon & Hummon 1983b, but see also discussion in Weiss 2001). The cysts with 9–16 simplified non-flagellated sperm cells in various Paucitubulatina generally occur in one or two ventral pairs, while up to 12 sperm packets in one individual are possible (Weiss 2001). Within the ovary, a further resting egg is matured in these “sperm bearers”. Although not proven, a sexual reproduction with recombination, possibly with cross-fertilization in Lepidodermella squamata is assumed (Hummon 1986). Because sperm bearers were discovered in dozens of freshwater-dwelling species (Weiss 2001), a life history of initially pure

parthenogenetic populations with later occurring hermaphroditic animals is proposed for all Paucitubulatina exclusive of the ancestrally marine taxa Xenotrichulidae and Muselliferidae (see chapter Reproductive Biology). It has to be mentioned that sperm-bearing freshwater gastrotrichs as well as fully mature individuals of the Xenotrichulidae exhibit an accessory structure referred to as the x-body or x-organ (Fig. 1.32 C, D). This secretory structure derives from undifferentiated female germ cells (as demonstrated in Lepidodermella squamata: Hummon 1984c) and appears in the region of the hind gut. It is regarded to play a role in sexual reproduction and was interpreted as a copulatory organ, a bursal organ, an oviduct, or a vitellarium repeatedly. However, all these hypotheses have never been confirmed (see discussion in Weiss 2001 and references therein). According to Hummon (1986), the cells of the developing x-body in L. squamata also contribute to the vitellogenesis of the postparthenogenic egg.

1.2.9.5 Additional accessory structures Glandular tissues comparable to the above mentioned paucitubulatinan x-organ are known from several species of the Macrodasyida, too. Here, they are associated with the distal parts of the seminal ducts (Figs. 1.34 A and 1.35 A, C) and/or with the pore of the caudal organ (Fig. 1.35 C) and lie in close proximity to the hindgut and anus (e.g., species of Urodasys: Schoepfer-Sterrer 1974, Tetranchyroderma bunti, Thaumastoderma heideri, Diplodasys ankeli: Ruppert 1978b). Comparable structures may be present in species that lack a caudal organ altogether such as in Paraturbanella teissieri and other species of the Turbanellidae (Balsamo et al. 2002). In these cases, the glandular organ is also associated with the hindgut (Fig. 1.34 A). However, homology of the described structures with the x-organ of Paucitubulatina is improbable. Furthermore, there is no convincing hypothesis on the functional role of those “glands”, neither for the paucitubulatinan x-organ (see above) nor for glandular structures in different Macrodasyida. It is imaginable that these organs produce and release water-soluble substances used for chemical communication, for instance, to aid those small organisms attracting a proper reproductive partner. Such a function was already suggested for the x-organ of Lepidodermella squamata (Hummon 1986). Because these glands in Macrodasyida may be associated with the sperm ducts and/or the caudal organ, it seems also likely that they produce substances (e.g., mucus) that support sperm transfer. The recently described posterior gland organ of Dinodasys mirabilis is hypothesized to be used for gluing the fertilized eggs to sand grains. However, it is unlikely

1.2 Morphology 

that the posterior gland organ of D. mirabilis and the real caudal organ, present in most macrodasyidan species, are homologue features (Todaro et al. 2012a).

1.2.10 Gametes 1.2.10.1 Spermatozoa In early monographs (e.g., Zelinka 1889), the existence of male gonads as well as spermatozoa in Gastrotricha was keenly doubted. It should be noted that at that time almost only freshwater species of the taxon Paucitubulatina were known who develop highly simplified sperm cells in a rather short period of their lifespan. Those hermaphroditic specimens attracted little attention for a long time. The discovery of many new taxa of the gastrotrich subgroup Macrodasyida in the early 20th century (e.g., Remane 1924, 1927a) revealed that at least the new marine species were hermaphrodites with testes and the ability to develop proper spermatozoa. First descriptions of sperms based on light microscopy can be found in Remane’s monographs (Remane 1927c, 1929, 1936). In the later work (Remane 1936), he depicts drawings of the long and partly spiraled sperm cells of Oregodasys maximus and Ptychostomella ommatophora as well as of the short-tailed and compact sperm of Neodasys chaetonotoideus. Remane (1927c) already introduced the two general sperm types of the Gastrotricha, the nowadays termed (a) filiform spermatozoon (“fadenförmig”) of most Macrodasyida and many Paucitubulatina and the (b) commaform spermatozoon (“eiförmig mit kurzem Schwanz”=ovoid with short tail) of Neodasys. Later, microscopic observations of further macrodasyidan species confirmed the filiform sperm type with spiraled sperm head and/or middle region, for instance, in Mesodasys laticaudatus (synonym: Cephalodasys lobocercus), Cephalodasys cambriensis (synonym: Paradasys cambriensis), and Oregodasys styliferus (see Boaden 1960, 1963, 1965). However, those pure light microscopic observations already demonstrated a certain morphological complexity and diversity among gastrotrich spermatozoa. Microscopic observations of spermatozoa of different species of the marine paucitubulatinan taxon Xenotrichulidae (Wilke 1954) demonstrated a filiform sperm type without any spiraled parts in that group. Furthermore, the sperms of Heteroxenotrichula squamosa displayed some peculiar unknown differentiations, the two “thin accessory flagella”, today known as the para-acrosomal bodies (see below). A misinterpretation that unites almost all of the early light-optical investigations of sperm morphology in Gastrotricha is the assumption that the anterior tip of

 57

the filiform cell represents the nucleus (see, e.g., Remane 1936, Wilke 1954). Resolving power of the TEM opened a window to an unexpected ultrastructural complexity of sperm cells. The first comprehensive ultrastructural study of spermatozoa (and sperm development, see below) in Gastrotricha was carried out by Teuchert (1975b, 1976b) who investigated formation and submicroscopic morphology of sperms of Turbanella cornuta. This pioneering study was shortly followed by an investigation of sperm transfer modalities in an undetermined species of Macrodasys that also contains many data on sperm ultrastructure (Ruppert 1978a). The mass of sperm cell reconstructions based on TEM studies was carried out during the past 20 years beginning in the mid-1990s. Since the first investigation of Teuchert (1976b), spermatozoa of at least 33 species from all major subtaxa of the Gastrotricha have been studied ultrastructurally (see Tab. 1.5): Neodasys/Multitubulatina, 1 species (Guidi et al. 2003a), Paucitubulatina, 7 species (Balsamo 1992, Balsamo et  al. 2010a, Ferraguti et  al. 1995, Guidi et al. 2003a, Hummon 1984b), and Macrodasyida, 25 species (Balsamo et al. 2002, 2007, Ferraguti & Balsamo 1994, 1995, Fischer 1994, 1996, Fregni et al. 1999, Guidi et al. 2004, 2009, 2011, Pierboni et al. 2003, 2004, Marotta et al. 2005, Pierboni & Kristensen 2007, Ruppert 1978a, Teuchert 1976b, Todaro et al. 2000a, 2012a). Most recently, Guidi et al. (2014) published the reconstruction of the spermatozoon of yet another taxon of the Macrodasyida, Megadasys sterreri (not yet included in Tab. 1.5). The general gastrotrich sperm types already described by Remane (1927c, 1929, 1936) and Wilke (1954) were supported by the ultrastructural studies: filiform sperms with spiraled head in most Macrodasyida (Fig. 1.38 A–F), filiform sperms without spiraled regions in (basal) Paucitubulatina (Fig. 1.38 G–M), the latter sometimes with elongated para-acrosomal bodies (Fig. 1.40 A–D), and commaform sperms in Neodasys (Fig. 1.39 A–F). Because of a certain degree of similarity between spermatozoa of different species of the Macrodasyida, a “basic sperm plan” for that group could be compiled. According to this, the macrodasyidan sperm is composed of an acrosomal anterior region, a nuclear central region, and a flagellar distal region. The long and corkscrewshaped acrosome is often composed of two different portions (anterior and basal) and contains an internal striated tube. The spring-shaped nucleus surrounds several small or one giant, tube-shaped mitochondrion. The flagellum (cilium) is composed of a normal axoneme (9×2+2 pattern of microtubules) that is surrounded by a striated cylinder (Balsamo et al. 1999, Ferrraguti & Balsamo 1995, Marotta et al. 2005; Fig. 1.44 C–G). The spermatozoon of

Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform Filiform-aciliar Filiform-aciliar Filiform Filiform Commaform Rod-like Rod-like Filiform Filiform Filiform Filiform Filiform

0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 1 1 0 0 0 0 0

0

Vestigial spermatozoa

Sperm shape

Lepidodasys sp. Lepidodasys unicarenatus Lepidodasys sp. (different species than in Guidi et al. 2004) Mesodasys laticaudatus Mesodasys adenotubulatus Acanthodasys aculeatus (see Fig. 1.38 A–F) Diplodasys ankeli Pseudostomella etrusca Tetranchyroderma sp. 1 Tetranchyroderma sp. 2 Tetranchyroderma papii Urodasys anorektoxys Urodasys acanthostylis Macrodasys sp. Macrodasys caudatus Crasiella diplura Turbanella cornuta (see Fig. 1.43 E–H) Turbanella ambronensis Paraturbanella teissieri Dinodasys mirabilis Dactylopodola baltica (see Fig. 1.41 A) Dactylopodola typhle Xenodasys sp. Xenodasys eknomios Neodasys ciritus (see Fig. 1.39 A–F) Chaetonotus maximus Lepidodermella squamata (see Fig. 1.41 B) Musellifer delamarei (see Fig. 1.38 G–M) Diurunotus aspetos Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata (see Fig. 1.40 A–D)

2

1

Cephalodasys maximus

Character-number (1–33) according to Marotta et al. (2005)

General morphology

Tab. 1.5: Morphological and ultrastructural characters of spermatozoa of 33 species of Gastrotricha.

1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 0a 0a 1 1 1 0 0 1 1 1 1 1

1

Flagellum (cilium) of mature sperm

3

0 0 0 0 0 0 0 0 0 0 0 1 1 0 0 0 0 0 0 0 N.A. N.A. 0 0 0 N.A. N.A. 1 1 1 1 1

1

Accessory fibers or globules

4

N.A. N.A. N.A. Multilayered Multilayered Monolayered Monolayered Monolayered Monolayered Monolayered ? N.A. Monolayered Monolayered Monolayered N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A. N.A.

0 0 1 1 1 1 1 1 1 1 0 1 1 1 0 0 0 0 0 N.A. N.A. 0 0 0 N.A. N.A. 0 0 0 0 0

(Continued)

Monolayered

0

Striated cylinder thickness

6

1

Striated cylinder

5

Flagellum/cilium

58   1 Gastrotricha

Cephalodasys maximus Lepidodasys sp. Lepidodasys unicarenatus Lepidodasys sp. (different species than in Guidi et al. 2004) Mesodasys laticaudatus Mesodasys adenotubulatus Acanthodasys aculeatus (see Fig. 1.38 A–F) Diplodasys ankeli Pseudostomella etrusca Tetranchyroderma sp. 1 Tetranchyroderma sp. 2 Tetranchyroderma papii Urodasys anorektoxys Urodasys acanthostylis Macrodasys sp. Macrodasys caudatus Crasiella diplura Turbanella cornuta (see Fig. 1.43 E–H) Turbanella ambronensis Paraturbanella teissieri Dinodasys mirabilis Dactylopodola baltica (see Fig. 1.41 A) Dactylopodola typhle Xenodasys sp. Xenodasys eknomios Neodasys ciritus (see Fig. 1.39 A–F) Chaetonotus maximus Lepidodermella squamata (see Fig. 1.41 B) Musellifer delamarei (see Fig. 1.38 G–M) Diurunotus aspetos Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata (see Fig. 1.40 A–D)

Character-number (1–33) according to Marotta et al. (2005)

Tab. 1.5: (Continued)

Bubble of cytoplasm ? ? ? ? Thin Hollow+thin ? ? Hollow+thin ? ? Thin Thin ? Twisted ? ? ? ? Rounded N.A. N.A. ? ? ? N.A. N.A. Thin Thin Thin Thin Thin

Helical Parallel Parallel Parallel Helical Parallel Parallel Parallel Helical ? ? Helical Parallel Parallel Parallel Parallel Helical Helical Helical Helical N.A. N.A. Parallel Parallel Parallel N.A. N.A. Parallel Parallel Parallel Parallel Parallel

Endpiece

Axonemal microtubule arrangement Parallel

8

7

Not swollen

Not swollen

Not swollen

Not swollen

Not swollen

N.A.

N.A.

Not swollen

Not swollen

Not swollen

N.A.

N.A.

Slightly swollen

Swollen

Swollen

Swollen

Swollen

Swollen

Swollen

Not swollen

Not swollen

?

Swollen

Swollen

Swollen

Not swollen

Not swollen

Not swollen

Not swollen

Not swollen

Not swollen

Not swollen

Swollen

Axonemal plasma membrane

9

Flagellum/cilium

One

One

One

0

One

N.A.

N.A.

Two

One

One

N.A.

N.A.

One

One

One

One

One

One

One

One

One

One

One

One

One

One

One

One

One

One

One

One

One

(Continued)

Number of centriolesb

10

1.2 Morphology 

 59

Cephalodasys maximus Lepidodasys sp. Lepidodasys unicarenatus Lepidodasys sp. (different species than in Guidi et al. 2004) Mesodasys laticaudatus Mesodasys adenotubulatus Acanthodasys aculeatus (see Fig. 1.38 A–F) Diplodasys ankeli Pseudostomella etrusca Tetranchyroderma sp. 1 Tetranchyroderma sp. 2 Tetranchyroderma papii Urodasys anorektoxys Urodasys acanthostylis Macrodasys sp. Macrodasys caudatus Crasiella diplura Turbanella cornuta (see Fig. 1.43 E–H) Turbanella ambronensis Paraturbanella teissieri Dinodasys mirabilis Dactylopodola baltica (see Fig. 1.41 A) Dactylopodola typhle Xenodasys sp. Xenodasys eknomios Neodasys ciritus (see Fig. 1.39 A–F) Chaetonotus maximus Lepidodermella squamata (see Fig. 1.41 B) Musellifer delamarei (see Fig. 1.38 G–M) Diurunotus aspetos Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata (see Fig. 1.40 A–D)

Character-number (1–33) according to Marotta et al. (2005)

Tab. 1.5: (Continued)

0 0 0 1 0 0 0 0 0 ? ? ? 1 0 ? ? 0 1 1 N.A. 0 N.A. N.A. 0 ? 1 N.A. N.A. N.A. N.A. 0 0 N.A.

1 1 1 1 1 1 1 1 ? ? ? 1 1 ? 0 0 1 1 0 1 N.A. N.A. 1 1 1 N.A. N.A. 0 0 1 1 0

Cap-like fibers

Cap-like,or conical structure 1

12

11

0

0

0

0

0

N.A.

N.A.

0

0

0

N.A.

N.A.

0

0

0

0

1

0

0

0

0

0

0

0

0

0

0

0

0

0

0

0

0

Sheath around central microtubules

Flagellum/cilium

0

0

0

0

0

N.A.

N.A.

0

0

0

N.A.

N.A.

0

0

0

0

1

0

0

0

0

0

0

0

0

0

0

0

0

0

0

0

0

(Continued)

Curled plasma membrane at flagellum

60   1 Gastrotricha

Cephalodasys maximus Lepidodasys sp. Lepidodasys unicarenatus Lepidodasys sp. (different species than in Guidi et al. 2004) Mesodasys laticaudatus Mesodasys adenotubulatus Acanthodasys aculeatus (see Fig. 1.38 A–F) Diplodasys ankeli Pseudostomella etrusca Tetranchyroderma sp. 1 Tetranchyroderma sp. 2 Tetranchyroderma papii Urodasys anorektoxys Urodasys acanthostylis Macrodasys sp. Macrodasys caudatus Crasiella diplura Turbanella cornuta (see Fig. 1.43 E–H) Turbanella ambronensis Paraturbanella teissieri Dinodasys mirabilis Dactylopodola baltica (see Fig. 1.41 A) Dactylopodola typhle Xenodasys sp. Xenodasys eknomios Neodasys ciritus (see Fig. 1.39 A–F) Chaetonotus maximus Lepidodermella squamata (see Fig. 1.41 B) Musellifer delamarei (see Fig. 1.38 G–M) Diurunotus aspetos Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata (see Fig. 1.40 A–D)

Character-number (1–33) according to Marotta et al. (2005)

Tab. 1.5: (Continued)

Spiral/slightly spiral Ribbon-like Ribbon-like Ribbon-like Ribbon-like Ribbon-like Ribbon-like Ribbon-like Spiral/slightly spiral Ribbon-like Ribbon-like Ribbon-like Spiral+straight (at base) Complex spiral Spiral/slightly spiral Spiral/slightly spiral Spiral/slightly spiral Ribbon-like Ribbon-like Ribbon-like Ribbon-like Alternate layers Alternate layers Ribbon-like Ribbon-like Straight Straight Straight Straight Straight Straight Straight Straight

Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Spiral/helical Straightc Spiral/helical Spiral/helical Spiral/helical Straight Straight Straight Straight Straight Straight Straight Straight

Shape of nucleus

Shape of sperm head Spiral/helical

14

13

0

0

0

1

1

0

0

0

1

1

0

0

0

0

?

0

0

0

?

1

1

0

0

0

0

0

0

1

1

0

0

0

1

Reduction of nuclear diameter

15

Nucleus/sperm head

Partial

Partial

Partial

Complete

Partial

?

?

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Fingerprint-like

Fingerprint-like

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

Complete

(Continued)

Chromatin condensation

16

1.2 Morphology 

 61

Cephalodasys maximus Lepidodasys sp. Lepidodasys unicarenatus Lepidodasys sp. (different species than in Guidi et al. 2004) Mesodasys laticaudatus Mesodasys adenotubulatus Acanthodasys aculeatus (see Fig. 1.38 A–F) Diplodasys ankeli Pseudostomella etrusca Tetranchyroderma sp. 1 Tetranchyroderma sp. 2 Tetranchyroderma papii Urodasys anorektoxys Urodasys acanthostylis Macrodasys sp. Macrodasys caudatus Crasiella diplura Turbanella cornuta (see Fig. 1.43 E–H) Turbanella ambronensis Paraturbanella teissieri Dinodasys mirabilis Dactylopodola baltica (see Fig. 1.41 A) Dactylopodola typhle Xenodasys sp. Xenodasys eknomios Neodasys ciritus (see Fig. 1.39 A–F) Chaetonotus maximus Lepidodermella squamata (see Fig. 1.41 B) Musellifer delamarei (see Fig. 1.38 G–M) Diurunotus aspetos Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata (see Fig. 1.40 A–D)

Character-number (1–33) according to Marotta et al. (2005)

Tab. 1.5: (Continued)

18

With fossa Convex Convex N.A. N.A. With fossa Convex

Concave Convex Convex N.A. N.A. Flat/slightly concave Convex

Flat

Flat

Flat/slightly concave

Flat/slightly concave

With fossa

Flat/slightly concave

With fossa

With fossa

Flat/slightly concave

Flat

With fossa

Flat/slightly concave

Flat/slightly concave

With fossa

Flat/slightly concave

Flat/slightly concave

?

Convex

With deep fossa

With fossa

Convex+spiraled

Convex

With fossa

Convex+spiraled

Convex

?

Flat/slightly concave

?

?

Flat/slightly concave

Flat/slightly concave

Flat

Flat/slightly concave

Convex

Flat

?

Flat/slightly concave

?

Flat/slightly concave Slightly concave

Slightly concave

Flat/slightly concave

Slightly concave

With fossa

Flat/slightly concave

Flat/slightly concave

Slightly concave

Flat/slightly concave

Convex

Slightly concave Slightly concave

Convex

Shape of nuclear apex Shape of nuclear base

17

Nucleus/sperm head

1

1

1

1

1

0

0

1

1

1

1

1

1

1

1

1

1

1

1

0

0

1

1

1

1

1

1

1

1

1

1

1

1

Mitochondria

19

One small

One small

One small

One giant

More than 1 (small)

N.A.

N.A.

More than 1 (small)

More than 1 (small)

More than 1 (small)

More than 1 (small)

More than 1 (small)

One giant

One giant

One giant

One giant

One giant

One giant

One giant

N.A.

N.A.

?

?

One giant

More than 1 (small)

One giant

One giant

One giant

More than 1 (small)

One giant

One giant

One giant

One giant

Number/size of mitochondria

20

At nuclear base

At nuclear base

At nuclear base

(Continued)

Around nuclear base

At nuclear base

N.A.

N.A.

Randomly around head

Around connecting piece

Around connecting piece

Inside nucleus

Along nucleusd

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Spirally around head

Spirally around head

N.A.

N.A.

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Inside nucleus

Position/arrangement of mitochondria

21

Mitochondrium/mitochondria

62   1 Gastrotricha

Cephalodasys maximus Lepidodasys sp. Lepidodasys unicarenatus Lepidodasys sp. (different species than in Guidi et al. 2004) Mesodasys laticaudatus Mesodasys adenotubulatus Acanthodasys aculeatus (see Fig. 1.38 A–F) Diplodasys ankeli Pseudostomella etrusca Tetranchyroderma sp. 1 Tetranchyroderma sp. 2 Tetranchyroderma papii Urodasys anorektoxys Urodasys acanthostylis Macrodasys sp. Macrodasys caudatus Crasiella diplura Turbanella cornuta (see Fig. 1.43 E–H) Turbanella ambronensis Paraturbanella teissieri Dinodasys mirabilis Dactylopodola baltica (see Fig. 1.41 A) Dactylopodola typhle Xenodasys sp. Xenodasys eknomios Neodasys ciritus (see Fig. 1.39 A–F) Chaetonotus maximus Lepidodermella squamata (see Fig. 1.41 B) Musellifer delamarei (see Fig. 1.38 G–M) Diurunotus aspetos Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata (see Fig. 1.40 A–D)

Character-number (1–33) according to Marotta et al. (2005)

Tab. 1.5: (Continued)

23 Shape of acrosome Corkscrew Cylindrical+corkscrew Cylindrical+corkscrew Cylindrical+corkscrew Cylindrical+corkscrew ? Corkscrew Corkscrew Corkscrew Corkscrew Corkscrew Corkscrew Cylindrical Helical Cylindrical+corkscrew Cylindrical+corkscrew Cylindrical+corkscrew Cylindrical+corkscrew Cylindrical+corkscrew Cylindrical Cylindrical+corkscrew Cylindrical Cylindrical Cylindrical Cylindrical Pear-shaped N.A. N.A. Cylindrical Cylindrical Cylindrical N.A. Cylindrical

22 Acrosome 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 0 0 1 1 1 0 1

25

26

With different regions With different regions With different regions With different regions With different regions ? With different regions With different regions With different regions With different regions With different regions With different regions With different regions Homogeneous With different regions With different regions With different regions With different regions With different regions With different regions With different regions ? ? With different regions With different regions With different regions N.A. N.A. With different regions Two cones Homogeneous N.A. Homogeneous

0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 1 1 1 1 0 0 0 1 0 N.A. N.A. 0 0 0 N.A. 0

1 0 0 0 0 0 1 1 1 1 1 1 1 0 1 1 1 0 0 0 1 0 0 0 0 1 N.A. N.A. 0 0 0 N.A. 0 (Continued)

Composition of acrosome Acrosomal thick Acrosomal tubular disks structure

24

Acrosome

1.2 Morphology 

 63

Cephalodasys maximus Lepidodasys sp. Lepidodasys unicarenatus Lepidodasys sp. (different species than in Guidi et al. 2004) Mesodasys laticaudatus Mesodasys adenotubulatus Acanthodasys aculeatus (see Fig. 1.38 A–F) Diplodasys ankeli Pseudostomella etrusca Tetranchyroderma sp. 1 Tetranchyroderma sp. 2 Tetranchyroderma papii Urodasys anorektoxys Urodasys acanthostylis Macrodasys sp. Macrodasys caudatus Crasiella diplura Turbanella cornuta (see Fig. 1.43 E–H) Turbanella ambronensis Paraturbanella teissieri Dinodasys mirabilis Dactylopodola baltica (see Fig. 1.41 A) Dactylopodola typhle Xenodasys sp. Xenodasys eknomios Neodasys ciritus (see Fig. 1.39 A–F) Chaetonotus maximus Lepidodermella squamata (see Fig. 1.41 B) Musellifer delamarei (see Fig. 1.38 G–M) Diurunotus aspetos Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata (see Fig. 1.40 A–D)

Tubular structure organization Ring-like N.A. N.A. N.A. N.A. N.A. Ring-like Ring-like Ring-like Ring-like ? Ring-like Ring-like N.A. Continuous Continuous Helical+radial N.A. N.A. N.A. Continuous N.A. N.A. N.A. N.A. Tubular structure-like N.A. N.A. N.A. N.A. N.A. N.A. N.A.

Uncondensed Condensed at base Condensed at apex Condensed at apex Condensed at apex ? Uncondensed Uncondensed Uncondensed Uncondensed Uncondensed Uncondensed Uncondensed Uncondensed Uncondensed Uncondensed Condensed at apex Condensed at base Condensed at base Condensed at base Condensed at apex Uncondensed Uncondensed Condensed at base Condensed Uncondensed N.A. N.A. Uncondensed Condensed+filamentous Uncondensed N.A. Uncondensed

28

Rectilinear+twisted N.A. N.A. N.A. N.A. N.A. Rectilinear+twisted Twisted Rectilinear Rectilinear Twisted ? Twisted N.A. ? ? ? N.A. N.A. N.A. Obliquely striated N.A. N.A. N.A. N.A. Rectilinear N.A. N.A. N.A. N.A. N.A. N.A. N.A.

Shape of tubular structure

29

Acrosome

Acrosomal material

Character-number (1–33) according to Marotta et al. (2005) 27

Tab. 1.5: (Continued)

0 N.A. N.A. N.A. N.A. N.A. 1 1 0 0 ? ? 1 N.A. 0 0 ? N.A. N.A. N.A. ? N.A. N.A. N.A. N.A. 0 N.A. N.A. N.A. N.A. N.A. N.A. N.A.

0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 ? 1 0 N.A. N.A. 0 0 0 N.A. 0 (Continued)

Tubular structure Ribbon helix around withdrawal acrosome

30

64   1 Gastrotricha

32

33

0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 1 0 1

0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

Urodasys acanthostylis Macrodasys sp. Macrodasys caudatus Crasiella diplura Turbanella cornuta (see Fig. 1.43 E–H) Turbanella ambronensis Paraturbanella teissieri Dinodasys mirabilis Dactylopodola baltica (see Fig. 1.41 A) Dactylopodola typhle Xenodasys sp. Xenodasys eknomios Neodasys ciritus (see Fig. 1.39 A–F) Chaetonotus maximus Lepidodermella squamata (see Fig. 1.41 B) Musellifer delamarei (see Fig. 1.38 G–M) Diurunotus aspetos Heteroxenotrichula squamosa Xenotrichula intermedia Xenotrichula punctata (see Fig. 1.40 A–D)

0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

0 0 0 0 0 0 0 1 0 1 1 0 0

Fischer 1994 Guidi et al. 2004 Guidi et al. 2004 Pierboni & Kristensen 2007 Ferraguti & Balsamo 1994 Fregni et al. 1999 Guidi et al. 2003b Ferraguti & Balsamo 1995 Ferraguti & Balsamo 1995 Ferraguti & Balsamo 1995 Ferraguti & Balsamo 1995 Ferraguti & Balsamo 1995 Todaro et al. 2000a, Pierboni et al. 2003, unpublished/Marotta et al. 2005, Balsamo et al. 2007 Pierboni et al. 2003, unpublished/Marotta et al. 2005, Balsamo et al. 2007 Ruppert 1978a Marotta et al. 2005 Guidi et al. 2011 Teuchert 1976b Ferraguti & Balsamo 1995 Balsamo et al. 2002 Todaro et al. 2012a Fischer 1996 unpublished/Marotta et al. 2005 Pierboni et al. 2004, unpublished/Marotta et al. 2005 Guidi et al. 2009 Guidi et al. 2003a Balsamo 1992 Hummon 1984b Guidi et al. 2003a Balsamo et al. 2010a Ferraguti & Balsamo 1995 Ferraguti et al. 1995 Ferraguti et al. 1995

Reference

a

Modified from Marotta et al. (2005) and amended. A question mark (?) indicates an unknown character state; N.A., not applicable; 0, absence; 1, presence. Fischer (1996) describes the penetrated spermatozoon of D. baltica. It is possible that mature ‟free” sperms do have a flagellum. b In several species, the centriole or basal body is strongly modified (see, e.g., Balsamo et al. 2002). c According to Marotta et al. (2005), D. baltica possesses a helical sperm head though it is straight according to Fischer (1996). d According to Marotta et al. (2005), D. baltica possesses mitochondria inside the nucleus though they lie along the nucleus according to Fischer (1996).

0 0 0 0 0 0 0 0 0 0 0 0 0

0 0 0 0 0 0 1 1 0 1 1 ? 0

Perinuclear helix Para-acrosomal bodies Basal crystal

31

Further structures

Cephalodasys maximus Lepidodasys sp. Lepidodasys unicarenatus Lepidodasys sp. (different species than in Guidi et al. 2004) Mesodasys laticaudatus Mesodasys adenotubulatus Acanthodasys aculeatus (see Fig. 1.38 A–F) Diplodasys ankeli Pseudostomella etrusca Tetranchyroderma sp. 1 Tetranchyroderma sp. 2 Tetranchyroderma papii Urodasys anorektoxys

Character-number (1–33) according to Marotta et al. (2005)

Tab. 1.5: (Continued)

1.2 Morphology 

 65

66 

 1 Gastrotricha

G H

aar

B ac

aa

A

sm

ncr

at

I

ab ncr

aar

C

ph mi

nu

J

fpr

D mi nu

sc

E

F

af

fpr

dc

L M

0.5 µm

Acanthodasys aculeatus (Figs. 1.38 A–F and 1.44 C–E) is a good representative of this “basic sperm plan”. It has to be stressed that this character pattern rather is a “generalized Bauplan” than a strictly phylogenetic reconstruction of the character pattern of the stem species of Macrodasyida. However, modifications from the aforementioned pattern are manifold. Microstructure and special differentiations of the acrosome, for example, are quite diverse. Acrosomal content may be condensed, uncondensed, or just condensed in the anterior or in the basal portion. Acrosomal tubular structures are absent in Lepidodasys, Mesodasys, Urodasys acanthostylis (present in U. anorektoxys), Turbanella, Paraturbanella, Dactylopodola, and Xenodasys (see Tab. 1.5 and references therein). Species of the Turbanellidae and Xenodasys eknomios (not Xenodasys sp.) possess piles of thick and electron-dense discs in their acrosomes. The chromatin

K se 0.5 µm

Fig. 1.38: Sperm ultrastructure (schematic) of Gastrotricha. (A–F) Spermatozoon of Acanthodasys aculeatus (Macrodasyida). (A) Whole sperm (131 µm total length). (B) Longitudinal sections of different regions. (C–F) Cross sections at different levels. (G–M) Spermatozoon of Musellifer delamarei (Paucitubulatina). (G) Whole sperm (28 µm total length). (H) Longitudinal sections of different regions. (I–M) Cross sections at different levels. Abbreviations: aa, anterior portion of acrosome; aar, anterior acrosomal region; ab, basal portion of acrosome; ac, acrosome; af, accessory fiber; at, axial tubular structure; dc, distal centriole; fpr, flagellar posterior region; mi, mitochondrion; ncr, nuclear central region; nu, nucleus; ph, perinuclear helix; sc, striated cylinder; se, septa; sm, supernumerary membrane. (A–F, According to Guidi et al. 2003a; G–M, according to Guidi et al. 2003b.)

is condensed in all species of the Macrodasyida investigated so far, unlike in species of Urodasys where sperm nuclei show a conspicuous fingerprint-like microstructure (Marotta et al. 2005, Balsamo et al. 2007). Many but not all of the studied species of Thaumastodermatidae (the only known exception is Pseudostomella etrusca) have a spring-shaped nucleus surrounded by a helically arranged cistern, the perinuclear helix (Ferraguti & Balsamo 1995, Guidi et  al. 2003b). Another unusual character of the nuclear central region was observed in species of Macrodasys: here, the giant mitochondrion winds spirally around the central and only slightly spiraled nucleus (Ruppert 1978a, Marotta et  al. 2005) and not vice versa like in most remaining macrodasyids. As already mentioned, there can be a single elongated mitochondrion associated with the nucleus or several small and ovoid ones (see Tab. 1.5 and the references

1.2 Morphology 

B

wf

C

aa

aar

mi cs tu

pb

D

A

B

ac pb

aa r

A

 67

ab

nu

E ncr

ncr

fpr

C

nu

1 µm pc

F

mi fpr

dc

Fig. 1.39: Sperm ultrastructure (schematic) of Gastrotricha. (A–F) Spermatozoon of Neodasys cirritus (Multitubulatina). (A) Whole sperm (approximately 13 µm total length). (B) Longitudinal sections of different regions. (C–F) Cross sections at different levels. Abbreviations: aa, anterior portion of acrosome; aar, anterior acrosomal region; ab, basal portion of acrosome; cs, crystalline structure; dc, distal centriole; fpr, flagellar posterior region; mi, mitochondrion; ncr, nuclear central region; nu, nucleus; pc, proximal centriole; tu, tubule of the acrosome; wf, “wig” of filaments. (A–F, According to Guidi et al. 2003b.)

in Tab. 1.5 are lost). In mature spermatozoa, mitochondria can also be absent at all as in species of Urodasys (Marotta et  al. 2005, Balsamo et  al. 2007). The striated cylinder surrounding the flagellar axoneme in the “basic sperm plan” of Macrodasyida is absent in several species, including all members of the Turbanellidae studied so far. A distal centriole (basal body) is reported from all studied species of Macrodasyida. However, shape and arrangement of basal structures vary a lot between species. There might be, for instance, accessory parts like conical or cap-like structures and thin fibers that support the connection between nucleus and axoneme (Tab. 1.5). Completely different sperm morphology can be observed in species of Dactylopodola (Fig. 1.41 A). Those sperms are filiform, too, but they

D

af 1 µm

Fig. 1.40: Sperm ultrastructure (schematic) of Gastrotricha. (A–D) Spermatozoon of Xenotrichula punctata (Paucitubulatina). (A) Whole sperm (approximately 10 µm total length without para-acrosomal bodies, length of para-acrosomal body is 20 µm). (B) Longitudinal sections of different regions. (C, D) Cross sections at different levels. Abbreviations: aar, anterior acrosomal region; ac, acrosome; af, accessory fiber; fpr, flagellar posterior region; mi, mitochondrion; ncr, nuclear central region; nu, nucleus; pb, paraacrosomal body. (A–D, According to Ferraguti et al. 1995.)

consist of a long, thin, and rod-shaped nuclear region and an adjacent rod-like compartment where small discshaped and piled mitochondria alternate with dense bodies of unknown function (Fischer 1996). Neither an acrosome nor a flagellum were observed, but it has to be mentioned that only penetrated spermatozoa of D. baltica were studied with TEM and no mature, testicular sperms. It is also possible that both structures disappear when the sperm cell enters the egg cell during fertilization (Fischer 1996). Further unusual sperm morphology was described for Dolichodasys carolinensis. Mature spermatozoa of that taxon appear like a “swollen semicolon” without a

68 

 1 Gastrotricha

A

B

C

D

mi nu db

E F 5 µm

1 µm

Fig. 1.41: Sperm ultrastructure (schematic) of Gastrotricha. (A) Longitudinal section of penetrated spermatozoon of Dactylopodola baltica (Macrodasyida). (B) Longitudinal section of sperm rod of Lepidodermella squammata (Paucitubulatina). (C–F) Different shapes of sperm cells in freshwater Paucitubulatina (all at same scale). (C) Filiform shape of Chaetonotus bisacer. (D) Rod-like shape of Chaetonotus sp. (E) Spindle-like shape of Polymerurus rhomboides. (F) Oval shape of Stylochaeta scirtetica. Abbreviations: db, dense bodies; mi, mitochondrion; nu, nucleus. (A, According to Fischer 1996; B, according to Hummon 1984b; C–F, according to Weiss 2001.)

flagellum (Ruppert & Shaw 1977). Unfortunately, spermatozoa of D. carolinensis were only studied with histological techniques and light microscopy. Subtle interspecific ultrastructural differences in spermatozoa are also quite diverse within the Chaetonotida (=Neodasys/Multitubulatina+Paucitubulatina). In addition, the gross sperm morphology, the “sperm types”, is more diverse among the major sub groups (Tab. 1.5). Accordingly, it was not possible to outline a “basic sperm model” for the Chaetonotida (Marotta et al. 2005). It has to be considered that Chaetonotida might not represent a monophylum (see chapter Phylogeny). In that case, a ground pattern reconstruction for such a group would be inappropriate. The tripartition into anterior acrosomal region, nuclear central region, and flagellar posterior region is also present in the sperm of Neodasys cirritus

(Fig. 1.44 A, B) and basal members of the Paucitubulatina such as Musellifer delamarei, Diuronotus aspetos (Guidi et  al. 2003b, Balsamo et  al. 2010a) and species of the Xenotrichulidae (Ferraguti et al. 1995). However, because neither the acrosomal nor the nuclear regions are spiraled or helically arranged, the external distinction between the three parts is less obvious. Neodasys is the only gastrotrich taxon that possesses the commaform (or dart-shaped, see Ruppert 1977) sperm type (Figs. 1.39 and 1.44 A, B). A somewhat conical to pear-shaped sperm head contains the anterior acrosome and the rod-shaped nucleus (Ruppert 1977, 1991, Guidi et al. 2003b, see also Kieneke et al. 2009). A short flagellum is attached perpendicular to the nuclear base, giving the whole cell the typical commaform shape (Fig. 1.39). Internally, the cone-shaped acrosome of Neodasys cirritus has a tubule (apical region) and a crystalline structure (basal region). Externally, it is enwrapped by a “wig” of short filaments in its apical region (Fig. 1.44 A), whereas few small mitochondria are situated near the transition from acrosome to nucleus (Guidi et al. 2003b). Until now, N. cirritus is the only known gastrotrich that has, in addition to the distal centriole (basal body), a remnant of the apical (accessory) centriole (Tab. 1.5). The not spiraled, filiform spermatozoa of basal marine and fully hermaphroditic members of the Paucitubulatina such as Musellifer delamarei (Fig. 1.38 G–M), Diuronotus aspetos, and species of the Xenotrichulidae (e.g., Fig. 1.40 A–D) are composed in sequence of an anterior acrosome, sometimes regionalized by different content or even separated into two parts (e.g., in Diuronotus aspetos), a rod-shaped and mostly incompletely condensed nucleus, a single or few mitochondria (e.g., in Musellifer delamarei, see Fig. 1.38 H, J) basal to the nucleus, and a long flagellum (Balsamo et al. 2010a, Ferraguti & Balsamo 1995, Ferraguti et  al. 1995, Guidi et  al. 2003b). The flagellum consists of a standard axoneme but with nine peculiar accessory fibers attached to the outer doublets of axonemal microtubules (see Figs. 1.38 K and 1.40 D). The three-dimensional microstructure of those accessory fibers is a complex system of undetermined filaments and rib-like structures (Ferraguti et  al. 1995). In cross section, each accessory fiber has a roughly triangular shape with a content of different electron transmissibility (see, e.g., Fig. 1.40 D). Analogue differentiations in the same position can be found in the flagellum of the spermatozoa of some macrodasyidan species, e.g., in Cephalodasys maximus (Fischer 1994) and in species of Urodasys (Balsamo et al. 2007). It has to be considered that these structures evolved convergently. Another instance of convergent evolution is the loss of mitochondria in sperms of Urodasys and derived members of Paucitubulatina (Marotta et al. 2005). Other differences between the sperm

1.2 Morphology 

cells of basal species of the Paucitubulatina exist in the shape and arrangement of the mitochondria. They might be quite big and barrel-shaped organelles like in Xenotrichula intermedia or X. punctata (Ferraguti et al. 1995, see Fig. 1.40 B), a large, cuff-like mitochondrion enwrapping the basal portion of the nucleus as in Diuronotus aspetos (Balsamo et al. 2010a), or, as already mentioned, a cluster of four small mitochondria like in Musellifer delamarei (Guidi et  al. 2003b, see Fig. 1.38 H). Peculiar differentiations of spermatozoa of some species of the Xenotrichulidae (e.g., Xenotrichula punctata, Fig. 1.40 A–D) are the para-acrosomal bodies, the function of which is still unknown (Ferraguti & Balsamo 1995, Ferraguti et al. 1995). Originating close to the transition between the real acrosome and nucleus, these long and thin structures (approximately 20×0.2 µm) are composed of piles of electron-dense discs (Fig. 1.40 A, B). Each disc is linked with its neighboring discs by thin filaments. The para-acrosomal bodies are extracellular formations and not surrounded by a cell membrane (Ferraguti et  al. 1995). It still remains unclear whether these presumably energy-costing para-acrosomal bodies are in any connection with the fertilization process (Ferraguti et al. 1995). More derived and mostly freshwater-dwelling species of the Paucitubulatina like Lepidodermella squamata or Chaetonotus maximus develop extremely modified spermatozoa after a phase of parthenogenetic reproduction (Hummon 1984b). These cells are nothing more but a rod-shaped nucleus with fully condensed chromatin surrounded by a cellular membrane (Hummon & Hummon 1983b, Hummon 1984b, Balsamo 1992). As nuclear envelope and cellular membrane lie close to each other, there is almost no cytoplasm present (Fig. 1.41 B). Comparably simplified spermatozoa as in L. squamata and C. maximus were first reported by a handful of researchers in cultured or environmentally sampled freshwater gastrotrichs (e.g., Weiss & Levi 1980, Kisielewska 1981, Balsamo & Todaro 1987, 1988). However, it was Weiss (2001) who demonstrated a widespread occurrence of simple, aflagellar sperm among 22 freshwater dwelling species of the Paucitubulatina. There is a certain degree of morphological diversity among these germ cells as they display filiform, rod-like, spindlelike, or oval shapes (Weiss 2001, Fig. 1.41 C–F). Because of their simplicity, these cells are frequently called vestigial spermatozoa (e.g., Marotta et al. 2005). However, though still not fully proven, an amphimictic function of these simplified sperms in a sexual phase of the life cycle after parthenogenesis is much likely (Balsamo 1992, Hummon 1984b, 1986, Weiss 2001, see also chapter Reproductive Biology). The occurrence of three morphologically different egg types in cultured and free-living freshwater

 69

gastrotrichs (subitaneous and resting eggs, both produced through apomictic parthenogenesis, and third morphologically different egg type, which maybe a product of amphimixis) is considered as possible evidence for sexual reproduction (Hummon & Hummon 1983a, Balsamo 1992 and references therein). Another sign for functionality of simplified sperm in freshwater Gastrotricha could be the observation that isolated postparthenogenetic individuals generally do not lay eggs, or if yet in rare cases, these eggs do not develop properly. This suggests that fertilization (amphimixis) is required for proper cleavage and embryonic development (Hummon 1984c, Balsamo & Todaro 1988, Balsamo 1992). Spermatozoa of the filiform and spiraled type can reach a considerable length, several dozen times longer than its diameter, that is mostly less than 1 µm. For instance, sperms of Acanthodasys aculeatus are approximately 131 µm long (Guidi et al. 2003b), of Macrodasys caudatus 75 µm (Marotta et  al. 2005), and of Megadasys sterreri 125–130 µm (Guidi et  al. 2014). However, there are also more compact filiform and spiraled sperm cells in species such as Urodasys acanthostylis (Balsamo et al. 2007, approximately 18 µm) or Turbanella ambronensis (Ferraguti & Balsamo 1995, no total length data provided). It seems that the filiform but not spiraled sperm cells of basal hermaphroditic species of the Paucitubulatina are generally less elongate. For instance, the mature spermatozoon of Musellifer delamarei has a length of 28 µm and a maximum width of almost 1 µm (Guidi et al. 2003a). Sperms of Xenotrichulidae, para-acrosomal bodies not considered, may be rather short with a length of 10 µm (X. punctata, Ferraguti et  al. 1995) or 14 µm (Heteroxenotrichula squamosa, Ferraguti & Balsamo 1995) but can be much longer as in X. intermedia (Ferraguti et  al., no total length data provided). The commaform sperms of Neodasys cirritus (ca. 13 µm length, Guidi et  al. 2003a), the aberrant sperms of Dactylopodola baltica (ca. 14 µm length, Fischer 1996), and the vestigial sperm cells of Lepidodermella squamata (7–8 µm length, Hummon 1984b), and other freshwater paucitubulatinans (Weiss 2001) are generally much shorter than those of most Macrodasyida (see Figs. 1.39 A–F and 1.41 A–F). Although motility of the flagellated sperm types of marine Gastrotricha is broadly assumed, there are, to our knowledge, only few documented observations of actively moving spermatozoa inside the testes, e.g., in an undescribed species of Neodasys (probably N. cirritus) when the specimen was slightly compressed (personal communication of Ruppert in Hummon & Hummon 1983b) and in different species of Xenotrichulidae (Ferraguti et al. 1995). If spermatozoa are artificially released

70 

 1 Gastrotricha

from the testes, they are usually inactive (Hummon & Hummon 1983b), but activity in freed spermatozoa was also observed, for instance, in Turbanella cornuta (Teuchert 1976b) or in Urodasys acanthostylis (Balsamo et al. 2007). There are, however, several reports of actively moving sperms inside the frontal organ of various taxa, for instance, in two species of Macrodasys (Ruppert 1978a), in two species of Crasiella (Guidi et  al. 2011), in Dinodasys mirabilis (Todaro et  al. 2012a), or in Neodasys chaetonotoideus (own unpublished observation). The configuration with one giant or several small mitochondria plus a well-developed axoneme also suggests activity in flagellated spermatozoa of the Gastrotricha. Whether sperm movement is already used during insemination or just to reach the mature egg via the internal pore of the frontal organ remains unclear. However, the rich muscular assembly of the caudal organ or parts of the sperm ducts, its absence in the frontal organ in many species and the numerous observations of sperm activity inside the frontal organ lumen (see Balsamo et al. 1999) might account for the latter possibility. However, the factor that activates sperm movement is still unknown (Balsamo et  al. 1999). Ruppert (1991) hypothesizes that the spiral shape of most spermatozoa of the Macrodasyida translates the beating action of the flagellum into a rotation of the whole sperm. This could facilitate movements of the sperm cell through the tissue or narrow ducts. As opposed to the flagellated sperm, the aberrant sperm cells without any flagellum in freshwater Paucitubulatina are definitely unable to perform active movements. The process how these immobile germ cells could be transferred from one specimen to the reproductive partner during cross fertilization, if happening at all, is still obscure (e.g., Balsamo et  al. 1999, Hummon 1986, Weiss 2001). As already mentioned, some Macrodasyida, e.g., species of Dolichodasys, and perhaps species of Dactylopodola, possess non-flagellated sperm, too (Fischer 1996, Ruppert & Shaw 1977). However, those animals exhibit, like Dolichodasys carolinensis (Ruppert & Shaw 1977), a muscularized caudal organ that is probably used for transferring the immobile sperm cells to the mating partner. In Dactylopodola baltica and D. typhle, male gametes are exchanged via spermatophores (Kieneke et al. 2008d, Teuchert 1968).

1.2.10.2 Spermatogenesis and spermiogenesis The formation and differentiation of male germ cells generally involve two consecutive developmental processes, i.e., (1) spermatogenesis and (2) spermiogenesis (=spermatohistogenesis) (see Gilbert & Singer 2006).

During spermatogenesis, a spermatogonion (diploid) matures into a primary spermatocyte I (still diploid). This cell undergoes the first meiotic division and becomes two secondary spermatocytes II (haploid but sister chromatids still united) and the second meiotic division leads to four spermatids (haploid and sister chromatids separated). Spermiogenesis is the cytomorphological process that matures spermatids into ripe spermatozoa through severe cellular transformations. Early reports of sperm cell formation within Gastrotricha based on light microscopic observations were made by Remane (1936) for Oregodasys (former Platydasys), Ruppert & Shaw (1977) for Dolichodasys ­carolinensis, or Ruppert (1978b) for several species of the Thaumastodermatidae. In detail and based on TEM studies, spermatogenesis has only been studied in the marine macrodasyidan species Turbanella cornuta (Teuchert 1976b, 1977a) and in the freshwater-dwelling paucitubulatinan Lepidodermella squamata (Hummon 1984b). Some more general data concerning spermatogenesis is meantime available for additional species, e.g., for Acanthodasys aculeatus, Lepidodasys sp., Dinodasys mirabilis, Crasiella diplura, or Urodasys anorektoxys (Balsamo et al. 2007, Guidi et al. 2003b, 2004, 2011, Todaro et  al. 2012a). Furthermore, there is some fragmentary data of spermatogenesis in Dactylopodola typhle and in Neodasys chaetonotoideus (Kieneke et al. 2008d, 2009). In T. cornuta, presumably most species of the Macrodasyida, spermatogenesis proceeds in the germinal epithelium caudofrontally, i.e., from the distal to the proximal pole of the testis (Fig. 1.42 A). Such a caudocephalic maturation of male germ cells has impressively been demonstrated for Lepidodasys sp. (see image 3B in Guidi et al. 2004) and is assumed to represent the ancestral mode of spermatogenesis (see chapter Reproductive Organs). However, spermatogenesis may also proceed in the opposite direction as in Urodasys anorektoxys (Balsamo et al. 2007). Since spermatogonia and primary spermatocytes are difficult to distinguish even at the ultrastructural level, Teuchert (1976b) simply described spermatogonial stages A–D of Turbanella cornuta, whereas stage A probably represents spermatogonia and stage D the early spermatids before spermiogenesis starts. In T. cornuta and other species of the Macrodasyida (see, e.g., Kieneke et  al. 2008d for Dactylopodola typhle; Fig. 1.37 E), the germinal epithelium consists of two or more layers of spermatogonial cells. Stage A cells in T. cornuta are characterized by big and active nuclei (regularly distributed euchromatin) and numerous free ribosomes. In stage B cells, electron-dense vesicles of 0.3 µm diameter and unknown origin and function appear. Stage C cells

1.2 Morphology 

A

frontal cc

spermiogenesis

gp

spermatogenesis

tl ge vd

tl caudal

B

Fig. 1.42: Spermatogenesis and spermiogenesis of Gastrotricha. (A) Schematic horizontal section of testes in Turbanella cornuta (Macrodasyida) indicating direction of spermatogenesis (gray arrow) and spermiogenesis (black arrows) (B) Diagrammatic spatial arrangement of the 16 spermatids (gray ovals) within a spermatogenic cyst of Lepidodermella squammata (Paucitubulatina). Observed (black lines) and assumed (broken lines) cytoplasmatic bridges between spermatids are indicated. Abbreviations: cc, cap cells of unknown function; ge, germinal epithelium; gp, male genital pore; tl, testicular lumen; vd, vas deferens. (A, Original according to data in Teuchert 1976b; B, according to Hummon 1984b.)

 71

are characterized by patterns of maturing divisions (condensed chromosomes, absence of nuclear envelop) and cytoplasmic bridges between adjacent cells. Stage D cells, spermatids, are characterized by rounded nuclei with central condensed and reticulate chromatin, peripherally electron-lucent nucleoplasm, voluminous and active Golgi apparatus, numerous mitochondria, and beginning formation of the flagellum (Teuchert 1976b). Investigations of further species support this scenario (see Balsamo et  al. 2007, Guidi et al. 2003b, 2004, Todaro et al. 2012a). Although only fragmentary data are available, spermatogenesis in Dactylopodola typhle seems to be different than in other macrodasyidan species. Dozens of spermatids are combined in a common cyst (Kieneke et al. 2008d). This is far more than a number of four spermatids that would result from meiosis of one primary spermatocyte. As opposed to spermatogenesis, spermiogenesis has been studied thoroughly in several species of the Macrodasyida: Turbanella cornuta (Teuchert 1975b, 1976b, Fig. 1.43 A–H), Cephalodasys maximus (Fischer 1994), Acanthodasys aculeatus (Guidi et  al. 2003b), Lepidodasys sp. (Guidi et al. 2004), Urodasys anorektoxys (Balsamo et al. 2007), Crasiella diplura (Guidi et  al. 2011), and Dinodasys mirabilis (Todaro et  al. 2012a). Spermiogenesis is a complex cytomorphological process and can involve up to five steps as in the species Cephalodasys maximus (Fischer 1994). Spermiogenesis usually proceeds from the periphery of the testis toward the lumen (centripetal direction, Guidi et  al. 2011), where mature spermatozoa are densely stocked (Fig. 1.42 A). Generalized and simplified, spermiogenesis in macrodasyidan gastrotrichs takes place as simultaneous acrosome (pro-acrosome) formation and growth of the flagellum. Both structures protrude parallel into the testicular lumen, hence giving the developing spermatid a characteristic U-shape (Figs. 1.42 A and 1.43 C). This was hypothesized as a common feature, an autapomorphy of the Macrodasyida (Balsamo et  al. 2007). During early (pro-)acrosome and flagellum formation, the nucleus also starts to grow out and follow the elongating and more and more twisting acrosome (Fig. 1.43 A, C). Several small mitochondria may follow the elongating nucleus and later fuse to form a single giant mitochondrion enclosed by the nuclear helix as in Turbanella cornuta (Teuchert 1975b, 1976b, Fig. 1.43 A, C, E) or Acanthodasys aculeatus (Guidi et  al. 2003b). However, it is also possible that a single mitochondrion at the base of the elongating nucleus starts to project into it like in Cephalodasys maximus, further mitochondria from the spermatid may later fuse with the initial one (Fischer 1994). Spiralization of the acrosome-nuclear complex

 1 Gastrotricha of three cells represent the spermatids of L. squamata. Each spermatid already possesses an electron-dense, rod-shaped nucleus, and the cytoplasm and organelles begin to reduce (inactive Golgi complexes, only a thin

A

E

np er

fv

B

F

sb ad

mi

nu

w

ad

mt

aar

nu

cl

pa nu 1 µm

go

C

G

cs

mi

ncr

generally seems to begin at the apex of the spermatid appendage that contains pro-acrosome and elongating nucleus. During the profound cytomorphological transformations of spermiogenesis such as elongation, shifting of organelles and spiralization of the acrosome-nuclear complex, microtubules seem to play an important role. In stage C of spermiogenesis of Cephalodasys maximus, for instance, the acrosome-nuclear complex is enclosed by a cuff of circular (helically?) arranged microtubules (Fischer 1994). Helically arranged microtubules encircle nucleus and mitochondrion during spermiogenesis of Turbanella cornuta but are absent in mature spermatozoa (Teuchert 1975b, 1976b, Fig. 1.43 B, D, G). Microtubules also occur during formation of the nucleus-mitochondrion complex in Crasiella diplura (Guidi et al. 2011). Comparably, microtubules surround the rod-shaped nucleus in spermatids of Lepidodermella squamata at the beginning of spermiogenesis but disappear later (Hummon 1984b, see below). The last step of spermiogenesis in macrodasyidan Gastrotricha is characterized by severe reduction of cytoplasm of the late spermatid. Because lysosomes and peroxisomes are frequently observed during that stage, it is hypothesized that reduction of cytoplasm happens by autolysis and reabsorption (Guidi et  al. 2003b). In species such as Urodasys anorektoxys or Dinodasys mirabilis, it is also possible that residual bodies are formed from the debris of the late spermatid (Balsamo et  al. 2007, Todaro et  al. 2012a). An interesting mode of metabolic recycling of those residual bodies is reported from U. anorektoxys. In that species, several macrophages per testis remove the residual bodies (Balsamo et al. 2007). Sperm cell formation, i.e., spermatogenesis and spermiogenesis, was intensely studied by Hummon (1984b) in the freshwater paucitubulatinan species Lepidodermella squamata. She distinguished four stages of sperm rod formation, spermatogenesis (until early spermatids) happens during stages 1 and 2 and spermiogenesis (until mature sperm rods) is covered by stages 2, 3, and 4. Four spermatogonia are connected by cytoplasmic bridges and characterized by fibrillar nuclear content; two developing cyst cells may already be present at that stage. Using the resolving power of the TEM, Hummon (1984b) was able to identify possible precursor cells of the four spermatogonia in specimens with temporally uneven development at both sides of the body. In later specimens, primary spermatocytes could unequivocally be identified through the presence of synaptonemal complexes that are diagnostic for pachytene of first maturing division of meiosis (Fig. 1.37 F). Afterward, a cluster of 16 intertwined cells connected by cytoplasmic bridges (Fig. 1.42 B) and enclosed within a cyst made

np nu dc nu

er

mi

D H

mi

mt

cl ci fpr

72 

ci 1 µm

0.5 µm

Fig. 1.43: Ultrastructure (schematic) of spermiogenesis in Turbanella cornuta (Macrodasyida). Gray arrows indicate direction of development. (A and B) Spermatid in an early stage of spermiogenesis. Nucleus and acrosome start to elongate and numerous small mitochondria are scattered along nucleus. (C and D) Spermatid in a late stage of spermiogenesis. Cilium (flagellum) has grown into testicular lumen and nucleus and giant mitocjondrion are already spiraled. Note the microtubules surrounding the mitochondrion-nuclear complex in B and D. (E–H) Mature spermatozoon. (A, C, and E) Longitudinal sections. (B, D, and F–H) Cross sections. Abbreviations: aar, anterior acrosomal region; ad, acrosomal discs (alternating light and dense discs); ci, cilium (flagellum); cl, clasp-like structure; cs, connecting structure between basal body and flagellum; dc, distal centriole (basal body); er, endoplasmic reticulum; fpr, flagellar distal region; fv, flagellar vacuole; go, Golgi cisterns; mi, mitochondrion; mt, microtubules; ncr, nuclear central region; np, nuclear plasm; nu, nucleus; pa, pro-acrosome; sb, spiraled band; w, “wall” of acrosome. (Modified from Teuchert 1976.)

1.2 Morphology 

layer of cytoplasm around one pole of nuclear rods), while mitochondria and multivesicular bodies are still numerous in some regions (Hummon 1984b). The rod shaped nucleus of late spermatids is surrounded by up to 10 microtubules, the cytoplasmic bridges between

 73

spermatids follow a regular pattern (see Fig. 1.42 B). In slightly older individuals, spermiogenesis has finished and 16 rod-shaped, 7- to 8-µm-long sperm cells, just consisting of rod-shaped condensed nucleus with surrounding cell membrane and one large residual body

B

A

wi

nu wi

ac

ac

nu

nu pc

C

*

2 µm

D

E ph

F

1 µm

G

cs

ac

nu mi

nu mi

*

2 µm

sc

1 µm

1 µm

1 µm

2 µm

Fig. 1.44: Ultrastructure (TEM micrographs) of the spermatozoa of Gastrotricha. (A) Neodasys ciritus (Multitubulatina). Longitudinal and cross sections of some spermatozoa within the testis. (B) Longitudinal and cross sections of developing sperm cells within the testis of N. chaetonotoideus. Note the different ultrastructure of the acrosome compared to N. ciritus. (C–E) Longitudinal sections of spermatozoa of Acanthodasys aculeatus (Macrodasyida). (C) Acrosomal region. (D) Nuclear region. (E) Flagellar region (cilium). (F and G) Longitudinal sections through 1 testis of Paraturbanella teissieri (Macrodasyida). (F) Overview with several longitudinally sectioned spermatozoa. (G) Close-up of some sperm cells showing the helically arranged nuclear-mitochondrial complex, the acrosome, and the twisted flagellum. Note the electron-dense structure that connects the flagellum with the nucleus (asterisk). Abbreviations: ac, acrosome; cs, clasp-like structure; fl, flagellum (cilium); mi, mitochondrion; nu, nucleus; pc, proximal centriole; ph, perinuclear helix; sc, striated cylinder; wi, wig of filaments. (Micrographs in A and C–G were kindly provided by Maria Balsamo and Loretta Guidi, Urbino.)

74 

 1 Gastrotricha

are placed within a cyst made of three cells. The residual body, up to 5 µm in diameter, contains presumptive cell debris (e.g., multivesicular bodies, various membranes, lysosomes) and disappears later. The whole process of sperm formation (spermatogenesis plus spermiogenesis) starts after deposition of the last parthenogenetic egg and lasts only 1 day (Hummon 1984b). Almost nothing is known about spermatogenesis and spermiogenesis in more basal, hermaphroditic taxa of the Paucitubulatina such as the Xenotrichulidae. In juvenile specimens, there are no traces of developing sperm cells but they are suddenly present in mature animals (Hummon & Hummon 1983b). This may point to a rather rapid sperm development in those groups. A process that might precede spermatogenesis is mitotic proliferation of spermatogonia by ongoing cell divisions within the germinal epithelium. Such a process within Gastrotricha has only been reported for Oregodasys cf. ocellatus (Ruppert 1978b, Balsamo et al. 1999). However, in Turbanella cornuta and Dactylopodola baltica a repeated alteration of male and female phases during a life span is described (Teuchert 1968). It is possible that mitotic proliferation at least of the male germ cells is involved in such an alternating and “phase-delayed” hermaphroditic reproduction mode (Hummon & Hummon 1983b, Balsamo et al. 1999). If mitotic proliferation within the testes of hermaphroditic gastrotrichs is a common process, long time assumed cell constancy and eutely of Gastrotricha must be restricted to the somatic cell line only (Balsamo et al. 1999). However, as regeneration capacity was demonstrated in Turbanella sp., which must involve mitotic activity of somatic cells (Manylov 1995), the idea of eutely in Gastrotricha, at least in Macrodasyida, has to be questioned in general.

outer layer consists of fibrous material, too, but without apparent orientation and with flocculent material dispersed among fibers (Rieger & Rieger 1980). Comparably, in Dactylopodola baltica, the inner layer of the mature egg envelope is fibrous and the outer layer consists of electron-dark filaments. However, these are covered by a membrane-like structure (Fischer 1996). Microscopically, the eggshell surface of species of Macrodasyida appears smooth and is uniformly sticky (Rieger & Rieger 1980, Ruppert 1991). It may additionally be provided with a thick covering of glutinous secretion, which may be produced by secretory cells of the epidermis or the uterus wall (Teuchert 1968) or by the recently described posterior gland organ in Dinodasys mirabilis (Todaro et al. 2012a). This adhesive covering enables the animals to stick the spawned eggs to the substratum such as sand grains (Hummon & Hummon 1983a). In species of the Paucitubulatina, the ultrastructural composition of the inner layer of the eggshell is fibrous, too. The outer eggshell layer displays a more homogenous and electron-dense texture (Rieger & Rieger 1980). The inner eggshell stratum of eggs of Lepidodermella squamata is described as a continuous layer of poorly stained material with a narrow dense outer layer that forms numerous dense caps (Hummon 1984a). It is the outer layer of the eggs of paucitubulatinans that may exhibit eggshell sculpturing of various shapes (e.g., Figs. 1.45 A–C and 1.46 C–F) such as tiny spinelets, humps, spines, or pillars, but eggshells may also be smooth (Remane 1936, Hummon & Hummon 1983a, Ruppert 1991). Furthermore, taxa like Aspidiophorus, maybe most paucitubulatinans possess a sticky attachment stalk at their eggs (Rieger & Rieger 1980). As those eggs are not uniformly sticky as the eggs of Macrodasyida, the attachment stalk is used to adhere the spawned egg to the

1.2.10.3 Eggs Mature eggs of Gastrotricha may be small spheres with a diameter of 35–45 µm but are predominantly ovals of up to 60×120 µm (Teuchert 1968, Rieger & Rieger 1980) (Fig. 1.36 A, D, E). Mean dimension for most taxa, however, is an oval of 40×60 µm (Hummon & Hummon 1983a). Both Macrodasyida and Chaetonotida have eggs with a welldifferentiated eggshell being stratified into two different layers (Rieger & Rieger 1980, Hummon 1984a, Ruppert 1991). In species of the Macrodasyida, both layers consist of fibrous material, whereas the innermost maybe organized into crossed fibers such as in Turbanella ocellata (Rieger & Rieger 1980) or in T. cornuta (Ruppert 1991). The

A

B

C

Fig. 1.45: Different egg shapes of freshwater Paucitubulatina. Drawings not in scale, diameter of eggs vary between 50 and 75 µm. (A) Egg of Chaetonotus maximus. (B, C) Eggs of 2 other species of Chaetonotus. (A–C, According to Remane 1936.)

1.2 Morphology 

A

 75

B C

ph

40 µm

ph

5 µm

E

35 µm

D

80 µm

F

10 µm

substratum (Ruppert 1991). The eggs of the Paucitubulatina show even more structural diversity: during their parthenogenetic phase, paucitubulatinan gastrotrichs like Lepidodermella squamata (Hummon 1984a) produce subitaneous eggs (so-called tachyblastic or quick developing eggs according to Brunson 1949; Fig. 1.46 A–C) and resting eggs (so-called opsiblastic or late developing eggs according to Brunson 1949). Both egg types may be structurally different within the same species, resting eggs being generally more heavily sculptured and with a darker egg shell (Hummon & Hummon 1983a). It is worth mentioning that resting eggs are of high ecological and biogeographical value because they facilitate the species to outlast periods of unfavorable environmental conditions like dry or cold seasons (Strayer & Hummon 1991). Furthermore, resting eggs may represent important indirect dispersal propagules (Strayer & Hummon 1991, Artois et  al. 2011). A third morphological egg type, the “plaque-bearing egg” (see Fig. 1.49), is reported in Lepidodermella squamata that possibly represents a fertilized egg (Levy & Weiss 1980, see also Hummon & Hummon 1983a, Strayer & Hummon 1991, Balsamo 1992). Apart from some information on oogenesis and vitellogenesis of Neodasys chaetonotoideus (Kieneke et al. 2009), there does exist no more data on the ultrastructure of mature eggs and the eggshell in Neodasys/Multitubulatina.

5 µm

Fig. 1.46: Egg types of Paucitubulatina. (A and B) Aspidiophorus polystictos. (A) Subitaneous egg (tachyblastic egg) with an embryo inside that has finished embryogenesis. (B) Surface of the egg shell with a fine granulation. (C–F) Chaetonotus sp. (C) Subitaneous egg with an embryo inside. (D) Surface differentiation of a subitaneous egg with pillar-like structures. (E) Close-up of the egg shell. (F) Pillar-like structures of a resting egg (opsiblastic egg). (A, C) DIC images. (B and D–F) SEM micrographs. Abbreviation: ph, pharynx of the embryo. (All images were kindly provided by Maria Balsamo & Loretta Guidi, Urbino.)

1.2.10.4 Oogenesis It has been discussed controversially if the ovaries of Gastrotricha are real ovaries because according to Hummon & Hummon (1983a) real ovaries are organs were primordial germ cells (oogonia) produce oocytes by mitotic proliferation (see also Ruppert 1991 for this issue). There are different reports of cell division patterns within the proximal part of the female gonads in several macrodasyidan gastrotrich species (e.g., Rieger et  al. 1974: Chordodasiopsis antennatus, Ruppert & Shaw 1977: Dolichodasys carolinensis, Ruppert 1978a: Macrodasys sp., Ruppert 1978b: Oregodasys cf. ocellatus and Acanthodasys thrinax). However, it is not clear whether these patterns represent mitotic or meiotic activities (see Hummon & Hummon 1983a, Ruppert 1991). The only certain instance of mitotic activity is known in the ovary of post-parthenogenic specimens of the freshwater paucitubulatinan species Lepidodermella squamata (Hummon 1984c). Apart from the aforementioned issue, oogenesis was ultrastructurally studied in different species of the Gastrotricha, most comprehensively in the marine hermaphrodites Turbanella cornuta (Teuchert 1977a) and Dactylopodola baltica (Fischer 1996) and in the parthenogenetic freshwater species Lepidodermella squamata (Hummon 1984a). Further data concerning oogenesis are

76 

 1 Gastrotricha

A cu mg

ov

* tes

mo

mo

nu tes

5 µm

B

tes

cu

C

ed

mo * mo

er tes ed lm

2 µm

5 µm

available for the macrodasyidan species Dactylopodola typhle (Kieneke et  al. 2008d), Crasiella diplura (Guidi et al. 2011), Dinodasys mirabilis (Todaro et al. 2012a), and for Neodasys chaetonotoideus (Kieneke et al. 2009). Early female germ cell development in hermaphroditic gastrotrich species definitely proceeds by meiosis because the occurrence of condensed chromosomes and synaptonemal complexes, diagnostic for synapsis during leptotene or zygotene of meiosis I, is frequently observed (e.g., Teuchert 1977a: Turbanella cornuta, Fischer 1996: Dactylopodola baltica). In the freshwater-dwelling gastrotrich Lepidodermella squamata, oogenesis leads to parthenogenetic eggs. Because patterns of meiosis (neither condensed chromosomes nor synaptonemal complexes) were never discovered in parthenogenetic individuals during intensive ultrastructural studies using TEM, it is hypothesized that diploid parthonegenetic eggs are produced by apomixis in L. squamata (Hummon 1984a, Ruppert 1991, Balsamo 1992). Generalized, egg development in Gastrotricha involves three major successive stages prior to spawning: growth, vitellogenesis, and egg shell formation. Young

mg

Fig. 1.47: Ultrastructure (TEM micrographs) of the oocytes of Gastrotricha. (A) Cross section through the trunk of Xenotrichula carolinensis (Paucitubulatina). Paired testes and ovaries in a lateral position. Maturing eggs lie in close proximity to the midgut. The large egg on the right side is still growing while that on the left already has different types of granules and vesicles. Note the rather big nucleus and nucleolus (asterisk) of the growing egg. (B) Cross section through the trunk of Diuronotus aspetos with a mature egg and spermatozoa within the testis. Note the different types of granules within the egg. (C) Cross section of the uterus of Neodasys chaetonotoideus (Multitubulatina). The maturing egg lies in close contact to gut cells. Vitellogenesis has initiated as can be seen by the numerous electron-dark granules. Note the active nucleus with prominent nucleolus (asterisk). Abbreviations: cu, cuticle; ed, epidermis; er, endoplasmic reticulum; lm, longitudinal muscle; mg, midgut; mo, mature/maturing oocyte; nu, nucleus; ov, ovary; tes, testes. (Micrograph B was kindly provided by Maria Balsamo and Loretta Guidi; Urbino; micrograph C, from Kieneke et al. 2009, with kind permission by Wiley.)

oocytes have a rather big and active nucleus with regularly scattered heterochromatin, almost no condensed chromatin and a distinct nucleolus (Fig. 1.47 A). Within the cytoplasm, mitochondria, free ribosomes, and rough endoplasmic reticulum are present as in Lepidodermella squamata, Turbanella cornuta, or Dactylopodola baltica (Teuchert 1977a, Hummon 1984a, Fischer 1996). Besides mitochondria and electron-lucent vesicles, Kieneke et  al. (2008d) report a high quantity of small electron-dense granules in early oocytes of Dactylopodola typhle, supposedly glycogen. In early stages of oogenesis, the centriole pair may be present like in the young egg cells of Lepidodermella squamata or Dactylopodola baltica (Hummon 1984a, Fischer 1996). Next, developing oocytes continuously migrate toward the distal uterus region. The direction of this migration may be directed frontally or caudally, depending on the anatomy of the female gonads (see chapter Reproductive Organs). In species with paired ovaries, maturing eggs alternately originate from the left and the right gonad such as in Lepidodermella squamata and Dactylopodola baltica (Hummon 1984a, Fischer 1996). During

1.2 Morphology 

this phase, the eggs grow considerably, for instance, six to eight times their original size like in T. cornuta (Teuchert 1977a). In Lepidodermella squamata, the egg cytoplasm grows several thousands of times, whereas its nuclear volume still enlarges by a factor of several hundreds (Hummon 1984a). Gastrotricha have a remarkably high ratio of egg and whole animal biomass, which means that they invest considerably high energy and matter into offspring. In gravid specimens of many species of the Paucitubulatina, for instance, length of the mature egg may encounter more than half of the whole animal. With such a size ratio, eggs of Paucitubulatina display the comparatively biggest eggs among Metazoa (Remane 1936). During the growing phase of the premature eggs, vitellogenesis begins. Mitochondria, Golgi complexes, and free ribosomes are very abundant within the cytoplasm during vitellogenesis, and the nuclear envelope may display distinct nuclear pores (Teuchert 1977a, Hummon 1984a). Furthermore, different inclusions of varying sizes (ranging around 0.5 µm), textures, and electron-transmissibility become visible within the cytoplasm (Fig. 1.47 B, C). Those structures are interpreted as lipid droplets and yolk granules (e.g., Teuchert 1977a, Ruppert 1978b, Rieger & Rieger 1980, Hummon 1984a, Fischer 1996, Kieneke et  al. 2008d, 2009, Guidi et  al. 2011). Fully mature eggs are densely packed with those rest substances. In most gastrotrich species, vitellogenesis is brought about by autosynthesis of the egg cell itself, as no nurse cells, follicle cells, or yolk contributing cells have been observed (Hummon & Hummon 1983a). However, a possible exception to this is reported later. Molecular components for yolk and other rest substance production must be transported from the gut cells to the oocytes via the intercellular space like in Lepidodermella squamata (Hummon & Hummon 1983a) or via direct cellular connections of gut cells with the oocyte like in Turbanella ocellata (Rieger & Rieger 1980). The latter mode of nutrient supply for the developing eggs was already assumed by Wilke (1954). In L. squamata, where no ovary wall epithelium is present, the egg plasma membrane facing the gut epithelium forms deep invaginations into the egg cytoplasm. These so-called vitellogenic channels facilitate a rapid transcytotic transport of substances from the gut cells via the intercellular space deep into the maturing oocytes (Hummon & Hummon 1983a). Another evidence of direct nutrient supply of the developing eggs by gut cells comes from TEM investigations of Neodasys chaetonotoideus. Early, pre-vitellogenic eggs lie in direct contact with the gut cells, only separated by a narrow intercellular cleft filled with ECM. The egg itself forms

 77

lateral extensions that enwrap the gut thereby enlarging the contact surface (Kieneke et al. 2009). This obviously facilitates a better exchange of substances. However, this peculiar species also offers a second mode of vitellogenesis. More distally in the ovary, a wall epithelium completely surrounds the developing egg. In this region of the so-called vitellogenic oviduct, the epithelial cells form protuberances that deeply invaginate into the oocyte and also constrict small spheres that may later become the yolk granules of the mature egg (Kieneke et  al. 2009). A comparable situation may be present in further species of the Gastrotricha. For instance, cross sections of Thaumastoderma sp. demonstrated a complete epithelial lining around the developing oocyte, which forms an invagination into the ventral side of the egg (Ruppert 1978b, Hummon & Hummon 1983a). Also, Teuchert (1977a) reports “thin processes of the ovarian wall epithelium that stretch in between single oocytes” in Turbanella cornuta. Furthermore, Remane (1934, 1936) describes nutritive cells in close adjacency to the ovaries of Paradasys subterraneus. If all the aforementioned structures and formations are actually involved in vitellogenesis and nutrient supply of the growing eggs needs further exploration. The final stage of oogenesis in Gastrotricha is the formation of the egg shell. Eggshell formation still takes place inside the gravid animal and was intensely investigated in two marine species (Turbanella ocellata, Macrodasyida, and Aspidiophorus sp., Paucitubulatina) by Rieger & Rieger (1980) and in the freshwater-dwelling paucitubulatinan species Lepidodermella squamata (Hummon 1984a). The formation of the eggshell in Gastrotricha occurs in late-vitellogenic eggs and is possibly initiated by sperm penetration in hermaphroditic species (Rieger & Rieger 1980). Eggshell precursor vesicles occur inside the vitellogenic oocyte (Rieger & Rieger 1980, Hummon & Hummon 1983a), in later stages in close proximity to the cellular membrane (Hummon 1984b). There is evidence that eggshell vesicles are produced and processed by the rough endoplasmic reticulum and the Golgi complexes of the oocyte. As the content of vesicles is quite comparable with the finished eggshell regarding its texture or even shape (small spinelets inside the vesicles that resemble the spinelets of the eggshell), it is assumed that they are transported to the surface and fuse with the cellular membrane of the egg. Thereby, content of the eggshell precursor vesicles is deposited to the exterior (Hummon & Hummon 1983a). During spawning, eggs of Gastrotricha from both major subtaxa (Macrodasyida and Paucitubulatina) are reported to be highly flexible and virtually

78 

 1 Gastrotricha

“flow out” when passing the female gonopore or ruptured body wall (Teuchert 1968, Hummon & Hummon 1983a). This implies that also the eggshell must be quite flexible because it is already formed within the gravid animal (see above). A hardening of the eggshell begins immediately after spawning when the egg is deposited to the substratum. Hardening continues throughout embryonic development (Hummon & Hummon 1989). The hardening process might possibly be induced once the eggshell gets into contact with the surrounding medium, i.e., seawater, brackish water, or freshwater. Although each of the paired female gonads of the freshwater inhabiting Lepidodermella squamata (Paucitubulatina) contains eight oocytes, only four, rarely five or six parthenogenetic eggs are matured and spawned (Levy & Weiss 1980, Hummon & Hummon 1983a, 1993). Most frequently, the last parthenogenetic egg is a resting (opsiblastic) egg (Hummon & Hummon 1983a, Hummon 1984a, see also chapter Reproductive Biology). In a marine species of Chaetonotidae, Aspidiophorus polystictos, usually three to six parthenogentic eggs are produced (up to 10) before the animals become hermaphrodites (Balsamo & Todaro 1987, Hummon & Hummon 1993). Parthenogenetic resting eggs generally start to develop after a certain period of dormancy. However, it is not known which factor exactly triggers the production of and hatching from resting eggs in Gastrotricha (Hummon & Hummon 1983a, Balsamo 1992). In post-parthenogenic specimens of L. squamata, the remaining cells of the female gonad undergo limited mitotic proliferation. Although some of the resulting secondary oocytes develop into secretory cells that later fuse to a secondary syncytium and build up the so-called X-body of unknown function, others may develop into eggs that are finally spawned. However, those secondary eggs do not undergo proper cleavage in laboratory cultures (Hummon 1984c). Regarding the Macrodasyida, there is no knowledge how many eggs are produced and laid during a lifetime of any species (Hummon & Hummon 1983a). An indirect estimate for this could be the observed total number of oocytes present inside the ovary. For instance, Fischer (1996) depicts 12 oocytes per lateral gonad of Dactylopodola baltica, in D. typhle, approximately 15 germ cells per female gonad are reported (Kieneke et al. 2008d), and in Crasiella diplura only four to five cells (Guidi et al. 2011). However, it is still under debate whether mitotic proliferation of oocytes occurs in the gonads of hermaphroditic gastrotrichs (see beginning of this section). If true, this could possibly mean that numeric egg production is much higher than the aforementioned numbers. Such a scenario may also match the expected lifetime of macrodasyidan gastrotrichs, assumed to last between 6 and 12  months

(Artois et al. 2011). However, exact life expectancy of specimens in wild populations has never been determined so far (Hummon & Hummon 1992). What is definitely known in some species is the number of laid eggs at a time. This ranges between a single egg as in Lepidodermella squamata and species of Cephalodasys and Mesodasys, occasionally two as in Turbanella, Dactylopodola, and Urodasys, and up to eight eggs laid at a time like in species of Macrodasys (Teuchert 1968, Hummon & Hummon 1983a).

1.3 Reproduction and development 1.3.1 Reproductive biology We are at an initial stage of understanding the reproductive biology of the predominantly hermaphroditic gastrotrich taxa Macrodasyida and Neodasys. One phenomenon related with hermaphroditism in both taxa is the chronology of maturation of both sexes in the same individual. Generally, macrodasyidan gastrotrichs may be protandrous or simultaneous hermaphrodites or may display an alteration of sexual phases (Balsamo 1992). Teuchert (1968) already identified different chronologies of sexual maturation: species such as Macrodasys caudatus, Urodasys mirabilis, Mesodasys laticaudatus, Acanthodasys aculeatus, and Cephalodasys maximus are true simultaneous hermaphrodites with only a slightly earlier development of the male gonads. The mode of simultaneous hermaphroditism is also present in Neodasys chaetonotoideus (Kieneke et  al. 2009). Urodasys cornustylis and U. spirostylis are clearly protandrous hermaphrodites that show a much earlier maturation of the testes and the male caudal organ before maturing oocytes are visible and before the animals have been grown up to their full length (Schoepfer-Sterrer 1974, Hummon & Hummon 1992). Meanwhile, species such as Dactylopodola baltica and different species of Turbanella are reported to be protandrous species with a following multiple alteration of each sex (Remane 1936, Teuchert 1968, Hummon & Hummon 1992). In Turbanella, both sexes may temporally overlap in a single individual while D. baltica either is in a male or in a female stage. Both examples of protandry with following alteration of sexes, termed “sequential hermaphroditism”, according to Hummon & Hummon (1992), furthermore indicate a certain temporal synchrony in reproductive activity within the same population (Teuchert 1968). The possible presence of an environmental factor for the control of such a synchrony in wild populations is discussed by Hummon



& Hummon (1992). However, other species of the same genus may display a different chronology of sexual maturity. For instance, Dactylopodola typhle might be a simultaneous hermaphrodite (Kieneke et al. 2008d) like the aforementioned species of Macrodasyida in contrast to its congeneric D. baltica. Simultaneous hermaphroditism in D. typhle was already suspected by Wilke (1954), who observed testes with spermatids in specimens of the assumed female stage. It has to be stressed that the alteration of sexes was purely deduced from the occurrence of different size classes in Turbanella during the season that have been either specimens in a more male or more female stage (Teuchert 1968). Based on the observations on a number of species from both Turbanella and Paraturbanella, Balsamo et al. (2002) reject the occurrence of sequential hermaphroditism at least for these two genera. The definite presence of sequential hermaphroditism in macrodasyidan Gastrotricha has to be confirmed urgently by tracing specific individuals with a known life history and individual age from laboratory cultures (Hummon & Hummon 1992). During her studies of the reproductive biology of marine gastrotrichs from the North Sea, and the Baltic Sea, Teuchert (1968) also noticed that species such as Mesodasys laticaudatus, Macrodasys caudatus, Dactylopodola baltica, or Acanthodasys aculeatus were characterized by a continuous reproductive activity during the warm season with a peak from late summer to autumn. However, a second group of species consisting of Cephalodasys maximus and two species of Turbanella (T. hyalina and T. cornuta) displayed two reproductive peaks, one in spring and, isochronic to that of the aforementioned species, a second during late summer to autumn (Teuchert 1968). Interestingly, a recent study focusing the molecular diversity of Turbanella cornuta and T. hyalina from the North Sea and Baltic Sea area gave evidence for at least two sympatric cryptic species within original T. hyalina (Kieneke et al. 2012). It is discussed whether the two reproductive peaks of T. hyalina reported by Teuchert (1968) could represent the reproductive maxima of in fact two separate cryptic species. Although explicitly studied and proofed in two instances only, all hermaphroditic species of the Macrodasyida and Neodasys (Multitubulatina) that possess a set of two accessory reproductive organs, i.e., the caudal organ plus frontal organ or at least one of both structures, should engage in mating and cross-insemination, during which spermatozoa are transferred from one mating partner to the other and vice versa. The mode of reciprocal sperm transfer/cross-insemination with the aid of accessory reproductive organs requires a certain kind of copulation

1.3 Reproduction and development 

 79

behavior. This has in fact been investigated and documented in two species of the Macrodasyida. Teuchert (1968) describes the copulation of Turbanella cornuta (see also Hummon & Hummon 1992 for a review), a macrodasyidan gastrotrich that has a frontal organ but obviously lacks a caudal organ (but see the interpretation of Ruppert 1991 that is presented later). In this species, an individual in the male stage lifts its caudum and performs pendulous movements in order to touch a specimen in its female stage ready to mate. Within a few seconds, the two individuals first form a loose, later a tight knot with their rear trunks (see figure 2 of Teuchert 1968). Tightening the knot is achieved by gluing to the substratum with the aid of the batteries of anterior adhesive tubes and stretching the trunks in opposite directions. At maximum 70 seconds later, both animals release again. Based on the insufficient knowledge of the reproductive anatomy of T. cornuta during that time, Teuchert (1968) was not able to clarify satisfactorily how sperm transfer really happens. As already mentioned, T. cornuta lacks a caudal organ as a sperm-transferring device. Just the unpaired ventral male gonopore of the fused sperm ducts can be used for insemination. The female gonopore of T. cornuta was assumed to be situated on the ventral side close to the anus. However, these anatomical conditions did not fit to the observed copula because in neither stage did male and prospective female pores of the mating animals get into contact (Teuchert 1968). Based on the well-known mating mechanism of Macrodasys sp. (see below) and on the unpublished morphological data of Turbanella ocellata, Ruppert (1991) reinterprets the mating and copulation in T. cornuta. He suggests that there are in fact two accessory reproductive organs in Turbanella, a frontal organ described by Teuchert (1977a) as the “gland organ of the intestinal region” with unknown function and a caudal organ close to the anus. Hence, Turbanella would first have to charge its own caudal organ like in Macrodasys before spermatozoa are transferred from this organ to the frontal organ of the mating partner (Ruppert 1991). The “gland organ of the intestinal region” is a paired structure situated close to the uterus region with the mature egg (Teuchert 1977a). Kieneke et  al. (2009) interpret this structure to be a socalled cervix, an outlet duct for the mature and fertilized egg (see also below). Such an interpretation still complicates the issue of mating and sperm transfer in T. cornuta. Further input to the mating and copulation in Turbanellidae comes from a con-familiar species of T. cornuta, Paraturbanella teissieri. This species has a frontal organ that definitely has a dorsolateral opening. The presence of a caudal organ, however, was rejected and therefore a copulation by contact of the slightly protrudable ventral male

80 

 1 Gastrotricha

gonopore (common opening of fused sperm ducts like in Turbanella and other Turbanellidae) and the dorsal pore of the frontal organ (see, e.g., Fig. 1.34 A) of two animals during mating is hypothesized to be the mode of copulation in P. teissieri and other Turbanellidae (Balsamo et al. 2002). Much better known in terms of reproductive biology and anatomy is Macrodasys. Based on intensive histological, SEM, TEM, and live observations of two undescribed species, Ruppert (1978a) was able to fully reconstruct the mating and copulation behavior and the structures and morphofunctional processes involved in reciprocal sperm transfer (see also Ruppert 1991, Hummon & Hummon 1992 and Balsamo 1992 for reviews on this issue). In this taxon, two mating animals pair and copulate by reciprocally protruding their tubular parts of the caudal organ (“copulatory tube”) into the frontal organ of the partner (Fig. 1.48). Openings of both accessory reproductive organs are

situated on the ventral surface of the animal. Once this copula is established, own spermatozoa (autosperm) are released from the testes and enter a posterior compartment of the own caudal organ called the antrum feminimum (see chapter Reproductive Organs). Owing to a severe bending of the rear trunk within a knot of two animals, both sperm duct openings are in close proximity to the ventral pore of the caudal organ. Hence, spermatozoa may easily reach the caudal organ after leaving the sperm ducts. From the antrum feminimum, spermatozoa enter the own copulatory tube that already sticks inside the frontal organ of the mating partner. Once the copulatory tube of each mating animal is charged with autosperm, it breaks off from the remaining tissue of the caudal organ (Fig. 1.48). Shortly after, the mating animals release and glide away with their frontal organs filled with foreign spermatozoa (allosperm) and the copulatory tube from the partner (Ruppert 1978a). The whole process of mating and copulation in Macrodasys

A I

F I

II

B

I

E I II

C

I

D I

II

II

testes frontal organ caudal organ & copulatory tube antrum feminimum spermatozoa

Fig. 1.48: Mating and reciprocal sperm transfer in Macrodasys. For simplicity, transfer is only displayed from one (I) to the other specimen (II). Second animal (II) drawn as a cross section at the level of its frontal organ. (A) Simplified diagrammatic reproductive anatomy of Macrodasys, ovary omitted. For more details, see Fig. 1.33 C. (B) Precopulatory behavior. The copulatory tube is still invaginated. (C) Attachment. Mating animals evert their copulatory tubes into the frontal organ of the partner. (D) Uptake of autosperm into the caudal organ via the antrum feminimum. (E) Reciprocal sperm transfer. Spermatozoa move, probably by beating action of their flagella, into the copulatory tube. (F) Separation of mating partners. (Modified from Ruppert 1978a, 1991 and Hummon & Hummon 1992.)



occurs in just a few seconds (see figure 2 in Ruppert 1978a). A quite differentiated behavior before and during the actual copulation is allied with these morphofunctional processes for reciprocal insemination in Macrodasys (Ruppert 1978a, reviewed in Ruppert 1991 and Hummon & Hummon 1992). Pre-copulatory actions may involve gregarious behavior of animals as a kind of prelude to mating. However, gregariousness in Gastrotricha could also have reasons different from reproduction and mating (Hummon & Hummon 1992, 1993). During copulation, two specimens of Macrodasys sp. form a complicated knot with their posterior trunk portions (Ruppert 1978a). Within the knot, the copulatory tube of one animal gets into contact with the frontal organ pore of the other and vice versa. Furthermore, each specimen within the knot brings its male gonopores in close proximity to its own antrum thereby enabling the transition of the autosperms from the testes to the caudal organ. The general scheme of reciprocal sperm transfer in Macrodasys – pairing, copulation, charging the caudal organ with autosperm, transferring sperm from the caudal organ into the frontal organ of the mating partner, and detachment – is assumed to occur in many species of hermaphroditic gastrotrichs (Ruppert 1991). Since anatomic conditions vary a lot among the different taxa (see chapter Reproductive Organs), different deviations from this general scheme will certainly exist. For instance, sperm ducts directly discharge into the caudal organ lumen in the taxa Mesodasys and Thaumastodermatidae. This means that the caudal organ may have already been charged with autosperm internally before the actual mating and copulation with a reproductive partner happens. In Thaumastodermatinae, a subtaxon of Thaumastodermatidae, the continuity of sperm duct, caudal organ, and frontal organ may also indicate the possibility of self-fertilization (Ruppert 1991, see also chapter Reproductive Organs). This hypothesis, however, has never been tested. Another deviation from the “Macrodasys scheme” of mating and copulation may be present in some species of Urodasys that possess a cuticularized stylet within their caudal organ (Schoepfer-Sterrer 1974). Those hard parts could in principle be used for hypodermal impregnation. Such a function of the stylet in different members of Urodasys, however, has never been demonstrated (Balsamo 1992) and was already rejected by Schoepfer-Sterrer (1974). Another mode of reciprocal sperm transfer in marine Gastrotricha is the formation and exchange of spermatophores (see also chapter Reproductive Organs). Spermatophores, i.e., oval to spherical packages of spermatozoa embedded inside a common sheath or matrix, definitely occur in species of Dactylopodola (Teuchert 1968, Ruppert 1991, Kieneke et al. 2008d) and Neodasys (Guidi et al. 2003, Kieneke et al. 2009) and have recently also been reported

1.3 Reproduction and development 

 81

from the newly described species Urodasys poculostylis (Atherton 2014). However, the exact mode of spermatophore exchange and the involved organs are still poorly understood. In Dactylopodola baltica, spermatozoa are internally packed into spermatophores but it is not known which organ facilitates spermatophore formation. The spermatophore is then extruded to the outside and glued to another animal where spermatozoa leave the later degenerating spermatophore by an undetermined mechanism (Teuchert 1968). For another species of Dactylopodola from the US Atlantic coast, it is hypothesized that the complete frontal organ is first charged with spermatozoa internally via a narrow channel that connects sperm ducts with the frontal organ lumen. Later, the whole frontal organ shall be extruded in toto as the actual spermatophore (Ruppert 1991). The function of the also present caudal organ in this species, however, is not clearly visible according to that hypothesis. Solely based on their morphological reconstructions, Kieneke et al. (2008d) hypothesize that in Dactylopodola typhle that spermatozoa must be transported externally from the testes into the lumen of the glandular caudal organ prior to mating. Here, mucous secretions are released and build up the spermatophore, which is then pressed from one animal, mediated by contractions of the rear trunk musculature, into the dorsolateral female gonopore of a mating partner. The spermatophore reaches a position beneath the mature oocyte inside the uterus via a short duct called the cervix. Again mainly based on ultrastructural morphological data, another mode of spermatophore formation was hypothesized for Neodasys chaetonotoideus (Kieneke et al. 2009). The formation of spermatophores in N. chaetonotoideus is assumed to occur completely external by the aid of secretion products that originate from the caudal organ. This assumption is supported by the observation that mature specimens may carry a spermatophore adhered to the posterior adhesive appendages (Kieneke et al. 2009). Inseminated specimens of N. chaetonotoideus display one, sometimes more (own unpublished observation, but see also figure 113 in Ruppert 1991) spermatophores inside their frontal organ. The mechanism how the spermatophore enters the frontal organ is obscure (Kieneke et al. 2009). All these examples highlight the urgent need of detailed behavioral studies of gastrotrichs in combination with morphological reconstructions to fully understand the reproductive biology of Macrodasyida and Neodasys. Apart from hermaphroditism, there are also few instances of parthenogenetic species among Macrodasyida that only possess female gonads and lack any accessory reproductive organs. Candidates for a solely unisexual reproduction are Urodasys viviparus, Paradasys subterraneus (and probably further species of Paradasys),

82 

 1 Gastrotricha

Redudasys fornerise (one of only two species of Macrodasyida inhabiting freshwater), and Anandrodasys agadasys (Wilke 1954, Kisielewski 1987a, Todaro et al. 2012b, Kieneke et al. 2013a). However, Valbonesi & Luporini (1984) described specimens of U. viviparus from the Somalian coast that obviously possessed a testis and a caudal organ that indeed indicates that hermaphroditic reproduction in that species occurs (see also Hummon & Hummon 1993). Apart from the questionable parthenogenesis in this species, U. viviparus is still of interest in terms of reproductive biology because it is the only known gastrotrich species that is ovoviviparous (Hummon & Hummon 1983a). Knowledge about reproductive biology of the gastrotrich subgroup Paucitubulatina is much more fragmentary than in Macrodasyida and Neodasys. Marine species of the taxa Xenotrichulidae and Muselliferidae are, with only few exceptions (the xenotrichulids Draculiciteria tesselata and Heteroxenotrichula pygmaea are much likely obligate parthenogenetic species, see Wilke 1954 and Ruppert 1979, reviewed in Balsamo 1992 and Hummon & Hummon 1993), hermaphroditic animals like most Macrodasyida and Neodasys. However, apart from species of Musellifer, which may possess putative accessory reproductive organs (Hummon 1969, but see Leasi & Todaro 2010), there are no comparable organs in the con-familiar taxon Diuronotus (Todaro et al. 2005, Balsamo et al. 2010a). Although Ruppert (1979) frequently reports an unpaired ventral copulatory organ in Xenotrichulidae at the site where both vasa defferentia fuse, such a structure has not been confirmed so far. There might be a glandular structure of unknown function in this region as in Xenotrichula punctata (Ferraguti et al. 1995). Owing to this predominant absence of accessory reproductive organs in simultaneous hermaphroditic Paucitubulatina (Muselliferidae and Xenotrichulidae), it is incomprehensible how sperm transfer might be achieved. To our knowledge, there is no assured report of reproductive behavior in any taxon of the Gastrotricha-Paucitubulatina. In their culture dishes, Balsamo & Todaro (1987) encountered some pairs of individuals of Aspidiophorus polystictos that were close to each other with their caudal ends. However, it was not possible to determine if this behavior enabled effective mating. By chance, we have uniquely observed two individuals of Xenotrichula intermedia in a Petri dish of freshly extracted meiofauna, which slowly orbited each other with slightly decreasing radius. The moment they were touching, the animals crawled around each other quite quickly but already released and departed after 1–2 seconds (unpublished observation). This was possibly the moment of mating and exchange of spermatozoa. However, due to the obvious absence of any copulatory or sperm storing device in Xenotrichulidae (see above), it is absolutely dubious

how cross-insemination will be facilitated by members of this taxon. The frequent occurrence of coiled spermatozoa on the surface of X. intermedia from an Arabian population is possibly related with sperm transfer modalities in this species (Leasi & Todaro 2009). However, this observation is raising more questions than providing answers in terms of reproductive biology in Xenotrichulidae. The predominantly freshwater inhabiting paucitubulatinan taxa Chaetonotidae and Dasydytidae were for long regarded to reproduce solely by parthenogenesis (e.g., Remane 1936, Hyman 1951). However, as already presented in the chapters about reproductive organs and gametes, reproductive development, and fine structure of the freshwater paucitubulatinan species Lepidodermella squamata (Chaetonotidae) was studied and described in detail in a series of papers (Hummon 1984a–c, 1986) and has finally demonstrated the frequent development of hermaphroditic specimens in this species. L. squamata first reproduces by parthenogenetic formation of tachyblastic (subitaneous) and opsiblastic (resting) eggs. After the parthenogenetic phase, packets of simple, rod-shaped sperms are produced and few oocytes develop into opsiblastic eggs. Spermatocytes and oocytes undergo meiosis during this hermaphroditic phase. A comparable mode of parthenogenesis followed by the production of simplified spermatoza was also demonstrated in a marine species of Chaetonotidae, Aspidiophorus polystictos (Balsamo & Todaro 1987, 1988). Hence, L. squamata, A. polystictos, and probably many other Chaetonotidae actually represent protogynous hermaphrodites rather than obligate parthenogens (Hummon & Hummon 1992). With the development of eggs and sperm in A. polystictos, L. squamata, and many more freshwater-dwelling species of the Chaetonotidae and Dasydytidae (e.g., Weiss & Levy 1979, Kisielewska 1981, Weiss 2001), essential features for a bisexual reproduction with recombination by cross-fertilization during a hermaphroditic phase are present in these taxa of the Paucitubulatina. However, it was until now not possible to demonstrate cross-fertilization in Chaetonotidae or Dasydytidae (e.g., Hummon & Hummon 1992, 1993, Weiss 2001), or even in the well-studied Lepidodermella squamata (Hummon 1986). Although this important proof was and is still missing a partly hypothetical biphasic life cycle for the Chaetonotidae (probably also applicable to the remaining parthenogenetic-hermaphroditic paucitubulatinan taxa Dasydytidae, Neogosseidae, Proichthydidae, and Dichaeturidae) has been developed (Levy 1984, see also Strayer & Hummon 1991 and Balsamo 1992 for summaries). According to this scenario, that is fully congruent with the results on reproductive development in L. squamata (Hummon 1984a–c, 1986), species of Chaetonotidae reproduce by two different reproductive phases (biphasic

1.3 Reproduction and development 



 83

HERMAPHRODITIC PHASE parthenogenetic female

hermaphrodites sp oo T T T

D juvenile

MEIOSIS

O PB 2n

oo F

sp

1n

AMPHIMIXIS

PARTHENOGENETIC PHASE (APOMIXIS)

life cycle). The first is the parthenogenetic phase (Fig. 1.49) where animals mostly produce four or five apomictic (2n) eggs, the last of which is almost always a resting (opsiblastic) egg while the others are subitaneous (tachyblastic) eggs. The parthenogenetic phase enables a rapid population growth and exploitation of resources under favorable conditions. Production of resting eggs serves as a “buffer” against periods of unfavorable ecological conditions (Strayer & Hummon 1991). When the parthenogenetic resting egg is spawned, animals enter the second, post-parthenogenetic (hermaphroditic) phase (Fig. 1.49) where bilateral packets of simplified sperm (1n) are produced by meiosis, generally 32–64 per animal (see also chapters about reproductive organs and gametes). The ovary produces meiotic eggs (1n) during the post-parthenogenetic phase. It is assumed that two of such hermaphroditic individuals exchange gametes by a yet undetermined mechanism by which fertilization is achieved and a zygote (2n) is formed. Owing to the complete absence of accessory reproductive structures as well as sperm ducts in L. squamata and other chaetonotids that may serve for sperm transfer (the enigmatic secretory x-organ is regarded to play another role in reproduction, see chapter Reproductive Organs), one possibility for sperm transfer, although quite bizarre, might be the ingestion of spermatozoa by cannibalism between hermaphrodites (Hummon 1986). The relative length of the hermaphroditic phase compared with the parthenogenetic phase (40%– 70% versus 20%–40%, a short postembryonic phase not considered for these values, see Balsamo 1992) may under-

F

Fig. 1.49: Hypothetical biphasic life cycle of Chaetonotidae (Paucitubulatina). Pharthenogenetic females generally produce 4 apomictic eggs; the last one is almost always a resting (opsiblastic) egg, whereas the others are subitaneous (tachyblastic) eggs. After depositing the last parthenogenetically produced egg, animals become hermaphrodites and develop simple sperm and further eggs that undergo meiosis. If fertilization really occurs has yet to be proved. A third egg type (PB) discovered in cultures of Lepidodermella squamata could represent fertilized eggs. The broken lines indicate still unproven pathways. Abbreviations: D, dormancy; F, fertilization; O, opsiblastic egg; oo, oocyte; PB, plaque-bearing egg; sp, sperm rod; T, tachyblastic egg. (Modified from Strayer & Hummon 1991 and Balsamo 1992.)

line the importance of presumed sexual reproduction with recombination in these predominantly freshwater taxa. Ovipository behavior was studied in several species of Macrodasyida by Teuchert (1968). Species such as Cephalodasys maximus and Mesodasys laticaudatus spawn only one egg at a time, whereas species of Turbanella, Dactylopodola baltica, or Urodasys mirabilis may deposit one to two mature eggs, Macrodasys caudatus is able to spawn seven to eight eggs at once. Because eggs of macrodasyidan gastrotrichs are provided with a sticky secretory covering (up to 20 µm thick like in Turbanella cornuta, see figure 7C of Teuchert 1968), the animals are able to glue the laid eggs immediately to sand grains. In Macrodasys caudatus, a short chain or clutch of eggs may be produced due to this glutinousness. A putative origin of the cohesive covering of readily laid eggs has recently been discovered in Dinodasys mirabilis. The so-called posterior gland organ of this species is hypothesized to synthesize the appropriate secretion (Todaro et al. 2012a). Egg deposition involves a special behavior where animals slow down their creeping locomotion and seem to sense the substratum for suitability (e.g., chemical factors, surface conformation, texture of sediment grains). It is possible that gravid animals engage in site selection to choose a spot with a certain protection for the laid egg (Hummon & Hummon 1983a). Such a behavior might be the only contribution to parental care in Gastrotricha. Immediately prior to egg deposition, species like Turbanella cornuta turn their body and

84 

 1 Gastrotricha

A

B

*

gc

mo

mg

em

tes ov

100 µm

25 µm

ph

press their back to the substratum to cause a rupture of the integument (Teuchert 1968; Fig. 1.50 A). This behavior is supported by muscular activity and adherence with the anterior tubes. Macrodasys caudatus and Urodasys mirabilis engage in repeated contraction and stretching of the body to press the eggs to the outside via a lateral rupture of the body wall. Again, the animal uses its anterior adhesive tubes to attach to the sediment during egg deposition. Dactylopodola baltica suddenly stops its jerky movements and executes a frontally directed pressure, probably caused by muscle contractions. This pressure causes again a rupture of the body wall close to the uterus (Teuchert 1968). Contraction of the longitudinal muscles also precedes egg deposition in freshwater chaetonotids. Here, mature eggs leave the animal ventrally, possibly through a yet undiscovered ovipore (Hummon & Hummon 1983a). The rupture of the integument to release mature and fertilized eggs in Macrodasyida was repeatedly reported in different macrodasyidan gastrotrichs (Teuchert 1968, see above). Also, in the ovoviviparous species Urodasys viviparus, a rupture of the body wall of the mother animal was assumed to give birth to the juvenile because no female gonopore could be observed in this species (Wilke 1954). The rather bizarre process of body wall rupture gets support by the observation of presumptive wounds in the integument that may persist as small bulges for two or even more days in the area where the egg has left the trunk (Teuchert 1968; Fig. 1.50 A). However, in different species of the Thaumastodermatidae (Ruppert 1978b) and in Dactylopodola typhle a short outlet duct with an at least preformed dorsolateral pore is present. This organ was later termed the “cervix” (Kieneke et al. 2009). Presence of a cervix is also supposed in Turbanella cornuta

mg

Fig. 1.50: Spawning of Macrodasyida. (A) Turbanella hyalina that has currently deposited an egg. Note the area in the integument where the egg has left the body (asterisk). It is still dubious whether the egg leaves the mother by rupture of the body wall or through an inconspicuous pore (see main text for details). (B) An embryo of Urodasys viviparus inside the egg envelope and still inside the mother animal. U. viviparus is so far the only known ovoviviparous gastrotrich species. (A–B): DIC images. Abbreviations: em, embryo; gc, glutinous covering of the egg; mg, midgut; mo, mature (spawned) egg; ov, ovary; ph, pharynx; tes, testis.

(Kieneke et al. 2009 regard the “gland organ of the intestinal region” as a potential cervix, see above), a species in which egg deposition shall actually happen via rupture of the body wall. As the cervix and female gonopore in D. typhle was only detectable by TEM or serial histological sections, not even by differential interference contrast (DIC) microscopy, it is possible that such an outlet duct for egg deposition is in fact present (but has not been detected yet) in species that are believed to spawn by rupture of the body wall (Kieneke et al. 2009). The observed wounds in different species of Macrodasyida (Teuchert 1968) could in fact be swollen epidermal cells that surround the female gonopore after spawning. Such a situation was observed in a specimen of D. typhle that was sectioned for histological investigation (see figure 10E–F in Kieneke et al. 2008c).

1.3.2 Cleavage and development There are still quite few investigations on the cleavage of the fertilized egg. First observations were made by Ludwig (1875), who observed the first cleavage steps in Chaetonotus larus eggs. De Beauchamp (1929) and Brunson (1949) added further observations from a total of three chaetonotid species, and Sacks (1955) provided the most detailed descriptrion of the embryology of Lepidodermella squamata. Mock (1979) stated that the first cleavage steps in two Xenotrichula species correspond to Sacks’ reports. Macrodasyids were investigated even more rarely, initial observations come from Swedmark (1955) on Macrodasys affinis, whereas Teuchert (1968) provided a detailed description of the cleavage and development of Turbanella cornuta, with

1.3 Reproduction and development 



additional data from Cephalodasys maximus and Macrodasys caudatus. Cleavage starts within a few hours after oviposition, after already 25–35 minutes in L. squamata (Sacks 1955), after about 2 hours in T. cornuta, after 2–3 hours in M. caudatus, and after 3–4 hours in C. maximus (Teuchert 1968). The first cleavage is total and results in two blastomeres of equal size (named AB and CD; Fig. 1.51 A). The second cleavage plane is perpendicular to the first plane and results in a shift of two blastomeres in a 90° angle compared with the other two blastomeres. This stage is called rhomboid (Teuchert 1968) or tetrahedral (Malakhov 1994). To reach this stage, some differences exist between the observed chaetonotids and macrodasyids. In L. squamata, there is a slight asynchrony in the cleavage of the two blastomeres from the first cleavage and also a slight difference in size (Sacks 1955). Blastomere AB divides slightly earlier than blastomere CD. The resulting blastomeres C and D appear to be smaller than blastomeres A and B. In the observed macrodasyids, there are no size differences between the blastomeres A to D, but the asynchrony in division is more pronounced, which leads to an intermediate three-cell stage (Teuchert 1968). In L. squamata, the asynchrony in cleavage persists throughout the entire cleavage, the anterior blastomeres divide before the posterior ones. Additionally, size differences become pronounced and dorsal blastomeres are smaller than ventral ones (Sacks 1955). After the fifth cleavage, the 32-cell stage constitutes the blastula with a small blastocoel in the center. In the four-cell stage of macrodasyids, one of the posterior blastomeres (called C in Teuchert 1968) moves anteriorly to a central position where it touches all three other blastomeres, or in some cases even further, to the level of the blastomeres A and B. The position of this blastomere C marks the dorsal side of the developing embryo. The following cleavages are asynchronous. After the third cleavage, another shift of one cell occurs in T. cornuta. Blastomere D divides into a dorsal and a ventral cell, the ventral one is called E. This blastomere moves into a central, ventral position and becomes the “urmesoderm cell”. In this eight-cell stage, the blastomeres detach from each other in the central part and form a small blastocoel. Consequently, Teuchert (1968) calls this eight-cell stage already a coeloblastula. In M. caudatus, no “urmesoderm cell” or blastocoel exists. A blastocoel occurs during the fourth cleavage in this species. In T. cornuta, the subsequent cleavages are asynchronous, with the “urmesoderm cell” dividing as the

 85

A nu

nu

15 µm

B *

15 µm

C

15 µm

D ph

fu

15 µm Fig. 1.51: Development of the freshwater gastro­trich Lepidodermella squamata (Paucitubulatina). (A) Two-cell stage. Note the sculptured egg shell. (B) Two cells migrated between the two central ventral cells (to the bottom in the image) into the blastocoel (asterisk). The blastopore is now closed. (C) Multicel­lular embryo during organogenesis. (D) Almost fully shaped embryo. (A–D): DIC images. Abbreviations: fu, furca with adhesive tubes; nu, nucleus; ph, pha­r ynx. All images kindly by Andreas Hejnol, Bergen.

last cell. The two resulting “urmesoderm cells” remain undivided during the sixth and the seventh cleavage (Teuchert 1968). The two “urmesoderm cells” have extensions into the blastocoel, which can be interpreted as an early indication of gastrulation. Embryos of both T. cornuta and M. caudatus are bilaterally symmetrical. For gastrulation in T. cornuta, both “urmesoderm cells”

86 

 1 Gastrotricha

sink into the blastocoel. Owing to the small size, the resulting archenteron is more or less a ventral pit. By stretching and bending of the embryo, a ventral furrow is created, which is closed by growth of the lateral ectodermal cells. Mesodermal cells originate from cells bordering the two “urmesoderm cells”, they migrate into the embryo during gastrulation. In L. squamata, gastrulation follows the fifth cleavage (Sacks 1955). It consists of an inward migration of two ventral cells, called A5.3 and A5.4, into the blastocoel (Fig. 1.51 B). This progress is similar to T. cornuta, but the two ventral cells have originated in a different way. The further development of the entoderm is also described in a different way. According to Sacks (1955), the blastopore is closed for a brief moment, then cells in this region proliferate and form a ring around the more and more pronounced anterior invagination. Another invagination forms at the posterior end of the embryo, these two invaginations extend and approach each other. At this time, the anterior invagination can already be recognized as the pharynx, whereas the posterior invagination appears to be the developing midgut. Teuchert (1968) has produced a cell lineage for T. cornuta. As described above, the entoderm derives exclusively from the cell E, the “urmesoderm cell”. Mesoderm and ectoderm derive from different cells from all remaining lineages. Embryos develop gradually into juveniles, which resemble miniature adults when they hatch from the egg capsule (e.g., Remane 1936, Teuchert 1968; Fig. 1.51 C, D). In chaetonotids, cuticular structures are recognizable in late embryos inside the eggs (Mock 1979). No larval stages are present, and no molting takes place. Hatched juveniles grow and develop gradually into mature specimens. There are, besides size differences and the absence of the reproductive organs, two main differences between juveniles and adults. They have (in macrodasyids) fewer adhesive tubes and fewer pairs of protonephridia than adults (Teuchert 1968), and the size relations between pharynx and midgut are different than in adults (Fig. 1.52 A–C). Usually, in juveniles, the pharynx is comparably longer than the midgut, whereas in adults, it is vice versa (e.g., Remane 1924 for Macrodasys buddenbrocki, Hochberg 1998 for Turbanella mustela). Some sources, especially from popular science, state that gastrotrichs are an example for eutely, which means that the cell number, including the germ line, is fixed. Hence, all cell divisions must be finished during embryogenesis with no further mitoses after this period

A

B

200 µm

50 µm

C

100 µm Fig. 1.52: Juvenile specimens of Gastrotricha. During postembryonic development organ sys­tems such as adhesive tubes, epidermal glands and protonephridia are increased and the go­nads occur. (A) Macrodasys caudatus (Mac­rodasyida), dorsal view. (B) Turbanella hyalina (Macrodasyida), ventral view. (C) Neodasys cha­etonotoideus (Multitubulatina), dorsal view. (A–C): BF images.

(see Balsamo et al. 1999). This requires a strictly determinate cleavage with an invariant cell line, two facts that are not known with certainty for gastrotrichs. At least in macrodasyids, eutely is certainly not the case. Manylov (1995) documented a complete regeneration after transsection in Turbanella sp. and also the egg release by rupture of the body wall in those species without a female gonopore can only be explained by regeneration capacity, which involves the generation of new cells. Also, the assumed mitotic proliferation of spermatogonia within the testes of at least some species of the Macrodasyida contradicts a strict definition of eutely (Balsamo et al. 1999).

1.4 Physiology 

1.4 Physiology Not much data are known about physiological properties of gastrotrichs. Most gastrotrichs are aerobic and live in the upper layers of sediments or above the sediment. Because some species occur deeper in the sediment, anaerobiosis is assumed to be possible (Boaden 1985), although it is not known whether this is just a tolerance of low oxygen levels or whether anaerobic metabolic pathways are present. Boaden (1974) described three species [Turbanella reducta, T. thiophila (=T. bocqueti) and Thiodasys sterreri (=Megadasys sterreri)] from gray and black sands rich in sulfide. The three species form a series occurring in more and more anoxic and sulfidic sediments. Megadasys sterreri is the species that appears most adopted to anoxic conditions. It was kept in sealed jars with anoxic sand for more than 2  months (Boaden 1974). An ultrastructural investigation showed that mitochondria seem to have fewer cristae than in other species, and it is possible that CO2 can be fixed (Boaden 1974). The complete absence of mitochondria has been observed in the spermatozoa of two Urodasys species (Balsamo et  al. 2007). One possible explanation is that this is an adaptation to anoxic sediments. Neodasys sp. has a blood-red appearance, which is caused by red cells that are assumed to be homologous to Y-cells (Kraus et al. 1981, Ruppert & Travis 1983). This is in contrast to other species with colorless Y-cells and was investigated in further detail. The red cells are arranged in two longitudinal rows dorsolateral of the gut and make up about 14% of the body volume (Ruppert & Travis 1983). Branches of these cells surround perikarya of muscle and nerve cells, whose mitochondria are close to this connection site (Ruppert & Travis 1983). The cells contain intracellular hemoglobin (Colacino & Kraus 1984), as had been suspected by Kraus et  al. (1981). Colacino & Kraus (1984), and Kraus & Colacino (1984) compared the oxygen consumption rate of Neodasys sp. with that of Turbanella ocellata and Dolichodasys carolinensis, but the rate of Neodasys sp. is between that of the two other species. Therefore, it is assumed that the possession of hemoglobin does not affect oxygen consumption, but may be important for oxygen storage, probably for sporadic movements to anoxic sediments.

1.5 Phylogeny In the 1920s, the German zoologist Adolph Remane discovered dozens of new taxa of marine gastrotrichs inha-

 87

biting the spaces between sand grains. By then, a variety of freshwater species, one of the earliest descriptions was provided by Ehrenberg (1838), and two marine ones, Hemidasys agaso (Claparède 1867) and Turbanella hyalina (Schultze 1853) were known to science. The freshwater species were considered to represent a separately evolved group of microscopic animals, the Gastrotricha (e.g., Mečnikow 1865, Zelinka 1889), which initially included the marine Turbanella (see Mečnikow 1865). Later, an affiliation of the two only known marine species (T. hyalina, H. agaso) to this taxon was dismissed (e.g., Wagner 1893). But in view of the large quantity of new marine forms, which were first called the “aberrante Gastrotrichen” (Remane 1924, 1925a, b), a revision of this group of exclusive meiobenthic organisms was required. At the latest with Remane’s monograph (Remane 1936) the traditional system of Gastrotricha has been shaped. He separated two orders among the Gastrotricha, which, at that time, represented a class of the Aschelminthes (see Schmidt-Rhaesa 2013): (1) the exclusive marine to brackish Macrodasyida (originally termed Macrodasyoidea) and (2) the marine as well as fresh and brackish water inhabiting Chaetonotida (originally termed Chaetonotoidea). Decades later, it was d’Hondt (1971) who erected two chaetonotidan suborders to highlight the isolated position of the marine taxon Neodasys. From then on, Gastrotricha comprised the two sister taxa Macrodasyida and Chaetonotida with Multitubulatina (=Neodasys) and Paucitubulatina (=all remaining chaetonotids) being sister taxa of highest rank within the Chaetonotida. When originally described, Remane (1927a, 1929) provisionally assigned Neodasys to the Macrodasyida but later he relocated it to the Chaetonotida based on histological findings (Remane 1936). Much later, a cladistic analysis of a variety of morphological characters (Hochberg & Litvaitis 2000; Fig. 1.53 A; see also Hochberg & Litvaitis 2001e) confirmed this traditional system and further provided ideas of phylogenetic relationships among the different genera of Gastrotricha. Monophyly of several of the traditional families was supported, while for others, it was declined (Hochberg & Litvaitis 2000, 2001e). Phylogenetic discussions that were focused on ultrastructural features of, e.g., the pharynx or the body wall of different gastrotrich species (Ruppert 1982, Travis 1983) confirmed the traditional thoughts of the basal internal relationships and provided some hypotheses on evolutionary traits. According to these studies, species of the subtaxon Macrodasyida would be characterized by the possession of pharyngeal pores and a pattern of an inverted Y of the cross-sectioned pharyngeal lumen, whereas members of the taxon Chaetonotida lack pharyn-

88 

 1 Gastrotricha

geal pores and show a Y-shaped pharyngeal lumen when cross-sectioned (see, e.g., Ruppert 1991). However, these characteristics of the within-groups of highest rank leave unresolved the character pattern of the common ancestor because we have conflicting character states in the basal node of Gastrotricha (absence versus presence of pharyngeal pores; Y-shaped versus inverted Y-shaped pharyngeal lumen). Initial DNA sequence-based cladistic analyses or combined morphological and molecular studies revealed successively the paraphyly of Paucitubulatina (Wirz et al. 1999; Fig. 1.53 B), the paraphyly of Macrodasyida (Zrzavý 2003; Fig. 1.53 D) or even polyphyly of whole Gastrotricha with both major subgroups (Chaetonotida, Macrodasyida) being monophyletic each but with Chaetonotida

A

B

C

D

E

F

G

H

more closely associated with taxa such as Plathelminthes, Gnathostomulida or Rotifera rather than with Macrodasyida (Manylov et al. 2004; Fig. 1.53 E). However, all these pioneer studies lack important taxa such as Neodasys, which are of highest systematic relevance. More recent cladistic analyses based on partial 18S rRNA gene sequences comprise a more satisfying taxon sampling among the gastrotrich genera (Todaro et al. 2003, 2006a, Petrov et  al. 2007, Paps & Riutort 2012). The mentioned analyses reveal Paucitubulatina as a monophyletic group but Macrodasyida being paraphyletic with Neodasys nesting within a sub clade comprising different macrodasyid genera (Todaro et al. 2003: ML analysis; Fig. 1.53 C, Todaro et al. 2006a; Fig. 1.53 F). Alternatively, Macrodasyida form a monophyletic group but include again Neo-

Fig. 1.53: Basal internal relationships of Gastrotricha according to different analyses using different character systems and methods of inference. In some cases, the positions of putative basal genera (in italics) are indicated. Note that the tree topologies are strongly simplified. (A) Relationships according to consensus tree (50% majority rule, maximum parsimony) of Hochberg & Litvaitis (2000). This scenario is also congruent with late systematization of the Gastrotricha (Remane 1936, d’Hondt 1971). (B) Relationships according to neighbor joining tree of Wirz et al. (1999). (C) Relationships according to maximum likelihood tree of Todaro et al. (2003). (D) Relationships according to summary tree of Zrzavý (2003). (E) Relationships according to Bayesian tree of Manylov et al. (2004). (F) Relationships according to Bayesian tree of Todaro et al. (2006). (G) Relationships according to Bayesian tree of Petrov et al. (2007)/maximum likelihood tree of Paps & Riutort (2012)/ single most parsimonious tree of Todaro et al. (2003). This scenario is also congruent with early systematization of the Gastrotricha (Remane 1929). (H) Relationships according to consensus tree (50% majority rule, maximum parsimony) of Kieneke et al. (2008). Abbreviations: Cephal., Cephalodasys; Dact., Dactylopodola; PAUCIT., Paucitubulatina; Xeno., Xenodasys. *The lineage comprises Redudasys and Marinellina.

1.5 Phylogeny 

dasys (Todaro et al. 2003: MP analysis, Petrov et al. 2007, Paps & Riutort 2012; Fig. 1.53 G). The latter scenario ironically represents Remane’s earlier system of Gastrotricha (e.g., Remane 1927a, 1929) before he assigned Neodasys to the Chaetonotida. The use of characters related with sperm morphology and sperm ultrastructure of 28 species of Gastrotricha grouped most of the included species of Macrodasyida as a monophyletic clade but did not reveal the Paucitubulatina as a monophylum (Marotta et al. 2005). Furthermore, the maximum parsimony analysis placed two included macrodasyidan species of the taxon Dactylopodola as the sister group of two paucitubulatinan species of the taxa Chaetonotus and Lepidodermella, possibly due to a similar aberrant but probably not synapomorphic sperm morphology of Dactylopodola and many freshwater dwelling Paucitubulatina (see chapter Gametes). Although Neodasys ciritus was included in that analysis, it is not possible to infer its phylogenetic position because it was used to root the tree. Unfortunately, a non-gastrotrich outgroup taxon was not used. In general, sperm characters are considered to bear a high content of phylogenetic information, but also to be highly homoplastic (Marotta et al. 2005). Another cladistic analysis of 135 phenotypic characters often used for taxonomy of Gastrotricha (e.g., number and arrangement of adhesive tubes, length/ width ratios, cuticular differentiations, shape of “head”, trunk, and “tail”, sensory appendages) provides a slightly different phylogeny for the internal relationships than the hitherto proposed scenarios. According to that analysis, Gastrotricha splits into the sister taxa Neodasys and so called Eutubulata, which comprise all remaining gastrotrich species (Kieneke et  al. 2008a, Fig. 1.53 H). The Eutubulata are characterized by the possession of real adhesive tubes, consisting of a duo-gland complex of two secretory cell types and a tube-shaped cuticular protrusion through which the cells discharge. Sometimes an associated ciliary sensory cell may be present as well (see chapter integument). Within Eutubulata, two monophyla, Macrodasyida sensu stricto (all traditional macrodasyids exclusive of Redudasys fornerise and Marinellina flagellata; characterized by the possession of epidermal glands and pharyngeal pores) and the so-called Abursata (characterized by the absence of the unpaired accessory reproductive frontal and caudal organs) are sister taxa of highest rank. Abursata comprise as sister taxa the monophyletic traditional Paucitubulatina and a monophylum consisting of the only two freshwater macrodasyidan taxa Redudasys and Marinellina. However, a recent analysis of a fragment of the 18S rRNA gene from numerous gastrotrich species has revealed a sister group relationship

 89

of Redudasys and Anandrodasys (together forming the new family Redudasyidae) with a well-supported position within the Macrodasyida and no obvious affiliations to the Paucitubulatina (Todaro et  al. 2012b). A comprehensive overview of the major hypotheses regarding the internal relationships but also the phylogenetic position of Gastrotricha among the Bilateria can be found in the review paper of Balsamo et al. (2010b). A common phenomenon in cladistic analyses using phenotypic (morphological) characters as data are the low recovery values of especially basal (early) nodes when using permutation tests such as the bootstrap to infer the robustness of phylogenetic trees (see node support values of, e.g., Hochberg & Litvaitis 2000, Marotta et  al. 2005, Kieneke et  al. 2008a). Owing to the much higher characters to taxa ratio in studies using DNA sequences as data (several hundreds to thousands of characters), the resolved nodes generally get much higher support values in those analyses (Wägele 2005). However, it has to be stressed that there are also internal nodes with low or just moderate bootstrap support in different 18S-based cladistic analyses of the Gastrotricha (see, e.g., Manylow et al. 2004, Paps & Riutort 2012, Todaro et al. 2012b). The amount of noise in the data set is possibly much more important than bare characters to taxa ratio. Especially in morphological data matrices, there can be many homoplastic character states. Homoplasy may result from, e.g., convergent transformations or parallel character losses. The latter is regarded as one major reason for incongruence between trees generated on the basis of molecular versus morphological data sets (see Bleidorn 2007). The latest chapter in gastrotrich phylogeny reconstruction has recently been started with cladistic analyses that use multiple gene loci for phylogenetic inference. Although very promising, these studies so far comprise only a fraction of gastrotrich taxa and focus on certain groups such as the Thaumastodermatidae (Todaro et  al. 2011a), the Chaetonotidae, or the Dasydytidae (Kånneby et al. 2012, 2013). Such analyses provide interesting new insights into the relationships of, for instance, the speciesrich but probably polyphyletic Chaetonotidae. However, support values for the important basal (early) nodes apparently decrease the more taxa and positions an alignment includes (e.g., compare Kånneby et al. 2012 with Kånneby et al. 2013). Instead of providing detailed descriptions of the different alternative in-group relationships of Gastrotricha, which are far from being settled, we here focus on taxa that show a certain probability to occupy a basal position within their respective superordinate taxon. Pragmatically, “probability” is here equalized with the frequency

90 

 1 Gastrotricha PLATHELMINTHES or GNATHOSTOMULIDA / GNATHIFERA or CYCLONEURALIA or ECDYSOZOA

Xenotrichulidae Polymerurus

“Chaetonotidae”

Dasydytidae

1

PAUCITUBULATINA

2

Muselliferidae

Neogosseidae Proichthydidae Dichaetura 3

Neodasys (MULTITUBULATINA) Dactylopodolidae or Xenodasys/Xenodasyidae or Cephalodasys “Planodasyidae”

“Cephalodasyidae” Lepidodasys Redudasyidae Turbanellidae Thaumastodermatidae

Diplodasyinae Thaumastodermatinae

of a taxon being resolved as a basal lineage among the various phylogenetic analyses. To simplify matters, we in the following refer to genera or even families although in most analyses, molecular as well as morphological, distinct species have been used as terminal taxa. However, we use traditional family names only if their monophyly is not in doubt. Within the Paucitubulatina, the initial split leads to the genus Musellifer (Hochberg & Litvaitis 2000, Todaro et al. 2006a) or, if Musellifer forms the sister group of the arctic marine taxon Diuronotus (Todaro et al. 2005, Leasi & Todaro 2008), to Muselliferidae (Fig. 1.54). Such a basal position of Musellifer/Muselliferidae is supported by the pattern of body musculature of several investiga-

MACRODASYIDA

“Macrodasyidae”

4

Fig. 1.54: Summary tree of the Gastrotricha that shows the known and unresolved internal phylogenetic relationships. The tree was manually constructed and depicts the congruent results between phylogenetic analyses of the past 2 decades (see text for more details). The indicated family names follow the recent systematization of Gastrotricha according to Balsamo et al. (2009), Hummon & Todaro (2010), and Todao et al. (2012) (erection of family Redudasyidae). Lineages that end with a black circle are monophyletic clades; groups marked with gray boxes and quotation marks are polyphyletic or paraphyletic. Dashed lines indicate the uncertain position of the Xenotrichulidae and the putative basal lineages within Macrodasyida (i.e., Dactylopodolidae, Xenodasyidae, or Cephalodasys). Unambiguous autapomorphies of the Gastrotricha and the 3 major internal lineages as follows (according to several authors): (1) external cilia covered with epicuticle, visceral muscular double helix, mechanoreceptive cells with 10 circumciliary microvilli, (2) tenpin-shaped habitus, loss of lateral tubes, incomplete visceral dorsoventral muscles, visceral muscle helix spans to one third of the intestine, bicellular terminal organ with composite filter, aciliar canal cell with convoluted distal lumen, (3) adhesive tubes as non-duo-gland organs, club-shaped mouth tube, loss of vasa deferentia, (4) existence of epidermal glands, existence of pharyngeal pores.

ted paucitubulatinan species (Leasi & Todaro 2008) and recently also by a molecular phylogeny using partial DNA sequences of the nuclear 18S rRNA gene (Kånneby et al. 2014). Besides Musellifer (Muselliferidae), the monophyletic family Xenotrichulidae (genera Draculiciteria, Xenotrichula, and Heteroxenotrichula) is also resolved to occupy a basal position within Paucitubulatina (Todaro et al. 2003, Petrov et al. 2007), sometimes as the result of the second speciation event after the first split that leads to Musellifer/Muselliferidae (Hochberg & Litvaits 2000, Todaro et  al. 2006a). It is also possible that both Muselliferidae and Xenotrichulidae are sister taxa and occupy a basal position within Paucitubulatina (Kieneke et  al. 2008a,

1.5 Phylogeny 

Fig. 1.54). Data of sperm ultrastructure give strong support for such a sister group relationship of Muselliferidae and Xenotrichulidae (Balsamo et  al. 2010b). Patterns of the anatomy of body musculature and of the protonephrida ultrastructure indicate that the next branch within Paucitubulatina leads to the freshwater taxon Polymerurus (Leasi & Todaro 2008, Kieneke & Hochberg 2012). Within the Macrodasyida, species of the genus Dactylopodola, or the family Dactylopodolidae as a whole (Dactylopodola, Dendrodasys, Dendropodola), have frequently been resolved as the earliest branch (Ruppert 1982, Travis 1983, Ruppert 1991, Hochberg & Litvaitis 2000, Todaro et  al. 2003: MP analysis, Zrzavý 2003: Dactylopodola as most basal lineage within whole Gastrotricha, see Zrzavý’s figure 5). However, according to some analyses, they share this putative basal position with others such as Xenodasys and Acanthodasys (Todaro et al. 2003: ML analysis) or with Paradasys, Cephalodasys, Urodasys, and Neodasys in a common clade (e.g., Todaro et al. 2006a). Exceptions from a basal position of Dactylopodolidae provide the studies of Petrov et al. (2007) and Kieneke et al. (2008a). According to nucleotide sequence data of the 18S rRNA gene, Cephalodasys is the most basal lineage within Macrodasyida; however, no sequence of a dactylopodolid species was included (Petrov et  al. 2007). According to phenotypic data (Kieneke et al. 2008a), Xenodasys could also represent the sister group of all remaining lineages of Macrodasyida (Fig. 1.54). Interestingly, such a result is supported by the analysis based on sperm morphology even though the position of Dactylopodola remains unclear (Marotta et  al. 2005). It has to be stressed that so far, the DNA sequence-based studies comprise a fraction of gastrotrich genera only. Hence, we have to keep in mind that assuming the position of a certain taxon, close to the base or highly derived, can be strongly biased by the incomplete taxon sampling. The monophyletic status of most of the gastrotrich genera is not challenged and for many taxa, very specific generic diagnoses with unique characters (putative autapomorphies) have been published, which indicates that genera indeed represent natural units. However, definite tests on monophyly have not been carried out very often (see, e.g., Kieneke 2010 for the genus Thaumastoderma). When we think about monophyly of the traditional families of Gastrotricha, the situation is getting much more difficult. For instance, Turbanellidae and Thaumastodermatidae (Macrodasyida) or Xenotrichulidae, Dasydytidae, and Proichthydidae (Paucitubulatina) turned out to form monophyletic clades according to different analyses (e.g., Hochberg & Litvaitis 2000, 2001a, Todaro et  al. 2006a, 2011a, 2012b, Kieneke et al. 2008a, Leasi & Todaro 2008,

 91

Kieneke & Ostmann 2012). Congruence among results of those analyses indicates that monophyly of the mentioned taxa is highly probable. In contrast, families such as Lepidodasyidae (Macrodasyida) or Chaetonotidae (Paucitubulatina) in most cases turned out to be paraphyletic or polyphyletic assemblages (e.g., Hochberg & Litvaitis 2000, 2001a, Todaro et al. 2006a, Kieneke et al. 2008a, Paps & Riutort 2012, Kånneby et al. 2013). Monophyly of Gastrotricha as a whole has also been questioned (Manylow et al. 2004, see above). However, there are at least two unique characters most likely shared by all members of Gastrotricha, i.e., the cuticular covering of all external cilia (Rieger & Rieger 1977; maybe also the multilayer condition of the epicuticle) and the presence of a visceral muscular double helix encircling the gut tube (Hochberg & Litvaitis 2001a, b). These two characters have recently been confirmed as substantial autapomorphies characterizing the Gastrotricha and supporting its monophyly (Kieneke et al. 2008a). Other potential apomorphies of the Gastrotricha that are mentioned from time to time, for instance, the hermaphroditism (Ax 2003, Balsamo et al. 2010b), or the direct transfer of filiform spermatozoa (Ax 2003), are less robust. First, members of main platyzoan lineages (Plathelminthes, Gnathostomulida) and of the earliest bilaterian branch Acoelomorpha (Acoela+Nemertodermatida) follow a hermaphroditic reproduction, too (see SchmidtRhaesa 2007). Second, occurrence of filiform spermatozoa could also represent a plesiomorphic character within the Bilateria: basal groups such as Nemertodermatida (Boone et al. 2011) and Acoela (Raikova et al. 1998) possess such a sperm type and so do further lineages (but see SchmidtRhaesa 2007 for the proposed ancestral sperm type of the Bilateria and the use of a phylogenetic character “filiform spermatozoon” in general). Inferring the internal relationships of Gastrotricha will make it possible to detect natural groups corresponding to “genera” or “families”. However, the system has not been settled yet (see above), and there are quite a lot of unresolved regions in the phylogenetic tree of Gastrotricha (Fig. 1.54). More extensive searches analyzing different gene loci in combination with testing the resulting topologies with morphological and ecological characters will probably bring us closer to the phylogenetic system of Gastrotricha. Furthermore, the analysis of extensive data sets of morphological characters will be an important source of information for understanding the phylogeny and evolution of this group of animals. In this context, the investigation of morphology and ultrastructure of different organ systems is still a valuable and necessary challenge. As already mentioned in the introduction to this volume (Schmidt-Rhaesa 2013), the phylogenetic position

92 

 1 Gastrotricha

of the phylum Gastrotricha is not yet clearly resolved. Different shared morphological characters of the Gastrotricha and the Cycloneuralia (e.g., a stratified cuticle, a terminal mouth opening, a cuticle-lined myoepithelial sucking pharynx) have been regarded as synapomorphies that support a clade Nemathelminthes (Ahlrichs 1995, Ehlers et al. 1996, Ax 2003). However, none of the expanding number of DNA sequence-based cladistic analyses has ever revealed a sister group relationship between gastrotrichs and cycloneuralians. Instead, studies that using sequences of one to several gene loci and more recently even data of whole transcriptomes (“expressed sequence tags”), repeatedly placed the Gastrotricha into a protostomian “superphylum” known as the Platyzoa (see Giribet 2002, Giribet et  al. 2007). Depending on the particular analysis, within Platyzoa, Gastrotricha may either be allocated as sister group of the Plathelminthes (e.g., Giribet et al. 2000, Dunn et al. 2008) or of the Gnathostomulida (e.g., Hejnol et  al. 2009, Paps et  al. 2009). The last mentioned analysis, however, did not resolve the Platyzoa as a monophyletic group (Paps et al. 2009). A clade Platyzoa was already suggested by Cavalier-Smith (1998) with Gastrotricha and Gnathostomulida as sister taxa united as the socalled Monokonta. As diagnostic characters for Monokonta, Cavalier-Smith (1998) mentions the monociliated epidermis, the simple protonephridia, the compact body organization, and the shared hermaphroditism. However, all these characters more likely represent plesiomorphies among the Bilateria (see, e.g., SchmidtRhaesa 2007) rather than apomorphic features of a clade Monokonta. Also, Zrzavý et  al. (1998) revealed a sister group relationship between Gastrotricha and Gnathostomulida, called the Neotrichozoa, with their combined analysis of molecular and morphological data. Furthermore, Schmidt-Rhaesa et al. (1998) discuss the possibility that Gastrotricha could be placed as the sister group of a large clade comprising all molting animals – the Ecdysozoa (=Cycloneuralia+Panarthropoda). Character states that could support such a grouping are almost the same ones as those used to support the Nemathelminthes (see above): a stratified cuticle, a muscular sucking pharynx, and a terminal mouth opening. However, such a sister group relationship has not been revealed by DNA sequence-based phylogenetic analyses, so far. In summary, there are four potential sister groups for the phylum Gastrotricha: Cycloneuralia, Plathelminthes, Gnathostomulida (or Gnathifera as a whole), or Ecdysozoa (Fig. 1.54). Hence, resolving the phylogenetic position of the Gastrotricha still remains a major task for evolutionary-systematic research.

1.6 Systematics In the following, we give a short overview on the diversity of gastrotrichs. There are some excellent sources to approach gastrotrich systematics. For marine taxa, macrodasyids, and chaetonotids, Todaro & Hummon (2008) provide a useful key to the genera and Hummon & Todaro (2010) give an exhaustive species list with synonyms and some taxonomic remarks. See also Ruppert (1988) for an overview. The most extensive guide to freshwater chaetonotids, though in German language, is from Schwank (1990) for Central European species. Balsamo & Todaro (2002) provide a recent key to the freshwater genera. There are attempts to build a video database (Hummon et al. 2005). Very helpful in terms of taxonomy and systematics of Gastrotricha is the regularly updated webpage “Gastrotricha World Portal” of Antonio Todaro (http:// www.gastrotricha.unimore.it/). We list here 801 species of currently known gastrotrichs. When describing or determining gastrotrichs, it is important to have some sense for potential intraspecific variability. For example, in those chaetonotids with cuticular scales, it is important to know whether the scale patterns are fixed or variable and, if so, in which range. It is assumed that some variability exists (Forneris 1966), but very little is known about the extent of this phenomenon. Amato & Weiss (1982) approached this problem in clones of Lepidodermella squammata and found some differences in the scale pattern as well as some asymmetries. In macrodasyids, some variation, especially in the position and probably also in the number of adhesive tubes, appears to be possible. Recently, molecular approaches, especially the “barcoding” of species by investigation of genes, preferentially the cytochrome oxidase 1, are applied in many animal taxa and often reveal cryptic species among morphologically indistinguishable populations. In gastrotrichs, such approaches are still at the beginning, but will be of great help to distinguish species in the future (see below under Biogeography for an example). In the following, we briefly characterize both orders (Macrodasyida and Chaetonotida), the suborder Paucitubulatina, as well as all families and genera of gastrotrichs. We do not provide full taxonomic diagnoses; however, several characteristic features of each taxon are given. Species are listed, but not further described. Below the order/suborder level, categories other than genus and family are not included here (there are, for instance, some subfamilies and subgenera). Redundant categories are given in parenthesis, e.g., when a family includes only one genus. The systematics followed below is mainly based on Hummon & Todaro (2010)

1.6 Systematics 

for marine taxa and Balsamo et  al. (2009, 2014) for f­reshwater taxa. These two publications are also the source for the species lists, whereas new descriptions since 2009/2010 have been added. See these publications for synonyms, species inquirendae and nomina nuda.

1.6.1 Order Macrodasyida Brunson, 1950 Remane (1924) introduced the name “Macrodasyoidea“ for the increasing number of marine gastrotrichs that were morphologically different from the gastrotrichs known up to that date. This name replaced the term “aberrant gastrotrichs” (in German original “aberrante Gastrotrichen”, see also chapter Phylogeny), which was used for these species (e.g., Remane 1924, 1925a, b, 1926a). Brunson (1950) used the writing “Macrodasyida”, which was suggested to be the preferable name by Chandrasekahara Rao & Clausen (1970) (although these authors still used “Macrodasyoidea”). Macrodasyids are characterized by the presence of adhesive tubes in different positions on the body, usually on the ventral or lateral side of the head region (anterior adhesive tubes, TbA), along the trunk in a lateral, dorsal, or ventral position (lateral, dorsal, or ventral adhesive tubes, TbL, TbD, TbV), and in the posterior end (posterior adhesive tubes, TbP). The pharyngeal lumen has an unpaired dorsal branch and paired ventrolateral branches (inverted Y-shaped). Pharyngeal pores are present in most species. Almost all other characters, the body shape, further appendages, cuticular structures, etc., are quite variable. Nine families are currently recognized: Cephalodasyidae, Dactylopodolidae, Lepidodasyidae, Macrodasyidae, Planodasyidae, Thaumastodermatidae, Turbanellidae, Xenodasyidae, and Redudasyidae.

 93

flat cross section; the body is strap-shaped, sometimes very long and in some species with a delimited head; no cuticular structures are present (“naked” cuticle); the ventral cilia form paired longitudinal bands, which may join anteriorly and posteriorly; the TbP are along the blunt end of trunk; the pharyngeal pores are close to pharynx-intestine transition; the radial pharyngeal musculature is striated; circular muscles are present in lateral body wall; and Y-cells are absent. Genus Cephalodasys Remane, 1926 (Fig. 1.55) This genus includes Psammodasys (see d’Hondt 1974, Hummon & Todaro 2010). The body is elongate, and a head region is slightly delimited. The TbA are on fleshy “hands” (Fig. 1.7 I). The ventral ciliation is on the entire

The six genera Cephalodasys, Dolichodasys, Megadasys, Mesodasys, Paradasys, and Pleurodasys were united as Cephalodasyidae by Hummon & Todaro (2010); formerly, they belonged to the family Lepidodasyidae. Most recently, Guidi et al. (2014) discovered different characters of the reproductive system and the spermatozoa shared between Megadasys and Crasiella and therefore assigned Megadasys to the family Planodasyidae (see below). Some characters of species in Cephalodasyidae, especially in comparison to Lepidodasys, are the following (from Hummon & Todaro 2010): a more or less

100 µm

1.6.1.1 Family Cephalodasyidae Hummon & Todaro, 2010

Fig. 1.55: Horizontal view of Cephalodasys sp. (Macrodasyida, Cephalodasyidae) from sublittoral calcareous sand around Lee Stocking Island, Bahamas. Note the characteristic neck constriction at the level of the anterior adhesive tubes (triangle). DIC image.

94 

 1 Gastrotricha

ventral surface; it is dense in the anterior region and fades toward the posterior. Twelve species are known from the eastern North Atlantic coast, the Baltic Sea, the Mediterranean Sea, the Red Sea, the Caribbean, and the Indian Ocean around India: C. cambriensis (Boaden, 1963), C. caudatus Chandrasekhara Rao, 1981, C. dolichosomus Hummon, 2011, C. hadrosomus Hummon, Todaro & Tongiorgi, 1993, C. littoralis Renaud-Debyser, 1964, C. maximus Remane, 1926, C. miniceraus Hummon, 1974, C. pacificus Schmidt, 1974, C. palavensis Fize, 1963, C. saegailus Hummon, 2011, C. swedmarki Hummon, 2008, and C. turbanelloides (Boaden, 1960).

Genus Mesodasys Remane, 1951 (Fig. 1.56) No head region is delimited. TbA are several to many; in most species, they are in a traverse row or arc. Many short TbL are present, and sometimes, additional ventral or even dorsal (M. ischiensis, see Hummon et  al. 1993) adhesive tubes are present. TbP are along the posterior margin of the slender terminal end or on a broad caudal plate. The pharyngeal pores are close to the pharynx-intestine junction. A further peculiarity of Mesodasys are the posteriorly directed sperm ducts, which directly discharge into the caudal organ (e.g., Ferraguti & Balsamo 1994, Fregni et al. 1999). Eight species were described from the North Sea and western Atlantic, the eastern Atlantic (Carolina), the Mediterranean Sea, the Caribbean and India: M. adenotubulatus Hummon, Todaro & Tongiorgi, 1993, M. britanicus Hummon, 2008, M. hexapodus Chandrasekhara Rao & Ganapati, 1968, M. ischiensis Hummon, Todaro & Tongiorgi, 1993, M. laticaudatus Remane, 1951, M. littoralis Remane, 1951, M. rupperti Hummon, 2008, and M. saddlebackensis Hummon, 2010.

tes 0.5 mm

Genus Dolichodasys Gagné, 1977 Dolichodasys is characterized by the following characters (after Ruppert & Shaw 1977). It has a large body size (up to 2.7 mm). One or two fused TbA are on each body side, TbL are absent or only present as indistinct adhesive papillae. TbP patterns vary from one single to several adhesive tubes, with distinct changes during development (Gagné 1977). The ventral ciliation is in four bands in the anterior end and two bands along the trunk. The small pharyngeal pores are close to the pharynx-intestine junction. Spermatozoa are aberrant, commaform, and probably aflagellate. Three species are known from the eastern and western coasts of the Atlantic: D. carolinensis Ruppert & Shaw, 1977, D. delicatus Ruppert & Shaw, 1977, and D. elongatus Gagné, 1977.

vd co

Fig. 1.56: Ventral view of Mesodasys cf. laticaudatus (Macrodasyida, Cephalodasyidae) from sublittoral sand of the Ria de Ferrol, Spain. Abbreviations: co, caudal organ; tes, testis; vd, conspicuous swelling of the vas deferens. BF image.

Genus Paradasys Remane, 1934 (Fig. 1.57) In most species, a strong ciliation is present around the head region, and the ventral ciliation extends in two ventral bands along the trunk. There are one or two closely located TbA; TbL are absent. Several TbP are either along the posterior margin or on paired caudal feet. A ciliated muzzle is around the small mouth opening, the pharyngeal pores are close to the pharynxintestine junction. Apart from two species (Paradasys lineatus and P. littoralis), no traces of male gonads have been found in the remaining ones so far. This could indicate a parthenogenetic reproduction (see Wilke 1954). Seven species with wide, but scattered distribution (Baltic Sea, western North Atlantic, India, Japan, Galapagos) are known: P. bilobocaudus Hummon, 2008, P. hexa-

1.6 Systematics 

 95

lateral row. At about the middle level of the trunk, an appendage composed of three long adhesive tubes is present. TbP are along the rounded posterior edge.

1.6.1.2 Family Dactylopodolidae Strand, 1929 This family contains three genera, which all have in common a more or less tenpin-shaped body form, a well-delimited head, and the presence of cross-striated muscles. The pharyngeal pores are close to pharynxintestine junction. Three genera, Dactylopodola, Dendrodasys, and Dendropodola are included in this family. mg

ov vc?

100 µm Fig. 1.57: Horizontal view of Paradasys subterraneus (Macrodasyida, Cephalodasyidae) from the beach slope of Schillig, Germany. Abbreviations: mg, midgut; ov, ovary; vc?, presumptive vitellocytes. DIC image.

dactylus Karling, 1954, P. lineatus Chandrasekhara Rao, 1980, P. littoralis Chandrasekhara Rao & Ganapati, 1968, P. nipponensis Sudzuki, 1976, P. pacificus Schmidt, 1974, and P. subterraneus Remane, 1934. Genus Pleurodasys Remane, 1927 (Fig. 1.58 A–C) There is probably only one species in this genus, P. helgolandicus Remane, 1927 (see Hummon & Todaro 2010), which was found in the western Atlantic. Most conspicuous are the unique “drumstick-like” organs on the ventral side behind the head. These were investigated by Marotta et al. (2008) and may be gravity receptors (see also chapter Sensory Structures). The head is distinctly separated from trunk. TbA originate from short fleshy hands; TbL are in a lateral and a dorso-

Genus Dactylopodola Strand, 1929 (Fig. 1.59) Originally named Dactylopodella (Remane 1926a), the genus had to be renamed because the name was preoccupied. Strand’s (1929) proposition of Dactylopodola was 2 months earlier than Remane’s (1929) Dactylopodalia and therefore is the valid genus name (Blake 1933). Apart from the family characters, Dactylopodola species have bilateral posterior feet carrying the TbP. Most species have few TbL. Some species have eyes, and in some species, the TbA are on fleshy hands. See Hummon & Todaro (2010) for synonyms and Von und zu Gilsa et al. (2014) for an overview and an updated determination key to species. Nine species are known from scattered locations in the entire world, ranging from Australia to India, the Caribbean, and the western Atlantic: D. australiensis Hochberg, 2003, D. axi Von und zu Gilsa, Kieneke, Hochberg & Schmidt-Rhaesa, 2014, D. baltica (Remane, 1926b), D. cornuta (Swedmark, 1956), D. deminuitubulata Von und zu Gilsa, Kieneke, Hochberg & Schmidt-Rhaesa, 2014, D. indica (Chandrasekhara Rao & Ganapati, 1968), D. mesotyphle Hummon, Todaro, Tongiorgi & Balsamo, 1998, D. roscovita (Swedmark, 1967), and D. typhle (Remane, 1927). Genus Dendrodasys Wilke, 1954 (Fig. 1.60) The species have a well-delimited head; in some species, with lateral extensions. There is only one pair of solid TbA. TbL are lacking. The caudal end is elongated and terminally bifurcated. One large adhesive tube is on the basis of each caudal branch, and two smaller adhesive tubes are on the caudal end of each branch. Very conspicuous is the presence of cilia in the pharynx and in the hindgut. These are present at least in D. affinis and D. gracilis (Wilke 1954); they were not observed in D. pacificus (Schmidt 1974). The pharyngeal cilia seem to function as a kind of weir to filter particles when water is extruded through the pharyngeal pores. A pair of testes or a single testis are present.

96 

 1 Gastrotricha

A

B

gr 50 µm

C

100 µm

mo 50 µm

Fig. 1.58: Pleurodasys helgolandicus (Macrodasyida, Cephalodasyidae) from the intertidal beach of Saint-Lunaire, France. (A) Habitus of a subadult specimen in dorsal view. (B) Dorsal view of the anterior end with the conspicuous gravireceptor organs. (C) Ventral view of the anterior end. Note the hand-like arranged anterior adhesive tubes (triangles). Abbreviations: gr, external portion of gravireceptor organs, mo, mouth opening. (A) DIC image. (B and C) BF images.

Lee, Chang & Kim, 2014, D. gracilis Wilke, 1954, D. pacificus Schmidt, 1974, D. ponticus Valkanov, 1957, and D. rubomarinus Hummon, 2011. pp

mo 100 µm Fig. 1.59: Ventral view of Dactylopodola typhle (Macrodasyida, Dactylopodolidae) from the intertidal beach of Saint-Lunaire, France. Abbreviations: mo, mature egg; pp, pharyngeal pores at the level of the neck constriction. BF image.

Six species are known from the Mediterranean Sea, the Black Sea, the Red Sea, the East China Sea, the Sea of Japan, and Galapagos: D. affinis Wilke, 1954, D. duplus

Genus Dendropodola Hummon, Todaro & Tongiorgi, 1993 One species, D. transitionalis Hummon (Todaro & Tongiorgi, 1993) was described in this genus. It resembles Dactylopodola with a tenpin shape and a well-delimited head. The caudal end is elongated, a pair of TbP is present at the base of the extension, and another pair at its tip. There are three pairs of TbL along the trunk and one pair of TbA. The pharynx is relatively long, the pharyngeal pores are close to the pharynx-intestine junction and behind the head constriction (see Hummon et al. 1993 for more data).

1.6.1.3 [Family Lepidodasyidae Remane, 1927] Genus Lepidodasys Remane, 1926 (Fig. 1.61) After removal of six genera to Cephalodasyidae (see above), Lepidodasyidae contains only one genus, Lepidodasys, with the following characteristics (especially in comparison to Cephalodasyidae) (from Hummon & Todaro 2010): The specimens are circular in cross section, the body is strap-shaped, without a delimited head. The body is covered with conspicuous scales (Fig. 1.4 F). Pharyngeal pores absent, a very unique feature

1.6 Systematics 

 97

A TbA

50 µm

B TbP

200 µm

50 µm Fig. 1.60: Dendrodasys cf. gracilis (Macrodasyida, Dactylopodolidae) from the sublittoral of Ria de Ferrol, Spain. Note that the specimen is slightly twisted. (A) More ventral view. (B) More dorsal view. Abbreviations: TbA, anterior adhesive tubes; TbP, posterior adhesive tubes. (A and B) BF images.

among macrodasyids. The radial pharyngeal musculature is not striated. TbP are present along the blunt end of trunk. Y-cells are present; they contain myofibrils. Circular muscles in the body wall were claimed to be absent (Hummon & Todaro 2010), but were recently shown with phalloidin staining (Hochberg et  al. 2013). Lepidodasys species are slowly gliding, their scale pattern is the most important taxonomic feature (Hochberg & Atherton 2011, Lee & Chang 2011, Hochberg et al. 2013). Nine species are known from the North Atlantic, the Caribbean, the Mediterranean Sea, and the western Pacific around Korea (see Hochberg et al. 2013). A key to species is provided by Lee & Chang (2011). Species are: L. arcolepis Clausen, 2004, L. castoroides Clausen, 2004, L. laeviacus Lee & Chang, 2011, L. ligni Hochberg, Atherton & Gross, 2013, L. martini Remane, 1926, L. platyurus Remane, 1927, L. tsushimaenensis Lee & Chang, 2011, L. unicarenatus Balsamo, Fregni & Tongiorgi, 1994, and L. worsaae Hochberg & Atherton, 2011.

Fig. 1.61: Lepidodasys sp. (Macrodasyida, Lepidodasyidae) from coral sand of Lee Stocking Island, Bahamas. Note the characteristic keeled scales on the surface. DIC image.

1.6.1.4 Family Macrodasyidae Remane, 1926 This family unites the two genera Macrodasys and Urodasys. Both are strap-shaped gastrotrichs with anterior, lateral, and posterior adhesive tubes, a medium-sized mouth opening, pharyngeal pores significantly anterior of the pharynx-intestine junction and usually a ventral ciliation covering the entire ventral side. The posterior end terminates in many species in an unpaired tail. Although this tail is moderately long in Macrodasys, it is very long in Urodasys species. Genus Macrodasys Remane, 1924 (Fig. 1.62) The TbA are usually in bilateral rows, TbL occur along the entire trunk and TbP are present in various distribution and density on the posterior region of the animals. A clear-cut delimitation between TbL and TbP is often not possible. Species of Macrodasys generally possess welldeveloped frontal and caudal organs. This is a species-rich genus with 36 species described to date; they occur in many places in the world oceans: M. achradocytalis Evans, 1994, M. acrosorus Hummon & Todaro, 2009, M. affinis Remane, 1936, M. africanus Remane, 1950,

98 

 1 Gastrotricha

M. ancocytalis Evans, 1994, M. andamanensis Chandrasekhara Rao, 1993, M. balticus Roszczak, 1939, M. blysocytalis Evans, 1994, M. buddenbrocki Remane, 1924, M. caudatus Remane, 1927, M. celticus Hummon, 2008, M. cephalatus Remane, 1927, M. cunctatus Wieser, 1957, M. deltocytalis Evans, 1994, M. digronus Hummon & Todaro, 2009, M. dolichocytalis Evans, 1994, M. fornerisae Todaro & Rocha, 2004, M. gerlachi Papi, 1957, M. gylius Hummon, 2010, M. hexadactylis Chandrasekhara Rao, 1970, M. imbricatus Hummon, 2011, M. lakshadweepensis Hummon, 2008, M. macrurus Hummon, 2011, M. meristocytalis Evans, 1994, M. neapolitanus Papi, 1957, M. nigrocellus Hummon, 2011,

A

M. nobskaensis Hummon, 2008, M. ommatus Todaro & Leasi, 2013, M. pacificus Schmidt, 1974, M. plurosorus Hummon, 2008, M. remanei Boaden, 1963, M. scleracrus Hummon, 2011, M. stenocytalis Evans, 1994, M. syringodes Hummon, 2010, M. thuscus Luporini, Magagnini & Tongiorgi, 1973, and M. waltairensis Chandrasekhara Rao & Ganapati, 1968. Genus Urodasys Remane, 1926 (Fig. 1.63) Species in this genus resemble Macrodasys in many respects, but are clearly recognizable by the extremely long

B

mo

co

Fig. 1.62: Two undetermined specimens of Macrodasys (Macrodasyida, Macrodasyidae) from the Caribbean Sea. (A) Specimen from the sublittoral around Little Caiman Island, horizontal view. (B) Individual from coral sand of Lee Stocking Island, Bahamas. The tail with the posterior adhesive tubes is in focus. Abbreviations: co, caudal organ; mo, mature eggs; TbP, posterior adhesive tubes. (A and B) DIC images.

100 µm

100 µm

TbP

100 µm

ta

Fig. 1.63: Urodasys sp. (Macrodasyida, Macrodasyidae) from San Salvador Island, Bahamas. The tail with the posterior adhesive tubes is strongly contracted. Abbreviations: ta, tail. DIC image.

1.6 Systematics 

 99

and contractile tail. Some species show peculiarities in their reproductive anatomy or biology, such as the presence of stylets (Fig. 1.36 G, H) or vivipary (Fig. 1.50 B). Fifteen species are known from the western and eastern North Atlantic, the Baltic Sea, the Mediterranean Sea, the Caribbean, the Maledives, and French Polynesia: U. acanthostylis Fregni, Tongiorgi & Faienza, 1998, U. anorektoxys Todaro, Bernhard & Hummon, 2000, U. apuliensis Fregni, Faienza, Grimaldi, Tongiorgi & Balsamo, 1999, U. bucinastylis Fregni, Faienza, Grimaldi, Tongiorgi & Balsamo, 1999, U. calicostylis Schoepfer-Sterrer, 1974, U. cornustylis SchoepferSterrer, 1974, U. elongatus Renaud-Mornant, 1969, U. mirabilis Remane, 1926, U. nodostylis Schoepfer-Sterrer, 1974, U. poculostylis Atherton, 2014, U. remostylis SchoepferSterrer, 1974, U. spirostylis Schoepfer-Sterrer, 1974, U. toxistylis Hummon, 2011, U. uncinostylis Fregni, Tongiorgi & Faienza, 1998, and U. viviparus Wilke, 1954. 1.6.1.5 Family Planodasyidae Chandrasekhara Rao & Clausen, 1970 When Clausen (1968) introduced the genus Crasiella, he was not sure about its placement among known gastrotrichs. After the description of Planodasys marginalis (Chandrasekhara Rao & Clausen 1970), the family Planodasyidae was introduced to unite these two genera. Species of both genera possess caudal feet with the TbP. They also have a “corona”, i.e., a ring of cuticular structures around the mouth opening and a buccal cavity in common. Recently, Guidi et  al. (2014) assigned a third genus, Megadasys (formerly included within Cephalodasyidae, see above) to Planodasyidae. An amended diagnosis of Planodasyidae, now comprising the three genera Crasiella, Megadasys, and Planodasys, is presented, but shared characters between Megadasys and Crasiella are mainly ultrastructural features of their spermatozoa (see Guidi et al. 2014). Genus Crasiella Clausen, 1968 (Fig. 1.64) Species have small, sometimes hardly distinguishable caudal feet carrying the TbP. A head is delimited in some species, but not in others. The ventral side is covered either entirely by cilia or cilia are in paired longitudinal rows. The mouth opening is surrounded by a ring of hook-like structures, and a buccal cavity is present. The pharyngeal pores are in the posterior part of the pharynx, close to the pharynx-intestine junction. The TbA are in some species in continuity of the ventrolateral TbL (e.g., C. azorensis), but in others clearly separated and form paired diagonal rows (e.g., C. oceanica, see Hummon 2008a). The TbL are scarce or dense, they may

100 µm Fig. 1.64: Crasiella cf. diplura (Macrodasyida, Planodasyidae) from sublittoral sand of the island Elba, Italy. BF image.

be of unequal size. In several species, the posterior part of the intestine has an S-shaped course. Eight species have been described from the western North Atlantic, the Caribbean Sea, the Indian Sea, the East China Sea, and the Pacific around Galapagos: C. azorensis Hummon, 2008, C. clauseni Lee & Chang, 2012, C. diplura Clausen, 1968, C. fonseci Hochberg, 2014, C. indica Chandrasekhara Rao, 1981, C. oceanica d’Hondt, 1974, C. pacifica Schmidt, 1974, and C. skaia Hummon, 2010. A determination key to species is provided in Lee & Chang (2012). Genus Megadasys Schmidt, 1974 (Fig. 1.65) Two species were described in the same year under different genus names, and because of a slightly earlier publication, the name Megadasys (Schmidt 1974) is valid and Thiodasys (Boaden 1974) is not (see Kisielewski 1987b). Specimens are very long and slender (up to >3 mm). They lack TbA; short TbL are numerous along the trunk and several TbP are on a caudal lobe along the posterior edge. The pharyngeal pores are close to pharynx-intestine junction, with a probable more anterior position in M. pacificus (see Schmidt 1974) (see Kisielewski 1987b for more information on the genus).

100 

 1 Gastrotricha

0.5 mm eg

co

Fig. 1.65: Megadasys minor (Macrodasyida, Planodasyidae) from sublittoral sand of the island Elba, Italy. Abbreviations: co, caudal organ; eg, epidermal glands. BF image.

Three species are known from the western Atlantic coasts and Galapagos: M. minor Kisielewski, 1987, M. pacificus Schmidt, 1974, and M. sterreri (Boaden, 1974). Genus Planodasys Chandrasekhara Rao & Clausen, 1970 Specimens are quite large, up to 0.8 mm in P. littoralis and up to 1.5 mm in P. marginalis. The caudal feet are clearly demarcated and appear as lobe-like structures covered with TbP. The TbA are arranged in a medially interrupted crescent. A head region is not delimited. TbL are numerous in P. marginalis (120–140), whereas P. littoralis has fewer (about 30) (see Chandrasekhara Rao & Clausen 1970 and Chandrasekhara Rao 1993). There are two species, both from the Bay of Bengal in the Indian Ocean: P. littoralis Chandrasekhara Rao, 1993, and P. marginalis Chandrasekhara Rao & Clausen, 1970.

1.6.1.6 Family Thaumastodermatidae Remane, 1927 Species in this large family are very diverse. Ruppert (1978b) has given an amended diagnosis including the following

characters: usually caudal feet are present, the body is often covered with cuticular structures, TbL are often in a ventrolateral position, epidermal cells are multiciliary, radial parynx musculature is reduced, pharyngeal pores are small, and at the end of the pharynx, Y-cells contain myofibrils, the caudal organ is well developed and close to the anus, oviduct is present. Ruppert (1978b) proposed two subfamilies, Thaumastodermatinae (including genera Thaumastoderma, Tetranchyroderma, Oregodasys, Pseudostomella, and Ptychostomella) and Diplodasyinae (including genera Acanthodasys and Diplodasys), with Hemidasys being of uncertain assignment. The most conspicuous difference is that the left side of the gonad is reduced in Thaumastodermatinae. This is most obvious in the testis, which is only present on the right side of animals (Fig. 1.36 C), whereas the unpaired (right) ovary is often more or less central in position. Diplodasyinae retain the paired nature of the gonad. Eight genera (Acanthodasys, Diplodasys, Hemidasys, Oregodasys, Pseudostomella, Ptychostomella, Tetranchyroderma, and Thaumastoderma) belong to this family. Genus Acanthodasys Remane, 1927 (Fig. 1.66 A–C) Species are slender and covered with cuticular spines, which are often called uniancres. These are spined scales. Unspined scales are additionally present in some species. Adhesive tubes are very variable. TbA are in diverse arrangements on the anterior end; TbL can be in dorsolateral or ventrolateral distribution or completely lacking. The TbP are along the posterior margin or on small caudal feet. Several new species names were mentioned by Ruppert (1978b), but full species descriptions were not provided. Hence, each of these names is considered as nomen nudum (see Hummon & Todaro 2010). Clausen (2004a) gives a key to the five species known at that time. Twelve species are known from both sides of the North Atlantic, the Caribbean, the Mediterranean Sea, the Maledives, the seas around Korea and India: A. aculeatus Remane, 1927, A. algarvensis Hummon, 2008, A. arcassonensis Kisielewski, 1987, A. caribbeanensis Hochberg & Atherton, 2010, A. carolinensis Hummon, 2008, A. comtus Lee, 2012, A. ericinus Lee, 2012, A. fibrosus Clausen, 2004, A. flabellicaudus Hummon & Todaro, 2009, A. lineatus Clausen, 2000, A. paurocactus Atherton & Hochberg, 2012, and A. silvulus Evans, 1992. Genus Diplodasys Remane, 1927 (Fig. 1.67 A, B) This genus includes flattened specimens with a dorsal covering of flat scales (Figs. 1.4 A and 1.67 B). Scales are also present on the ventral side; the ventral distribution of cilia is often interrupted and appears in isolated patches.

1.6 Systematics 

A

 101

B

ph

tes

50 µm

C

50 µm

ov

100 µm

Especially the dorsal scales may exhibit different patterns of ornamentation such as depressions, pores, or ridges. A head region is usually visible, but this may depend on the state of contraction of the animals (Kieneke et al. 2013b). Very characteristic is the presence of a row of strong spines along the lateral margin of the trunk (Fig. 1.4 I). The TbA are in varying number and arrangement on the ventral side of the head region. TbL are present in paired ventrolateral rows, and TbP are present on either caudal feet or on the posterior margin. Ten species have been described from diverse locations in the world oceans (eastern North Atlantic, Mediterranean Sea, Caribbean, Galapagos, Indian Ocean): D. ankeli Wilke, 1954, D. caudatus Kisielewski, 1987, D. meloriae Todaro, Balsamo & Tongiorgi, 1992, D. minor Remane, 1936, D. pacificus Schmidt, 1974, D. platydasyoides Remane, 1927, D. remanei Chandrasekhara Rao & Ganapati, 1968, D. rothei Kieneke, Narkus, Hochberg & Schmidt-Rhaesa, 2013, D. sanctimariae Hummon & Todaro, 2009, and D. swedmarki Kisielewski, 1987.

Fig. 1.66: Acanthodasys aculeatus (Macrodasyida, Thaumastodermatidae) from the shallow sublittoral of Elba, Italy. (A) Habitus in horizontal view. (B) Dorsal view of the anterior end covered with cuticular uniancres and small scales. (C) Ventral view of the posterior end with the caudal pedicles and posterior adhesive tubes. Abbreviations: ov, ovary; ph, pharynx; tes, testes. (A–C) BF images.

Genus Hemidasys Claparéde, 1867 Hemidasys agaso was the second species of Macrodasyida to be described (Claparéde 1867). It was found in muddy sediments of the port of Naples (Mediterranean Sea), both free living and epizoic on the polychaete Neanthes caudatus (Claparéde 1867). Since then, it has not been found again, despite repeated attempts (see Hummon & Todaro 2010). Hemidasys agaso, the single species in the genus, has a narrow mouth opening, and a large buccal cavity being covered by something like an oral hood. Ventral cilia are present only in the anterior body region and few cuticular plates are found in the region of the male genital pore. TbA are in one row, TbL are few. Genus Oregodasys Hummon, 2008 (Fig. 1.68 A, B) Remane (1927a) introduced the genus Platydasys, but because this name was preoccupied, Hummon (2008a) renamed it as Oregodasys. Specimens are very flat and broad, and some can be quite large (up to 800 µm in

102 

 1 Gastrotricha

A

B

lsp

50 µm

TbVL

100 µm

A

Fig. 1.67: (A) Diplodasys cf. meloriae (Macrodasyida, Thaumastodermatidae) from calcareous sand of Lee Stocking Island, Bahamas. Ventral view with focus on adhesive tubes. (B) Close-up of the dorsal scales of D. rothei from Norman’s Pond Cay, Bahamas. Abbreviations: lsp, lateral spines, TbVL ventrolateral adhesive tubes. (A and B) DIC images. (Micrograph in B was from Kieneke et al. 2013b, with kind permission by Verlag Dr. Friedrich Pfeil.)

B

pp

mg

TbP 100 µm

50 µm

P. ruber and P. tentaculatus, Swedmark 1956). The broad mouth opening is directed anteroventrally. Eyes are present in some species (O. ocellatus, O. norenburgi).

Fig. 1.68: (A) Oregodasys cirratus (Macrodasyida, Thaumastodermatidae) from sand out of a submarine cave on Tenerife, Canary Islands. Dorsal view showing the numerous secretory papillae. (B) Subadult specimen of Oregodasys caymanensis from Little Cayman Island. Abbreviations: mg, midgut; pp, pharyngeal pores; TbP, posterior adhesive tubes. (A and B) DIC images. (Micrograph in A was from Rothe & Schmidt-Rhaesa 2010, with kind permission by Verlag Dr. Friedrich Pfeil.)

The body is covered by papillae. TbA are present along the posterior margin of the mouth opening and in irregular arrangement posterior of it. TbL are

1.6 Systematics 

present in a lateral and a ventral row, and TbP are either along the posterior margin or on small caudal feet. In six species, cirri (=compound cilia) are present. These cirri are present in the posterior part or, in O. cirratus, in a row along the entire trunk. See Hochberg (2010a) and Rothe & Schmidt-Rhaesa (2010) for more information. Fifteeen species have been described to date with distribution in the eastern North Atlantic including the Canary Islands, the Mediterranean, the Caribbean, Galapagos, and Japan: O. caymanensis Hochberg, Atherton & Kieneke, 2014, O. cirratus Rothe & Schmidt-Rhaesa, 2010, O. itoi (Chang, Kubota & Shirayama, 2002), O. katharinae Hochberg, 2010, O. kurnowensis Hummon, 2008, O. mastigurus (Clausen, 1965), O. maximus (Remane, 1927), O. norenburgi Hochberg, 2010, O. ocellatus (Clausen, 1965), O. pacificus (Schmidt, 1974), O. phacellatus (Clausen, 1965), O. rarus (Forneris, 1961), O. ruber (Swedmark, 1956), O. styliferus (Boaden, 1965), O. tentaculatus (Swedmark, 1956). Genus Pseudostomella Swedmark, 1956 (Fig. 1.69 A, B) Species in the genus Pseudostomella are very distinctive because they have very conspicuous anterior extensions, the pre-buccal palps. These serve as grasping structures. Papillae of different size can be present on these palps and dorsal and ventral of the mouth rim. The mouth opening is very broad. TbA are present along the ventral side of the mouth opening. TbL are along the lateral sides of the animals; additionally, a group of ventral adhesive tubes is present in some species. The TbP are on small caudal feet. Cuticular structures in the form of triancres, tetrancres, and pentancres, i.e., with three, four, or five tips are present, the triancres can be in the form of scaled triancres (or feathered triancres) (see Ruppert 1970, Hochberg 2002). Keys to species were provided by Ruppert (1970), Lee & Chang (2002), Hochberg (2002), Clausen (2004b), and Todaro (2012). Sixteen species are currently known from diverse locations of the North Atlantic, the Mediterranean, the Indian Ocean, Australia, Malaysia, and the western Pacific around Korea: P. andamanica Chandrasekhara Rao, 1993, P. cataphracta Ruppert, 1970, P. cheraensis Pryialakshmi, Menon & Todaro, 2007, P. dolichopoda Todaro, 2012, P. etrusca Hummon, Todaro & Tongiorgi, 1993, P. faroensis Clausen, 2004, P. indica Chandrasekhara Rao, 1970, P. klauserae Hochberg, 2002, P. koreana Lee & Chang, 2002, P. longifurca Lee & Chang, 2002, P. malayica Renaud-Mornant, 1967, P. megapalpator Hochberg, 2002, P. plumosa Ruppert, 1970, P. roscovita Swedmark, 1956, P. squamalongispina Araujo, Balsamo & Garraffoni, 2013, and P. triancra Hummon, 2008.

 103

Genus Ptychostomella Remane, 1926 Species in this genus are short and straight. The mouth opening is large, and the anterior rim is folded. Eyes are present in one species (P. ommatophora). Some species have tentacles. The TbA are in an arc, TbL are concentrated in groups and may be lateroventral or ventral. The posterior end forms lateroterminal tips of caudal feet, both covered with TbP. Cuticular structures are either absent (most species) or present on the dorsal side. However, the characteristic hook-shaped ancres that occur in three other genera of the subfamily Thaumastodermatinae (Pseudostomella, Tetranchyroderma, Thaumastoderma) are always absent in Ptychostomella. See Lee & Chang (2003) for a comparison of species. Todaro (2013) provides a key to species. Thirteen species are currently known from the eastern North Atlantic, the Baltic Sea, the Mediterranean, and the Northwest Pacific (Korea): P. bergensis Clausen, 1996, P. brachycephala (Levi, 1954), P. helana Roszczak, 1939, P. higginsi Clausen, 2004, P. jejuensis Lee, Hwang &

A

B pa

50 µm

50 µm

Fig. 1.69: Pseudostomella roscovita (Macrodasyida, Thaumastodermatidae) from the intertidal beach in Saint-Lunaire, France. (A) Horizontal view. (B) Dorsal view. Abbreviation: pa, prebuccal apparatus. (A and B) BF images.

104 

 1 Gastrotricha

Chang, 2009, P. lamelliphora Todaro, 2013, P. lepidota Clausen, 2000, P. mediterranea Remane, 1927, P. ommatophora Remane, 1927, P. orientalis Lee & Chang, 2003, P. papillata Lee & Chang, 2003, P. pectinata Remane, 1926, and P. tyrrhenica Hummon, Todaro & Tongiorgi, 1993. Genus Tetranchyroderma Remane, 1926 (Fig. 1.70) This species-rich genus includes stout to strap-shaped specimens with a covering of cuticular structures in the form of triancres, tetrancres, or pentancres; pentancres are the

100 µm Fig. 1.70: Tetranchyroderma sp. (Macrodasyida, Thaumastodermatidae) from an intertidal beach close to Ribadeo, Spain. BF image.

most abundant structures (Fig. 1.4 D, E). Some species do not have a complete covering by cuticular structures. Usually, only one type of cuticular structures is present, but some species have a mixture of different types. Cephalic tentacles are present in few species. The mouth opening is very broad, and the anterior mouth rim forms a characteristic oral hood. Adhesive tubes occur in the usual groups, TbA, TbL, and TbP; ventral and dorsal adhesive tubes on the trunk are present in some species. Most, but not all species have caudal feet, the TbP are present on these feet, and in some species, additionally between the feet. Todaro (2002) provides a helpful key to species of the genus. Currently, 82 species have been described from many locations in the world oceans: T. aapton Dal Zotto, Ghiviriga & Todaro, 2010, T. adeleae Hochberg, 2009, T. aethesbregmum Lee & Chang, 2012, T. anisoankyrum Lee 2012, T. anomalopsum Hummon, Todaro, Balsamo & Tongiorgi, 1996, T. antenniphorum Hummon & Todaro, 2010, T. aphenothigmum Hummon, Todaro, Tongiorgi & Balsamo, 1998, T. apum Remane, 1927, T. arcticum Clausen, 1999, T. australiense Nicholas & Todaro, 2006, T. boadeni Schrom, in Riedl 1970, T. boreale Clausen, 2000, T. bronchostylus Atherton & Hochberg, 2012, T. bulbosum Clausen, 2000, T. bunti (Thane-Fenchel, 1970), T. canariense Todaro, Ancona, Marzano, D’Addabbo & De Zio Grimaldi, 2003, T. cirrophorum Lévi, 1950, T. coeliopodium Boaden, 1963, T. copicirratum Hummon, 2010, T. corallium Hummon, 2011, T. coreensis Lee, 2012, T. dendricum Saito, 1937, T. dragescoi Swedmark, 1967, T. enallosum Hummon, 1977, T. esarabdophorum Tongiorgi & Balsamo, 1984, T. faroense Clausen, 2004, T. gausancrum Hummon, 2008, T. gracilium Chang, Lee & Clausen, 1998, T. heterotentaculatum Chang & Lee, 2001, T. heterotubulatum Hummon, Todaro & Tongiorgi, 1993, T. hirtum Luporini, Magagnini & Tongiorgi, 1973, T. hoonsooi Chang & Lee, 2001, T. hyponiglarum Hummon & Todaro, 2009, T. hypopsilancrum Hummon, Todaro & Tongiorgi, 1993, T. hystrix Remane, 1926, T. inaequitubulatum Todaro, Balsamo & Tongiorgi, 2002, T. indicum Chandrasekhara Rao & Ganapati, 1968, T. insolitum Lee & Chang, 2012, T. insulare Balsamo, Fregni & Tongiorgi, 1994, T. interstitiale Hummon, 2008, T. kontosomum Hummon, Todaro, Balsamo & Tongiorgi, 1996, T. korynetum Hummon & Todaro, 2009, T. lameshurense Hummon, 2008, T. littorale Chandrasekhara Rao, 1981, T. longipedum Hummon, 2008, T. mainensis Hummon & Guadiz, 2009, T. massilense Swedmark, 1956, T. megabilubulatum Lee & Chang, 2012, T. megastomum (Remane, 1927), T. monokerosum Lee & Chang, 2007, T. multicirratum Lee & Chang, 2007, T. norvegicum Clausen, 1996, T. oblongum Lee, 2012, T. oligopentacrum Hummon & Todaro, 2009, T. pachysomum Hummon, Todaro & Tongiorgi, 1993, T. pacificum Schmidt, 1974, T. papii Gerlach,

1.6 Systematics 

Genus Thaumastoderma Remane, 1926 (Fig. 1.71 A–C) The species in this genus resemble those of Tetranchyroderma in the general body shape. Their body is covered exclusively with tetrancres. Additionally, specimens have at least two pairs of cephalic appendages: one pair of

A

rt

50 µm

spatulate tentacles and one pair of lateral tentacles (Fig. 1.18 F and 1.71 A–C). Apart from T. renaudae, all remaining species have another pair of anterior, rod-shaped tentacles (Kieneke 2010). Very conspicuous are paired dorsal appendages, the dorsal cirrata, which are present in 4, 5, or 6 pairs. Kieneke (2010) provides an overview on the characters and a phylogenetic analysis of the genus. Seventeen species are currently known, most from the eastern North Atlantic and the waters around Korea and Japan; fewer species from the Baltic Sea, the Mediterranean, the western Atlantic (South Carolina), and the Antarctic Deep Sea: T. antarctica Kieneke, 2010, T. appendiculatum Chang, Lee & Clausen, 1998, T. arcassonense d’Hondt, 1965, T. bifurcatum Clausen, 1991, T. cantacuzeni Lévi, 1958, T. clandestinum Chang, Kubota & Shirayama, 2002, T. copiophorum Chang, Lee & Clausen, 1998, T. coronarium Chang, Lee & Clausen, 1998, T. heideri Remane, 1926, T. mediterraneum Remane, 1927, T. minancrum Hummon, 2008, T. moebjergi Clausen, 2005, T. natlanticum Hummon, 2008, T. ramuliferum Clausen, 1965, T. renaudae Kisielewski, 1987, T. swedmarki Lévi, 1950, and T. truncatum Clausen, 1991. 1.6.1.7 Family Turbanellidae Remane, 1927 Species are strap-shaped, and the head may be delimited or not. Head appendages are present in some species. At

B

st

C

50 µm

Fig. 1.71: Thaumastoderma mediterraneum (Macrodasyida, Thaumastodermatidae) from the shallow sublittoral around Elba, Italy. (A) Ventral view. (B) Horizontal view. (C) Dorsal view. Abbreviations: rt, rod-shaped tentacles; st, spatulate tentacles. (A–C) BF images.

50 µm

1953, T. paradoxum Thane-Fenchel, 1970, T. paralittorale Chandrasekhara Rao, 1991, T. parapapii Hummon, 2009, T. pentasperum Nicholas & Todaro, 2006, T. pinnatum Lee, 2012, T. polyacanthum (Remane, 1927), T. polypodium Luporini, Magagnini & Tongiorgi, 1971, T. polyprobolostomum Hummon, Todaro, Balsamo & Tongiorgi, 1996, T. psilotopum Hummon, Todaro, Tongiorgi & Balsamo, 1998, T. pugetense Wieser, 1957, T. quadritentaculatum Todaro, Balsamo & Tongiorgi, 1992, T. rhopalotum Hummon, 2011, T. sanctaecaterinae Todaro, Balsamo & Tongiorgi, 1992, T. sardum Todaro, Balsamo & Tongiorgi, 1988, T. schizocirratum Chang, Kubota & Shirayama, 2002, T. sinaiensis Hummon, 2011, T. suecicum Boaden, 1960, T. swedmarki Chandrasekhara Rao & Ganapati, 1968, T. symphorochetum Hummon, Todaro, Tongiorgi & Balsamo, 1998, T. tanymesatherum Hummon, Todaro, Balsamo & Tongiorgi, 1996, T. tentaculatum Chandrasekhara Rao, 1993, T. thysanogaster Boaden, 1965, T. thysanophorum Hummon, Todaro & Tongiorgi, 1993, T. tribolosum Clausen, 1965, T. verum Wilke, 1954, T. weissi Todaro, 2002, and T. xenodactylum Hummon, 2011.

 105

106 

 1 Gastrotricha

A

B

TbA

TbP

50 µm

40 µm

Fig. 1.72: Desmodasys abyssalis (Macrodasyida, Turbanellidae), fixed animals. (A) Specimen from the Antarctic deep sea floor at depth of 5191 m, ventral view. (B) Specimen from artificial tubeworm aggregations deposited close to a hydrothermal vent of the East Pacific Rise at depth of 2500 m. Abbreviations: TbA, anterior adhesive tubes; TbP, posterior adhesive tubes. (A) DIC image. (B) SEM micrograph.

the posterior end there is a pair of caudal feet with adhesive tubes (TbP) on the posterior margin of the entire foot. The arrangement of adhesive tubes is variable. TbA are often concentrated on fleshy hands; TbL can be numerous, strongly reduced in number, or completely absent. In some species/genera, characteristically large adhesive tubes are present. The ventral ciliation is usually on the entire anterior region and in paired rows along the trunk. The pharyngeal pores are in the posterior region of the pharynx, with the exception of the genus Prostobuccantia, where they are significantly more anterior. The family includes six genera: Desmodasys, Dinodasys, Paraturbanella, Prostobuccantia, Pseudoturbanella, and Turbanella.

TbA

Genus Desmodasys Clausen, 1965 (Fig. 1.72 A, B) The genus includes two species from the coast of Norway (eastern North Atlantic) and one species from the deep sea at the East Pacific Rise. A head is more or less delimited; ventral ciliation is in two bands (D. borealis and D. phocoides) or on the entire ventral side (D. abyssalis). Eyes can be present. A small buccal cavity is present, and the pharyngeal pores are close to the pharynx-intestine junction. The cuticle is smooth. TbL are absent, only sensory cilia are arranged along the trunk. The TbA are arranged in paired anteroventral tufts. TbP are on caudal feet. The three species described to date are: D. abyssalis Kieneke & Zekely, 2007, D. borealis Clausen, 2000, and D. phocoides Clausen, 1965. Genus Dinodasys Remane, 1927 (Fig. 1.73) The two species of Dinodasys have a more or less welldefined head, which is bordered by a pair of large tentacles. Additional smaller “cephalic protrusions” are present in different number on the dorsal and the ventral side of the head. The TbA are on ventral fleshy

100 µm Fig. 1.73: Dinodasys mirabilis (Macrodasyida, Turbanellidae) from sublittoral sand of the outer Jade Bay, Germany. Note the cylindric buccal cavity. Abbreviation: TbA, anterior adhesive tubes. BF image.

1.6 Systematics 

A

B

C

an

TbA

50 µm

D

 107

50 µm lao

200 µm

hands, the TbP on caudal feet. TbL are numerous. The two species described were found on both sides of the North Atlantic, in the North Sea, and on the coast of Delaware: D. delawarensis Hummon, 2008, and D. mirabilis Remane, 1927. Genus Paraturbanella Remane, 1927 (Fig. 1.74 A–D) Very conspicuous is a paired accessory adhesive organ made up of a long and a shorter adhesive tube; this is present at the level of the anterior region of the pharynx (Fig. 1.74 D). TbA are on fleshy hands (Fig. 1.74 B); TbL are few and quite small or may be absent at all; TbP are on the caudal feet. A key to species was provided by Wieser (1957). Twenty-two species are currently known from the North Atlantic, the Mediterranean, the Caribbean, the Pacific, India, and Australia: P. aggregotubulata Evans, 1992, P. armoricana (Swedmark, 1954), P. boadeni Chandrasekahara Rao & Ganapati, 1968, P. brevicaudata Chandrasekahara Rao, 1991, P. cuanensis Maguire, 1976, P. dohrni Remane, 1927, P. dolichodema Hummon, 2010, P. eireanna Maguire, 1976, P. intermedia Wieser, 1957, P. levantia Hummon, 2011, P. manxensis Hummon, 2008, P. microptera Wilke, 1954, P. mesoptera Chandrasekahara Rao, 1970, P. pacifica Schmidt, 1974, P. pallida Luporini, ­Magagnini & Tongiorgi, 1971, P. palpibara Chandrasekahara Rao & Ganapati, 1968, P. pediballetor Hummon, 2008,

100 µm

Fig. 1.74: Paraturbanella pacifica (Macrodasyida, Turbanellidae) from sublittoral sand of Little Cayman Island. (A) Habitus in horizontal view. (B) Anterior end in ventral view showing the handlike arranged anterior adhesive tubes. (C) Ventral view of the posterior end. (D) Anterior end focused on the peculiar lateral accessory adhesive organs. Abbreviations: an, anus; TbA, anterior adhesive tubes; lao, lateral adhesive organ. (A–D) DIC images.

P. sanjuanensis Hummon, 2010, P. scanica Clausen, 1996, P. solitaria Todaro, 1995, P. stradbroki Hochberg, 2002, and P. teissieri Swedmark, 1954. Genus Prostobuccantia Evans & Hummon, 1991 A single species, P. brocha Evans & Hummon, 1991, has been described from Florida (Evans & Hummon 1991). The most characteristic feature is a “corona”, an anterior projection of the buccal cavity. The pharyngeal pores are, unusual for a member of Turbanellidae, not close to the pharyngointestinal border, but further anterior. The TbA are on hands, TbL are present posterior of about the level of the pharyngeal pores. Quite conspicuous is a pair of a very long and a shorter adhesive tube shortly behind the pharyngointestinal junction. The TbP are on caudal feet. Genus Pseudoturbanella d’Hondt, 1968 One species, P. stylifera d’Hondt, 1968, is known in this genus, and it was found on the French Atlantic coast (d’Hondt 1968). The TbA are on paired hands, one pair of very large tubes attaches on the posterior margin of the well delimited head. TbL are reduced to one single pair at the level of the pharyngointestinal junction. The caudal feet carry few adhesive tubes on their inner side and their terminal end.

108 

 1 Gastrotricha

A

B TbA

100 µm

pp mgp

C

200 µm

Genus Turbanella Schultze, 1853 (Fig. 1.75 A–C) A head region is more or less clearly delimited; in some species, there are lateral lobes (Fig. 1.75 A). TbA are on fleshy hands (Fig. 1.75 B); TbL are abundant along the entire trunk in almost all species, frequently arranged in ventral, ventrolateral, lateral, dorsolateral and/or dorsal series. TbP are on caudal feet (Fig. 1.75 C). An unpaired, median cone is present between the two caudal feet in many species. A key was provided by Wieser (1957). Twenty-nine species are currently known with a distribution in the Atlantic (both coasts of the North Atlantic plus western South Atlantic and Caribbean), the Baltic Sea (brackish), the Mediterranean and Black Sea, the Red Sea, the Pacific (western coast of North and Central America, Galapagos), India, and Australia: T. ambronensis Remane, 1943, T. aminensis Chandrasekahara Rao, 1991, T. amphiatlantica Hummon & Kelly, 2011, T. bengalensis Chandrasekahara Rao & Ganapati, 1968, T. bocqueti Kaplan, 1958, T. brusci Hochberg, 2002, T. caledoniensis Hummon, 2008, T. corderoi Dioni, 1960, T. cornuta Remane, 1925, T. erythrothalassia Hummon, 2011, T. hyalina Schultze, 1853, T. indica Chandrasekahara Rao, 1981, T. lutheri Remane, 1952, T. mikrogada Hummon, 2008, T. multidigitata Kisielewski, 1987, T. mustela Wieser, 1957, T. ocellata Hummon, 1974, T. otti Schrom, 1972, T. pacifica Schmidt, 1974, T. palaciosi Remane, 1953, T. petiti Remane, 1952, T. pontica Valkanov, 1957, T. reducta Boaden, 1974, T. remanei Forneris, 1961, T. scilloniensis Hummon, 2008, T. subterranea

50 µm

Fig. 1.75: (A) Turbanella ambronensis (Macrodasyida, Turbanellidae) from an intertidal sand bar close to Saint-Jacut-de-la-Mer, France. Habitus in ventral view. (B and C) Turbanella hyalina from the intertidal at Schillig, Germany. (B) Anterior end in ventral view. (C) Posterior end in ventral view. Abbreviations: mgp, male genital pore; pp, pharyngeal pores; TbA, anterior adhesive tubes. (A) BF image. (B and C) DIC images.

Remane, 1934, T. varians Maguire, 1976, T. veneziana Schrom, 1972, and T. wieseri Hummon, 2010.

1.6.1.8 Family Xenodasyidae Todaro, Guidi, Leasi & Tongiorgi, 2006 This family, including four species in two genera, includes conspicuous specimens with a clearly delimited head, from which a strong pair of segmented appendages, usually called tentacles, originates (Fig. 1.18 D). Additional pairs of dorsal unsegmented appendages may be present on the head. The dorsal surface is either covered by cuticular structures (Xenodasys) or smooth (Chordodasiopsis). TbL are few and difficult to observe, but at least one pair of large ventral adhesive structures, often called pedicles, is present. Each pedicle is composed of a pair of closely attached adhesive tubes. The caudal feet are elongate and directed posteriorly. Posterior of the intestine is a chordoid organ, a median longitudinal structure made up of modified muscle cells. Muscles are cross-striated, and the pharynx and intestine are, at least partially, ciliated (see Todaro et al. 2006 for a more detailed description). The first species was described by Swedmark (1967) as Xenodasys sanctigoulveni, the second by Schöpfer-Sterrer (1969) as Chordodasys riedli. A third species was described as Chordodasys antennatus (Rieger et al. 1974). Kisielewski (1987a) regarded Chordodasys as synonymous to Xenodasys (see also Hummon 1982), but Todaro et  al. (2006b),

1.6 Systematics 

 109

Shetland Islands in the eastern Atlantic and probably in South Africa (see Todaro et al. 2006b). Genus Xenodasys Swedmark, 1967 (Fig. 1.76) One pair of segmented tentacles is present; dorsal appendages may be present on the head. The dorsal cuticle is roughly structured into plates and spines. The TbA originate from fleshy hands. TbL are few, one, or two pairs of pedicles are present. TbP are present on the caudal feet. See Todaro et al. (2006b) for more information. Three species have been described from the eastern North Atlantic (X. sanctigoulveni Swedmark, 1967), the western North Atlantic and the Caribbean (X. riedli Schöpfer-Sterrer, 1969), and the Mediterranean Sea (X. eknomios Todaro, Guidi, Leasi & Tongiorgi, 2006).

1.6.1.9 Family Redudasyidae Todaro, Dal Zotto, Jondelius, Hochberg, Hummon, Kånneby & Rocha, 2012

100 µm Fig. 1.76: Xenodasys riedli (Macrodasyida, Xenodasyidae) from calcareous sand of San Salvador Island, Bahamas. Dorsal view. Note the seemingly segmented tentacles on the head region. DIC image.

when describing a fourth species, Xenodasys eknomios, regarded Xenodasys antennatus as significantly different from the three other species and erected a new genus, Chordodasiopsis, for it. Genus Chordodasiopsis Todaro, Guidi, Leasi & Tongiorgi, 2006 One pair of segmented tentacles and two pairs of unsegmented dorsal appendages are present on the head. The dorsal cuticle is smooth, only structured by sensory processes. Numerous of those so-called regular sensory processes ar arranged along the trunk in a lateral and dorsolateral position. The TbA originate directly from the surface of the body, not on hands. Adhesive tubes are not visible on the caudal feet, instead there is an adhesive pad. One pair of ventral pedicles is present. The single species, C. antennatus (Rieger, Ruppert, Rieger & SchoepferSterrer, 1974), was found on the western North Atlantic (North Carolina, Florida, Virgin Islands, Bermuda), the

This family includes two species, Redudasys fornerise and Anandrodasys agadasys. Both species cluster close together in an analysis of their 18S rDNA sequences (Todaro et al. 2012b). Most of the characters, in which the two species resemble each other (see Todaro et al. 2012b) are plesiomorphies, which are also present in several other gastrotrich species (Kieneke et al. 2013a), but a potential synapomorphy is that both species appear to be parthenogenetic (Todaro et al. 2012b, Kieneke et al. 2013a). Genus Redudasys Kisielewski, 1987 (see Fig. 1.26 B) Redudasys fornerise Kisielewski, 1987, is one of two species of Macrodasyida found in freshwater. It was originally described by Kisielewski (1987a) from Brazil and then rediscovered by Todaro et al. (2012b). The body is slender (Kisielewski 1987a) to tenpin-shaped (Todaro et al. 2012b), probably due to the presence of mature eggs. Two TbA are present on each side; they originate from a common base. At the posterior end of the animals, two TbP are present on each side. The ventral ciliation is in separate fields of unequal size. The pharyngeal pores are in the posterior region of the pharynx. Recently, Garraffoni et  al. (2010) reported the discovery of several specimens of Redudasys that possibly belong to another species. Genus Anandrodasys Todaro, Dal Zotto, Jondelius, Hochberg, Hummon, Kånneby & Rocha, 2012 (Fig. 1.77) Originally described as Dactylopodola agadasys from Australia (Hochberg 2003), Todaro et  al. (2012b) found this species to cluster close to Redudasys fornerise and created a new genus for it, Anandrodasys. Specimens have been

110 

 1 Gastrotricha

A

et al. 2013a). Five TbP fan from what might be considered as a very short caudal lobe on both sides of the body. The ventral ciliation is in paired columns.

B

1.6.2 Order Chaetonotida Brunson, 1950

*

100 µm

100 µm

*

Remane (1924) used the name “Chaetonotoidea” for the longknown freshwater gastrotrichs in comparison to the newly discovered, marine macrodasyids. Brunson (1950) used the writing “Chaetonotida”, which was suggested to be the preferable name by Chandrasekahara Rao & Clausen (1970). Chaetonotids live in freshwater as well as in the marine environment. They include one genus, Neodasys, specimens of which closely resemble the general macrodasyid shape of the body. All remaining chaetonotids, summarized as Paucitubulatina, have a more or less tenpin-shaped body and have adhesive tubes exclusively on the posterior end. In many, but not all species, the body is covered with cuticular scales or spines, which are important taxonomic markers. Characteristic for all chaetonotids is the absence of pharyngeal pores (this is, among macrodasyids, only known from Lepidodasys species) and a Y-shaped pharyngeal lumen, meaning that the triradiate lumen has an unpaired ventral branch and paired dorsolateral branches.

Fig. 1.77: Anandrodasys agadasys (Macrodasyida, Redudasyidae) from calcareous sand of Lee Stocking Island, Bahamas. (A) Dorsal view. (B) Ventral view. Note the ventrolateral adhesive tubes, which are restricted to the midtrunk region (asterisks). (A and B) DIC images. (from Kieneke et al. 2013a, with kind permission of Verlag Dr. Friedrich Pfeil.)

found, apart from the first record in Australia, in the Red Sea (Hummon 2011) and in several localities in the Caribbean (see Kieneke et al. 2013a). The animals are slender; the head is not clearly delimited. The buccal cavity is large, and the pharyngeal pores are in the posterior region of the pharynx. Three TbA are present per side; they form a short row and decrease in size from lateral to medial. Few ventrolateral adhesive tubes are present in about the middle region of the trunk; they have been found to be asymmetric in number and arrangement among several specimens from the Red Sea and the Caribbean (Kieneke

1.6.2.1 [Suborder Multitubulatina d’Hondt, 1971] 1.6.2.1.1 [Family Neodasyidae Remane, 1929] Genus Neodasys Remane, 1927 (Fig. 1.78) The three species of this genus resemble in their external morphology macrodasyidan gastrotrichs because they have an elongate body with a number of TbL. The orientation of the pharyngeal lumen (Y) and the absence of pharyngeal pores correspond to (other) chaetonotids and Neodasys is therefore traditionally regarded as member of the Chaetonotida. However, this systematic position of Neodasys within Chaetonotida is recently challenged by several phylogenetic analyses of either molecular data or coded morphological characters (see chapter Phylogeny). According to these, Neodasys could also be part of the Macrodasyida or may represent the earliest lineage within Gastrotricha. The genus is the only component of the suborder Multitubulatina and of the family Neodasyidae. A head region is evident in N. chaetonotoides and N. uchidai, but not in N. cirritus. TbA are not present; the TbL are inconspicuous and short. However, small papillae on the ventral side of the head of N. uchidai were regarded as remnants of TbA (Remane 1961). In the posterior

1.6 Systematics 

 111

end, a pair of caudal feet is present; in N. uchidai, the feet originate from a common unpaired stem. TbP are present on the caudal feet. The mouth opening is surrounded by a cup-shaped structure. The cuticle is smooth. Three species were described from the North Sea, the Western Mediterranean Sea, and from the Atlantic coast of Florida: N. chaetonotoideus Remane, 1927, N. cirritus Evans, 1992 (see Hummon & Todaro 2010 for a nomenclatural change), and N. uchidai Remane, 1961.

1.6.2.2 Suborder Paucitubulatina d’Hondt, 1971 All species of Paucitubulatina are characterized by the absence of adhesive tubes in places other than on the paired caudal extensions, which are usually named furca. With two exceptions (see Diuronotus and Dichaetura), only one single adhesive tube is present per side. TbA and TbL are always lacking. The caudal feet are lacking in some swimming species; in this case, no adhesive tubes are present at all. Most species have a more or less tenpin-shaped body form, with a head region, a thinner neck, and a wider trunk. Very important for the determination of species are the cuticular structures. These may be lacking, but many species are covered with scales or spines. There is a wide diversity of such cuticular structures (see, e.g., Schwank 1990). Scales are flat structures, these may carry a keel or a spine. Spines appear to be derived from spined scales; many spines have a small scale as basis and only few spines appear to originate directly from the cuticle. Scales may also be stalked. The pharynx has in some species one or even more bulbs. A buccal cavity is usually present, the mouth can be surrounded by a cuticular ring and/or spine-like rods. Further cuticular structures are present in the buccal cavity or pharynx of few species. Paucitubulatina are abundant in freshwater, but several species do occur in the marine environment or in brackish waters.

mo

co

200 µm

fo

Fig. 1.78: Neodasys chaetonotoideus (Multitubulatina) from an intertidal beach of river Ems estuary, Germany. Dorsal view. Abbreviations: co, caudal organ; fo, frontal organ; mo, mature egg. BF image.

1.6.2.2.1 Family Muselliferidae Leasi & Todaro, 2008 On the basis of a cladistic analysis of muscle characters, Leasi & Todaro (2008) erected the family Muselliferidae for the genera Musellifer and Diuronotus. The head region is weakly separated from the remaining body and it lacks appendages or extensions. In some species, the head characteristically tapers toward the anterior end and is densely covered with cilia thus forming a conspicuous “muzzle”. The mouth opening is surrounded by a ring of teeth-like cuticular ridges. One (Musellifer) or two (Diuronotus) pairs of adhesive tubes is present on the furca. The body is completely covered with uniform scales, which are either spined (Musellifer) or keeled (Diuronotus). The ventral ciliation is on the entire anterior ventral region and in two longitudinal bands on the posterior region (see

112 

 1 Gastrotricha

Leasi & Todaro 2008 for more characters). The species are hermaphrodites and occur only marine. Genus Diuronotus Todaro, Balsamo & Kristensen, 2005 (Fig. 1.79) The two species of Diuronotus resemble species of Musellifer in many respects, but differ from these in the following characters (Todaro et  al. 2005, Balsamo et  al. 2010a): a second pair of smaller adhesive tubes is present additional to the longer one in Diuronotus. The scales have a keel in Diuronotus, in Musellifer a keel is, when present, very weak and scales are spined. Finally, Diuronotus has a smaller furca to total body length ratio compared with Musellifer. Two species are known from sediment samples from the shore or near the shore in Denmark and Greenland: D. aspetos Todaro, Balsamo & Kristensen, 2005, and D. rupperti Todaro, Balsamo & Kristensen, 2005.

Genus Musellifer Hummon, 1969 (Fig. 1.80) The characters to distinguish Musellifer from Diuronotus are described above. Five species were described from the eastern North Atlantic, the Mediterranean Sea, the eastern Pacific, and recently from the western Atlantic and Caribbean Sea: M. delamarei (Renaud-Mornant, 1968), M. profundus Vivier, 1974, M. reichardti Kånneby, Atherton & Hochberg, 2014, M. sublitoralis Hummon, 1969, and M. tridentatus Kånneby, Atherton & Hochberg, 2014. 1.6.2.2.2 Family Xenotrichulidae Remane, 1936 This family includes small and very fast species, which typically occur in high-energy beaches (Ruppert 1979). See Ruppert (1979) for a review of the family. The most important character is that the ventral cilia are clustered into bundles called cirri. The caudal furca is very long, its base is covered by overlapping scales. In the head region, three pairs of sensory structures (often called sensoria) are present, there are at least five pairs of elongated sensory cilia along the body.

mz

100 µm

TbP

st Fig. 1.79: Diuronotus aspetos (Paucitubulatina, Muselliferidae) from the sublittoral off island Wangerooge, Germany. The secondary adhesive tube is out of focus. Abbreviations: st, secondary adhesive tube; TbP, posterior adhesive tube. DIC image.

50 µm Fig. 1.80: Musellifer cf. profundus (Paucitubulatina, Muselliferidae) from the Antarctic deep sea floor at depth of 5191 m. Fixed and stained specimen in horizontal view. Abbreviation: mz, ciliated muzzle. DIC image.

1.6 Systematics 

The three genera are usually split into two subfamilies (Ruppert 1979): Draculiciterinae (genus Draculiciteria) and Xenotrichulinae (genera Xenotrichula and Heteroxenotrichula). Genus Draculiciteria Hummon, 1974 (Fig. 1.81 A, B) The head is well separated from the trunk; the two posterior sensoria of each side are fused into one bundle of sensory cilia (all characters from Ruppert 1979). The furca is not well delimited from the trunk, the furcal branches are slightly curved. Most of the length of the furca branches (about 5/6) is covered with scales. The body is covered by scales and, on the lateral sides, with stalked scales. The dorsal side of the head is covered by a characteristic pattern of polygonal cuticular plates (Fig. 1.4 G). The pharynx has an anterior bulb, eyes are absent. One species is known from the eastern North Atlantic (Gulf of Mexico and Caribbean) and the Mediterranean Sea: D. tesselata (Renaud Mornant, 1968).

 113

Genus Xenotrichula Remane, 1927 (Fig. 1.82 A–C) The two genera, Xenotrichula and Heteroxenotrichula, have a poorly defined head, but furcal branches are well delimited from the trunk (Ruppert 1979). Scales are present up to two thirds of the furcal length. In Xenotrichula, all locomotory ventral cirri are of equal length (Fig. 1.82 A), the pharynx has no bulb, and the mouth opening is surrounded by folds and spines. Fifteen species are known from both sides of the North Atlantic Ocean, the Caribbean, the Baltic Sea, the Mediterranean Sea, the Pacific (Galapagos), and India: X. bispina Roszczak, 1939, X. carolinensis Ruppert, 1979, X. cornuta Wilke, 1954, X. floridana Thane-Fenchel, 1970, X. guadelupensis Kisielewski, 1984, X. intermedia Remane, 1934, X. laccadivensis Rao, 1991, X. lineata Schrom, 1972, X. micracantha (Remane, 1926a), X. paralineata Hummon & Todaro, 2007, X. punctata Wilke, 1954, X. quadritubulata Kisielewski, 1988, X. soikai Schrom, 1966, X. tentaculata Rao & Ganapati, 1968, and X. velox Remane, 1927.

A

100 µm

B cir

50 µm Fig. 1.81: Draculiciteria tesselata (Paucitubulatina, Xenotrichulidae) from calcareous sand of Lee Stocking Island, Bahamas. (A) Habitus in horizontal view. (B) Slightly smaller specimen in ventral view. Abbreviation: cir, locomotory cirri. (A and B) DIC images.

114 

 1 Gastrotricha

A

B

C

cir

hf

cir

Genus Heteroxenotrichula Wilke, 1954 (Fig. 1.83) Apart from the characters named under Xenotrichula, the most obvious character of Heteroxenotrichula species is the presence of ventral locomotory cirri in two size classes. The mouth is slightly subventral and surrounded by a collar, but not by folds or spines. The pharynx has an anterior bulb. Nine species have been described from both sides of the North Atlantic Ocean, the Caribbean, the Baltic Sea, the Mediterranean Sea, the Pacific (Galapagos), and India: H. affinis (Remane, 1934), H. arcassonensis Ruppert, 1979, H. pygmaea (Remane, 1934), H. simplex (Mock, 1979), H. squamosa Wilke, 1954, H. subterranea (Remane, 1934), H. texana Todaro, 1994, H. transatlantica Ruppert, 1979, and H. wilkeae Ruppert, 1979. 1.6.2.2.3 Family Chaetonotidae Gosse, 1864 A head region is more or less clearly separated, often the head is composed of three or five lobes. Bundles of cilia are present in the head region, often cephalic shields are present (Fig. 1.4 J, K). A furca with one pair of adhesive tubes is present. The cuticle may be naked, but in most species, it is covered with cuticular scales of diverse shapes. Ventral cilia are usually in two longitudinal bands, rarely in other patterns. The pharynx is without bulbs, only slight thickenings of the pharynx can be present. A weir is usually present at the transition to the intestine. Several species have rudimentary testes (e.g., Weiss & Levy 1979, Weiss 2001, see chapters Reproductive Organs and Reproductive Biology). This is a very large

50 µm

50 µm

50 µm

hf

Fig. 1.82: (A) Xenotrichula intermedia (Paucitubulatina, Xenotrichulidae) from Geniusstrand (no more existant) in the north of Wilhelmshaven, Germany. Ventral view. (B and C) X. velox. (B) Specimen from the beach of Tylosand, Sweden. Horizontal view with focus on internal organs. (C) Specimen from the intertidal sand flat at Les Hemmes Plage, France. Focus on the ventral side showing the hydrofoil scales. Abbreviations: cir, locomotory cirri; hf, hydrofoil scales. (A and B) DIC images. (C) BF image.

family with 12 genera (Arenotus, Aspidiophorus, Caudichthydium, Chaetonotus, Fluxiderma, Halichaetonotus, Heterolepidoderma, Ichthydium, Lepidodermella, Polymerurus, Rhomballichthys, and Undula); almost all genera belong to the subfamily Chaetonotinae, and only Undula belongs to the Undulinae (Kisielewski 1991). Schwank (1990, page 61) includes in a key one further genus, Hemichaetonotus, with H. clipeatus as type species and eight further marine species that are not listed. He obviously intended to move, Heterolepidoderma clipeatum Schrom, 1972, into a new genus, but as a diagnosis was not given, the name Hemichaetonotus has to be regarded a nomen nudum. Genus Arenotus Kisielewski, 1987 The body is covered with a thick cuticular layer, but there are no further cuticular structures like scales or spines (all data from Kisielewski 1987a). The mouth is surrounded by a complex collar, a pair of strong teeth originates dorsolaterally in the pharynx. In the head region, cuticular plates are present in the form of the unpaired cephalion (dorsal) and hypostomium (ventral) and two pairs of pleuria. One species, A. strixinoi Kisielewski, 1987, is described from freshwater in Brazil. Genus Aspidiophorus Voigt, 1903 (Fig. 1.84) Species of this genus have the more or less “usual” appearance of Chaetonotidae. Characteristic is that the body is densely covered by stalked scales. The scales are often rhombic, sometimes more oval, and usually have two to three keels (Fig. 1.4 C). The scale covering usually ends

1.6 Systematics 

 115

hy

50 µm Fig. 1.83: Heteroxenotrichula cf. affinis (Paucitubulatina, Xenotrichulidae) from sediment at the base of the dike in Wilhelmshaven, Germany. Ventral view. DIC image.

anterior of the furcal base. See Schwank (1990) for characters and a key and Todaro et al. (2009) for additional comments and a key to the marine species. Thirty-two species (22 from freshwater and 10 marine) have been described from freshwater environments in Europe, North America, Argentina, and Japan and from marine sediments from the North Atlantic, Baltic Sea, Mediterranean Sea, the Caribbean, and the Sea of Japan: A. aster Martin, 1981, A. bibulbosus Kisielewski, 1979, A. bisquamosus Mock, 1979,

50 µm Fig. 1.84: Aspidiophorus paramediterraneus (Paucitubulatina, Chaetonotidae) from the shallow sublittoral of the Bay of Fetovaia, Elba, Italy. Ventral view. Abbreviation: hy, hypostomion. BF image.

A. bramhsi Grosso, 1973, A. heterodermus Saito, 1937, A. heterodermus Saito, 1937, A. lilloensis Grosso & Drahg, 1983, A. lamellophorus Balsamo, Hummon, Todaro & Tongiorgi, 1997, A. longichaetus Kisielewski, 1986, A. marinus Remane, 1926, A. mediterraneus Remane, 1927, A. multitubulatus Hummon, 1974, A. microlepidotus d’Hondt, 1978, A. microsquamatus Saito, 1937, A. nipponensis Schwank, 1990, A. oculifer Kisielewski, 1981, A. oculatus Todaro, Dal Zotto, Maiorova &

116 

 1 Gastrotricha

Adrianov, 2009, A. ophiodermus Balsamo, 1983, A. ornatus Mock, 1979, A. paradoxus (Voigt, 1902), A. paramediterraneus Hummon, 1974, A. pleustonicus Kisielewski, 1991, A. polonicus Kisielewski, 1981, A. polystictos Balsamo & Todaro, 1987, A. pori Kisielewski, 1999, A. schlitzensis Schwank, 1990, A. semirotundus Saito, 1937, A. slovinensis Kisielewski, 1986, A. squamulosus Roszczak, 1936, A. tatraënsis Kisielewski, 1986, A. tentaculatus Wilke, 1954, and A. tetrachaetus Kisielewski, 1986.

Genus Caudichthydium Schwank, 1990 Schwank (1990) introduces this genus in a key with a very brief characterization of characters. Mock (1979) has already speculated that the three species united in this genus may have to be separated from the genus Ichthydium, in which they were originally described. The main character is that the furcal branches are fused more or less significantly at their base. The cuticle is naked and has no scales or spines. The ventral cilia occur in bundles. Three species have been described from the North Atlantic (North Carolina, Arcachon, Sylt): C. hummoni (Ruppert, 1977), C. rupperti (Mock, 1979), and C. supralitorale (Mock, 1979). Genus Chaetonotus Ehrenberg, 1830 (Fig. 1.85 A, B) Chaetonotus is a large genus with species carrying spined scales (Fig. 1.4 B). In some species, the spines on the scales are quite long and strong. Balsamo et  al. (2009) name 163 freshwater and 40 marine species in this genus, here we list 202 species compiled from Balsamo et al. (2009) and Hummon & Todaro (2010) and

100 µm

B

100 µm

A

including the new descriptions since 2010. Usually, the genus is subdivided into several subgenera, some of these subgenera are more or less provisional constructs and the list of subgenera has changed through time (see Schwank 1990, Kisielewski 1997, Balsamo et  al. 2009, with addition of Tristatachaetus by Kolicka et al. 2013). Taking records of all species into account, the genus has a worldwide distribution. C. acanthodes Stokes, 1887, C. acanthophorus Stokes, 1887, C. aculeatus Robbins, 1965, C. aegilonensis Balsamo, Todaro & Tongiorgi, 1992, C. aemilianus Balsamo, 1978, C. aequispinosus (Schrom, 1972), C. africanus Schwank, 1990, C. alatus Schwank, 1990, C. alni Nesteruk, 1991, C. angustus Schrom, 1972, C. annectens Grosso & Drahg, 1991, C. anomalus Brunson, 1950, C. antipai (Rodewald, 1938), C. apechochaetus Hummon, Balsamo & Todaro, 1992, C. apolemmus (Hummon, Balsamo & Todaro, 1992), C. arethusae Balsamo & Todaro, 1995, C. armatus Kisielewski, 1981, C. arquatus Voigt, 1903, C. atrox (Wilke, 1954), C. australiensis Schwank, 1990, C. chuni Voigt, 1901, C. cordiformis Greuter, 1917, C. balsamoae Kisielewski, 1998, C. beauchampi d’Hondt, 1967, C. benacensis Balsamo & Fregni, 1995, C. bifidispinosus Tretjakova, 1991, C. bisacer Greuter, 1917, C. bogdanovii (Schimkewitsch, 1886), C. brachyurus Balsamo, 1981, C. brasilianus (Kisielewski, 1991), C. brasiliensis Schwank, 1990, C. breviacanthus Kisielewski, 1991, C. brevispinosus Zelinka, 1889, C. caricicola Schwank, 1990, C. carpaticus Rudescu, 1967, C. caudalspinosus Visvesvara, 1964, C. cestacanthus Balsamo, 1990, C. chicous (Hummon, 1974), C. christianus Schwank, 1990, C. condensus Mock, 1979, C. corderoi Schwank, 1990, C. crassus Preobrajenskaja, 1926, C. dadayi Schwank, 1990, C. daphnes Balsamo & Todaro, 1995, C. decemsetosus Marcolongo,

fu

Fig. 1.85: Chaetonotus luporinii (Paucitubulatina, Chaetonotidae) from calcareous sand around the island of Pianosa, Italy. (A) Dorsal view showing the spined scales. Note the crest-shaped basal part of the scales. (B) Ventral view. Abbreviation: fu, furca with both posterior adhesive tubes. (A and B) BF images.

1.6 Systematics 

1910, C. disjunctus Greuter, 1917, C. dispar (Wilke, 1954), C. dracunculus Balsamo, 1990, C. dubius Daday, 1905, C. dybowskii Jakubski, 1919, C. elachysomus Hummon, 2010, C. elegans Konsuloff, 1921, C. enormis Stokes, 1887, C. eratus Hummon, 2010, C. erinaceus Daday, 1905, C. euhystrix Schwank, 1990, C. eximius Kolicka, Kisielewski, Nesteruk & Zawierucha, 2013, C. fenchelae d’Hondt, 1974, C. ferrarius Schwank, 1990, C. fluviatilis Balsamo & Kisielewski, 1986, C. formosus Stokes, 1887, C. fujisanensis (Sudzuki, 1971), C. furcatus Kisielewski, 1991, C. gastrocyaneus Brunson, 1950, C. greuteri Remane, 1927, C. heideri Brehm, 1917, C. heteracanthus Remane, 1927, C. heterochaetus Daday, 1905, C. heterospinosus Balsamo, 1978, C. hilarus (Schrom, 1972), C. hirsutus Marcolongo, 1910, C. hoanicus Schwank, 1990, C. hystrix (Metschnikoff, 1865), C. ichthydioides Tongiorgi, Fregni & Balsamo, 1999, C. illiesi Schwank, 1990, C. inaequidentatus Kisielewski, 1988, C. insigniformis Greuter, 1917, C. intermedius Kisielewski, 1991, C. italicus Balsamo & Todaro, 1995, C. jakubskii Roszczak, 1936, C. lacunosus (Mock, 1979), C. laroides Marcolongo, 1910, C. larus (Müller, 1773), C. laterospinosus Visvesvara, 1965, C. linguaeformis Voigt, 1902, C. lobo Kisielewski, 1991, C. longisetosus Preobrajenskaja, 1926, C. longispinosus Stokes, 1887, C. lucksi Voigt, 1958, C. lunatospinosus Balsamo, 1981, C. luporinii Balsamo, Fregni & Tongiorgi, 1996, C. machikanensis Suzuki & Furuya, 2011, C. macrochaetus Zelinka, 1889, C. macrolepidotus Greuter, 1917, C. magnificus Balsamo, Hummon, Todaro & Tongiorgi, 1997, C. majestuosus Grosso & Drahg, 1984, C. mariae (Todaro, 1992), C. maximus Ehrenberg, 1831, C. mediterraneus Balsamo, Hummon, Todaro & Tongiorgi, 1997, C. microchaetus Preobrajenskaja, 1926, C. minimus Marcolongo, 1910, C. mitraformis Greuter, 1917, C. modestus (Schrom, 1972), C. monobarbatus Visvesvara, 1965, C. montevideensis Cordero, 1918, C. multisetosus Preobrajenskaja, 1926, C. multispinosus Grünspan, 1908, C. murrayi Remane, 1929, C. mutinensis Balsamo, 1978, C. naiadis Balsamo & Todaro, 1995, C. napoleonicus Balsamo, Todaro & Tongiorgi, 1992, C. neptuni (Wilke, 1954), C. novenarius Greuter, 1917, C. oceanides (d’Hondt, 1971), C. octonarius Stokes, 1887, C. oculatus Schwank, 1990, C. oculifer Kisielewski, 1981, C. odontopharynx Grosso & Drahg, 1986, C. oligohalinus (Hummon, 1974), C. oplites Balsamo, Fregni & Tongiorgi, 1994, C. ornatus Daday, 1897, C. paluster d’Hondt, 1967, C. palustris Anderson & Robbins, 1980, C. parafurcatus Nesteruk, 1991, C. paraguayensis Schwank, 1990, C. paucisetosus Marcolongo, 1910, C. paucisquamatus Kisielewski, 1991, C. pawlowskii Kisielewski, 1984, C. pentacanthus Balsamo, 1981, C. persetosus (Zelinka, 1889), C. pilaga Grosso, 1982, C. ploenensis Voigt, 1909, C. polychaetus Daday, 1905, C. polyhybus Hummon, 2010,

 117

C. polyspinosus Greuter, 1917, C. poznaniensis Kisielewski, 1981, C. pratensis Schwank, 1990, C. pravus Kolicka, Kisielewski, Nesteruk & Zawierucha, 2013, C. pseudopolyspinosus Kisielewski, 1991, C. pungens Balsamo, 1990, C. puniceus Martin, 1990, C. pusillus Daday, 1905, C. quadratus Martin, 1981, C. quintospinosus Greuter, 1917, C. rafalskii Kisielewski, 1979, C. rarispinosus Roszczak, 1936, C. rectaculeatus Kisielewska, 1981, C. remanei Schwank, 1990, C. retiformis Suzuki & Furuya, 2011, C. rhombosquamatus Kolicka, Kisielewski, Nesteruk & Zawierucha, 2013, C. robustus Davison, 1938, C. rotundus Greuter, 1917, C. sagittarius (Evans, 1992), C. sanctipauli Kisielewski, 1991, C. schlitzensis Schwank, 1990, C. schoepferae Thane-Fenchel, 1970, C. schromi Hummon, 1974, C. schultzei (Metschnikoff, 1865), C. scoticus Schwank, 1990, C. scutatus Saito, 1937, C. scutulatus Martin, 1981, C. segnis Martin, 1981, C. semihamus Hummon, 2010, C. serenus (Schrom, 1972), C. sextospinosus Visvesvara, 1964, C. siciliensis Hummon, Balsamo & Todaro, 1992, C. silvaticus (Varga, 1963), C. similis Zelinka, 1889, C. simrothi Voigt, 1909, C. soberanus Grosso & Drahg, 1983, C. sphagnophilus Kisielewski, 1981, C. spinifer Stokes, 1887, C. spinulosus Stokes, 1887, C. splendidus Preobrajenskaja, 1926, C. stagnalis d’Hondt, 1967, C. striatus Preobrajenskaja, 1926, C. succinctus Voigt, 1902, C. sudeticus Kisielewski, 1984, C. tachyneusticus Brunson, 1948, C. tempestivus Mock, 1979, C. tenuis Grünspan, 1908, C. tenuisquamatus Grosso, 1982, C. triacanthus Todaro, 1994, C. trianguliformis Visvesvara, 1964, C. trichodrymodes Brunson, 1950, C. trichostichodes Brunson, 1950, C. tricuspidatus Schwank, 1990, C. trilineatus Valkanov, 1937, C. triradiatus Rao, 1991, C. trispinosus Balsamo, 1990, C. uncinus Voigt, 1902, C. vargai Rudescu, 1967, C. variosquamatus (Mock, 1979), C. vellosus Martin, 1990, C. ventrochaetus Kisielewski, 1991, C. venustus d’Hondt, 1967, C. veronicae Kånneby, 2013, C. voigti Greuter, 1917, C. vorax Remane, 1936, C. vulgaris Brunson, 1950, C. woodi (Thane-Fenchel, 1970), and C. zelinkai Grünspan, 1908. Genus Fluxiderma d’Hondt, 1974 Characteristic for the species in this genus is the presence of round scales. Usually the scales are in some distance from each other, but in one species, they attach each other. Three species are known from few freshwater localities in Europe and North America: F. concinnum (Stokes, 1887), F. montanum Rudescu, 1967, and F. verrucosum (Roszczak, 1936). Genus Halichaetonotus Remane, 1936 (Fig. 1.86) Originally a subgenus of Chaetonotus (Remane 1936), this taxon received genus level by Schrom (1972). The body is covered with large keeled scales, the keels can extend into short spines. Ventrolaterally, there are paired rows with lamellate spines.

118 

 1 Gastrotricha

Thirty-one marine species have been described from the North Atlantic, Baltic Sea, Mediterranean Sea, Black Sea, Northwestern Pacific, and Australia: H. aculifer (Gerlach, 1953), H. arenarius (d’Hondt, 1971), H. atlanticus Kisielewski, 1988, H. australis Nicholas & Todaro, 2005, H. balticus Kisielewski, 1975, H. bataceus Evans, 1992, H. batillifer (Luporini & Tongiorgi, 1972), H. clavicornis Balsamo, Fregni & Tongiorgi, 1995, H. decipiens (Remane, 1929), H. etrolomus Hummon, Balsamo & Todaro, 1992, H. euromarinus Hummon & Todaro, 2010, H. genatus Balsamo, Fregni

& Tongiorgi, 1995, H. italicus Balsamo, Hummon, Todaro & Tongiorgi, 1997, H. jucundus (d’Hondt, 1971), H. lamellatus Kisielewski, 1975, H. littoralis (d’Hondt, 1971), H. margaretae Hummon, Balsamo & Todaro, 1992, H. marivagus Balsamo, Todaro & Tongiorgi, 1992, H. paradoxus (Remane, 1927), H. parvus (Wilke, 1954), H. pleuracanthus (Remane, 1926), H. polonensis Hummon, 2008, H. riedli Schrom, 1972, H. sanctaeluciae Todaro, Dal Zotto, Perissinotto & Bownes, 2011, H. schromi Kisielewski, 1975, H. somniculosus (Mock, 1979), H. susi Hummon, 2010, H. swedmarki Schrom, 1972, H. tentaculatus (d’Hondt, 1971), H. testiculophorus (Hummon, 1966), and H. thalassopais Hummon, Balsamo & Todaro, 1992. Genus Heterolepidoderma Remane, 1927 (Fig. 1.87 A, B) Species in this genus are densely covered with keeled scales, the keels often create a longitudinally striated appearance. There is some nomenclatorial discussion (d’Hondt et al. 2010, Hummon & Todaro 2010). Thirty-four species have been described, 14 from marine or brackish environments (Hummon & Todaro 2010 and newer descriptions) and 21 from freshwater environments (Balsamo et  al. 2009 and newer descriptions); some species, especially H. ocellatum, seem to have some tolerance for salinity and occur in brackish and freshwater

lsp

A

20 µm

B

po 40 µm 50 µm Fig. 1.86: Halichaetonotus sp. (Paucitubulatina, Chaetonotidae) from sublittoral sand of the North Sea off the Danish coast at depth of 25 m. Ventral view. Abbreviation: lsp, lamellar spine. BF image.

Fig. 1.87: (A) Heterolepidoderma joermungandri (Paucitubulatina, Chaetonotidae) from peat bog (Sphagnum spp.) in a rock pool at Skarvesäter, Sweden. Dorsal view with focus on the keeled scales. (B) H. ocellatum from peat bog in a rock pool at Kristineberg, Sweden. Dorsal view. Abbreviation: po, pseudocellus. (A and B, DIC images were kindly provided by Tobias Kånneby, Stockholm.)

1.6 Systematics 

habitats (see Hummon & Todaro 2010). Species are H. acidophilum Kånneby, Todaro & Jondelius, 2011, H. arenosum Kisielewski, 1988, H. armatum Schrom, 1966, H. axi Mock, 1979, H. brevitubulatum Kisielewski, 1981, H. baium Hummon, 2010, H. caudosquamatum Grilli, Kristensen & Balsamo, 2009, H. clipeatum Schrom, 1972, H. contectum Schrom, 1972, H. dimentmani Kisielewski, 1999, H. famaillense Grosso & Drahg, 1991, H. foliatum Renaud-Mornant, 1967, H. gracile Remane, 1927, H. grandiculum Mock, 1979, H. hermaphroditum Wilke, 1954, H. illinoisense Robbins, 1965, H. istrianum Schrom, 1972, H. joermungandri Kånneby, 2011, H. jureiense Kisielewski, 1991, H. kossinense (Preobrajenskaja, 1926), H. lamellatum Balsamo & Fregni, 1995, H. longicaudatum Kisielewski, 1979, H. loricatum Schrom, 1972, H. macrops Kisielewski, 1981, H. majus Remane, 1927, H. marinum Remane, 1926, H. multiseriatum Balsamo, 1978, H. obesum d’Hondt, 1967, H. obliquum Saito, 1937, H. ocellatum (Metschnikoff, 1865), H. patella Schwank, 1990, H. pineisquamatum Balsamo, 1981, H. tenuisquamatum Kisielewski, 1981, and H. trapezoidum Kånneby, 2011.

Genus Ichthydium Ehrenberg, 1830 (Fig. 1.88 A, B) In species of this genus, cuticular structures are either completely absent or strongly reduced and the cuticle appears “naked” (Fig. 1.88 A, B). Twenty-nine species have been described, two from the marine environment (Mediterranean Sea: I. cyclocephalum

A

 119

and I. tergestinum) and the majority from freshwater environments worldwide. The species I. podura may be found in marine and freshwater (see Hummon & Todaro 2010). For a key and further information, see Schwank (1990) and Kånneby et al. (2009). Species are I. auritum Brunson, 1950, I. balatonicum Varga, 1950, I. bifasciale Schwank, 1990, I. bifurcatum Preobrajenskaja, 1926, I. brachykolon Brunson, 1949, I. cephalobares Brunson, 1949, I. chaetiferum Kisielewski, 1991, I. crassum Daday, 1905, I. cyclocephalum (Grünspan, 1908), I. diacanthum Balsamo & Todaro, 1995, I. dubium Preobrajenskaja, 1926, I. forcipatum Voigt, 1902, I. forficula Remane, 1927, I. fossae d’Hondt, 1972, I. galeatum Konsuloff, 1921, I. leptum Brunson, 1949, I. macrocapitatum Sudzuki, 1971, I. macropharyngistum Brunson, 1949, I. maximum Greuter, 1917, I. minimum Brunson, 1950, I. palustre Kisielewski, 1981, I. pellucidum Preobrajenskaja, 1926, I. plicatum Balsamo & Fregni, 1995, I. podura (Müller, 1773), I. rostrum Roszczak, 1969, I. squamigerum Balsamo & Fregni, 1995, I. sulcatum (Stokes, 1887), I. tanytrichum Balsamo, 1983, and I. tergestinum (Grünspan, 1908). Genus Lepidodermella Blake, 1933 (Fig. 1.89 A, B) The genus was originally introduced as Lepidoderma by Zelinka (1889), but as this name was preoccupied by slime molds(Myxogastria),Blake(1933)renameditLepidodermella.

A mo

40 µm

B

50 µm

B 50 µm

Fig. 1.88: (A) Ichthydium squamigerum (Paucitubulatina, Chaetonotidae) from peat bog (Sphagnum spp.) in a rock pool at Skarvesäter, Sweden. Horizontal view. (B) I. skandicum from an artificial pond on Skaftö, Sweden. Horizontal view. Note the naked cuticle in both species. (A and B, DIC images were kindly provided by Tobias Kånneby, Stockholm.)

50 µm Fig. 1.89: Lepidodermella squamata (Paucitubulatina, Chaetonotidae) from a drainage ditch in Oldenburg, Germany. (A) Ventral view. (B) Dorsal view showing the tile-like scales. Abbreviation: mo, mature egg. (A and B) DIC images.

120 

 1 Gastrotricha

The body is covered densely with attaching or overlapping simple scales without spines. Thirteen species have been described, one of them from the marine environment (Mediterranean Sea: L. limogena), the other species from freshwater. Lepidodermella squamata is one of the most well-known gastrotrich species. A key to species is given in Schwank (1990). The known distribution of the genus is in Europe, North and South America, India, and Japan: L. acantholepida Suzuki, Maeda & Furuya, 2013, L. amazonica Kisielewski, 1991, L. aspidioformis Sudzuki, 1971, L. broa Kisielewski, 1991, L. limogena Schrom, 1972, L. macrocephala d’Hondt, 1972, L. minor (Remane, 1936), L. serrata Sudzuki, 1971, L. spinifera Tretjakova, 1991, L. squamata (Dujardin, 1841), L. tabulata (Preobrajenskaja, 1926), L. triloba (Brunson, 1950), and L. zelinkai (Konsuloff, 1913). Genus Polymerurus Remane, 1926 (Fig. 1.90) This genus includes large specimens, the maximal size is up to 650 µm (all data from Schwank 1990). The head is

not clearly separated from the body. The cuticular plates in the head region are more or less well developed, usually a cephalion, a slender hypostomium, and one pair of pleuria are present. Two bundles of cilia are present in the head. The toes are extremely long, and the terminal adhesive tubes are strongly reduced (Fig. 1.90). In some species, the toes are covered with spined scales; in others, the scales are fused to form rings that give the toes a segmented appearance. The body is covered with simple scales of different shapes or with spines. Minute stalked spines are present in the species P. rhomboides. The pharynx is short and without a bulbus. A key to species is given in Schwank (1990). Eighteen species have been described from all continents except Antarctica, some species (P. nodicaudus, P. rhomboides) appear to be cosmopolites: P. andreae Hochberg, 2005, P. biroi (Daday, 1897), P. callosus Brunson, 1950, P. corumbensis Kisielewski, 1991, P. elongatus (Daday, 1905), P. entzii (Daday, 1882), P. hystrix (Daday, 1910), P. longicaudatus (Tatem, 1867), P. macracanthus (Lauterborn, 1894), P. macrurus (Collin, 1897), P. magnus Visvesvara, 1963, P. nodicaudus (Voigt, 1901), P. nodifurca (Marcolongo, 1910), P. paraelongatus Grosso & Drahg, 1986, P. rhomboides (Stokes, 1887), P. ringueleti Grosso, 1975, P. serraticaudus (Voigt, 1901), and P. squammofurcatus (Preobrajenskaja, 1926).

Genus Rhomballichthys Schwank, 1990 The characteristic feature of species in this genus is that the body is densely covered with rhomboidal scales that closely attach each other. On the surface, further structures such as short spines or secondary scales can be present (see Schwank 1990). Schwank (1990) includes three species in this genus, but Balsamo et  al. (2009) regard two of these species as species inquirenda (R. carinatus Schwank, 1990, and R. murrayi Schwank, 1990). The only remaining species is R. punctatus (Greuter, 1917). The species has a scattered distribution in Europe.

ce

50 µm

fu

Fig. 1.90: Polymerurus nodicaudus (Paucitubulatina, Chaetonotidae) from a ditch in northern Germany. Horizontal view. Note the annulated furca. Abbreviations: ce, cephalion; fu, furca. DIC image.

Genus Undula Kisielewski, 1991 The genus was created by Kisielewski (1991) for one conspicuous species, Undula paraënsis Kisielewski, 1991, from Brazil. It has a furca, but instead of adhesive tubes, the furcal branches end with a spine. The name-giving feature is an undulating band of cilia in the head region that ends in paired ventral tufts of cilia. Some spines are present in the posterior region of the trunk. The body is covered with very small and inconspicuous scales (see Kisielewski 1991 for more details).

1.6 Systematics 

1.6.2.2.4 Family Dasydytidae Daday, 1905 In most species, the head is clearly delimited and often has lobes. Dasydytids are pelagic and during swimming the head is held downward (Schwank 1990). A furca is absent, the cuticle is usually naked (but see especially different species from the tropical South America described by Kisielewski 1991), with the exception of several more or less long spines that either originate directly from the cuticle or from scale rudiments. Spines often occur in bundles and may sometimes be very long and movable; they are used for locomotion and defense (see Kieneke et  al. 2008b, Kieneke & Ostmann 2012). With exception of Halidytes, the ventral ciliation is in bundles. A putative marine dasydytid species, Metadasydytes quadrimaculatus (Roszczak 1971), is not a gastrotrich, but a polychaete larva (Hummon 2008b). All species live in freshwater. Eight genera are included in this family: Anacanthoderma, Chitonodytes, Dasydytes, Dichaetura, Ornamentula, Haltidytes, Setopus, and Stylochaeta.

 121

*

ms

Genus Chitonodytes Remane, 1936 On the lateral sides of the animals, bundles of long spines originate, and these extend behind the posterior end. The spines are either “simple”, meaning without lateral extensions, or they have one or two such extensions called denticles. The denticles are directed toward the animal. Three species are known from Europe: C. collini (Remane, 1927), C. longisetosus (Metschnikoff, 1865), and C. longispinosus (Greuter, 1917). Genus Dasydytes Gosse, 1851 (Fig. 1.91) The head is well delimited, and the trunk carries a large number of spines, which are grouped in bundles or as single spines. Spines have one lateral denticle and a terminal bifurcation. There are no appendages at the posterior end. The pharynx has no bulbs. When present, scales covering the body are keeled. Schwank (1990) divided the genus Dasydytes into three subgenera: Setopus, Setodytes, and Dasydytes. A year later, Kisielewski (1991) regarded Setopus to be at genus level, including the former Setodytes species. Within Dasydytes, he recognized three subgenera: Dasydytes, Prodasydytes, and Chitonodytes.

50 µm

Genus Anacanthoderma Marcolongo, 1910 The head is not delimited as in the other taxa of Dasydytidae. In the posterior region of the trunk are few (2–16) spines that are not movable. The pharynx has two bulbi; the anus is ventral. Two not well-known species were described from Italy and Romania: A. paucisetosum (Marcolongo, 1910) and A. punctatum Marcolongo, 1910.

Fig. 1.91: Dasydytes goniathrix (Paucitubulatina, Dasydytidae) from a ditch close to Oldenburg, Germany. Horizontal view. Note the rings of locomotor cilia on the head (asterisk). Abbreviation: ms, motile spines. DIC image.

At present, Chitonodytes is regarded as a genus and Dasydytes comprises the two subgenera Dasydytes and Prodasydytes (Balsamo et al. 2009). For the characterization of subgenera, see Kisielewski (1991). Nine species are currently included in this genus; they have been found in Europe, the USA, and Brazil: D. asymmetricus Schwank, 1990, D. carvalhoae Kisielewski, 1991, D. elongatus Kisielewski, 1991, D. goniathrix Gosse, 1851, D. lamellatus Kisielewski, 1991, D. monile Horlick, 1975, D. nhumirimensis Kisielewski, 1991, D. ornatus Voigt, 1909, and D. papaveroi Kisielewski, 1991.

122 

 1 Gastrotricha

Genus Haltidytes Remane, 1936 (Fig. 1.92) Species have one to three pairs of ventrolateral originating long jumping spines, in relaxed state they cross over the trunk (Fig. 1.92). There are further spines originating from the transition of the neck to the trunk. All spines are simple. The ventral cilia are in paired rows, which is unusual for this family. The pharynx has no bulbus. Five species have been described from Europe, North America, Central Russia, Argentina, and Brazil: H. crassus (Greuter, 1917), H. festinans (Voigt, 1909), H. ooëides Brunson, 1950, H. saltitans (Stokes, 1887), and H. squamosus Kisielewski, 1991. Genus Ornamentula Kisielewski 1991 The body is covered with well-developed and finely ornamented scales, and these are very large in the dorsal and lateral sides; ventrally, there are very small spined scales in the posterior half of the animal. Smaller scales are also present on the head. Long spines with a lateral denticle originate from the lateral scales (Kisielewski 1991).

One species is known from Brazil: O. paraënsis Kisielewski, 1991. Genus Setopus Grünspan, 1908 (Fig. 1.93 A, B) There is a pair of spines on the bilobed caudal trunk end. These terminal spines may be equal or unequal (Fig. 1.93 B) in length. The lateral spines occur in three to six paired groups or pairs of single spines. Spines are simple or may have one small lateral denticle. Some species have scales, and these are dorsally larger than ventral; the dorsal scales are incompletely keeled, whereas the ventral scales are keeled or spined (see Kisielewski 1991). Eight species are described from Europe, Brazil, India, Central Russia, and probably the USA: S. abarbitus (Visvesvara, 1964), S. aequatorialis Kisielewski, 1991, S. bisetosus (Thompson, 1891), S. chatticus (Schwank, 1990), S. dubius (Voigt, 1909), S. iunctus Greuter, 1917, S. primus Grünspan, 1908, and S. tongiorgii (Balsamo, 1983). Genus Stylochaeta Hlava, 1904 (Fig. 1.94) Paired club-like extensions (protuberances) with few fine bristles (cilia) are present in the caudal end. The lateral spines are long, and they have a sharp tip and two to three lateral denticles. The pharynx has one bulb.

*

ms

*

*

Fig. 1.92: Haltidytes crassus (Paucitubulatina, Dasydytidae) from a drainage ditch in Oldenburg, Germany. Ventral view. Note the rings of locomotor cilia on the head (asterisk). The most posterior pair of motile spines shows a characteristic crossing. Abbreviation: ms, motile spines. BF image.

ts

50 µm

50 µm

ms

50 µm

ms

Fig. 1.93: (A and B) Setopus tongiorgii (Paucitubulatina, Dasydytidae) from a drainage ditch in Oldenburg, Germany. A and B at slightly different focus. Note the rings of locomotor cilia on the head (asterisks) and the unequal length of the terminal spines. Abbreviation: ms, motile spines; ts, terminal spines. (A and B) DIC images.

1.6 Systematics 

Four species are known from Europe, Central Russia, and North America: S. fusiformis (Spencer, 1890), S. longispinosa Greuter, 1917, S. scirtetica Brunson, 1950, and S. stylifera (Voigt, 1901). 1.6.2.2.5 Family Neogosseidae Remane 1927 The most conspicuous character for members of this family is the presence of club-shaped tentacles on the head. A caudal furca and therefore adhesive tubes are absent. Locomotory cilia are present as short bands and bundles (tufts) around the head and along the ventral surface of the trunk. Most, but not, all species have spined scales, toward the posterior end the spines are usually longer. The mouth is surrounded by a mouth ring. The male reproductive system is unknown. The pharynx has between one and four bulbs. All species live planktonic or semipelagic in freshwater. For more information see Schwank (1990), Kisielewski (1991), and Todaro et al. (2013). Two genera are included in this family, Kijanebalola and Neogossea. Genus Kijanebalola Beauchamp, 1932 In the posterior end, a median group of spines is present. The body is covered by keeled scales with rudimentary

 123

spines, naked regions may occur. The pharynx has up to two bulbs, the mouth opening is surrounded by a cuticular ring. For more characters, see Todaro et al. (2013). Three species have been described from Lake Kijanebalola in Uganda and freshwater habitats in South Africa and Brazil: K. canina Kisielewski, 1991, K. devestiva Todaro, Perissinotto & Bownes, 2013, and K. dubia de Beauchamp, 1932. Genus Neogossea Remane, 1927 (Fig. 1.95) In the posterior end are small posterolateral projections carrying tufts of long spines. The body is covered by spined scales. The mouth opening is terminal, surrounded by a large, protruding cuticular ring with longitudinal ridges and spine-like structures. The pharynx has four bulbs, the posterior one is the largest (characters from Todaro et al. 2013). Six species are known from freshwaters in Brazil, Paraguay, South Africa, India, North America, and Central Europe, extending eastward to Central Russia and the Caspian Region: N. acanthocolla Kisielewski, 1991, N. antennigera (Gosse, 1851), N. fasciculata (Daday, 1905),

ct

ph

* ms

Fig. 1.94: Stylochaeta scirtetica (Paucitubulatina, Dasydytidae) from a ditch close to Leer, Germany. Ventral view. Note the rings of locomotor cilia on the head (asterisk). The first group of spines is spread. Abbreviations: ms, motile spines; sty, styli. DIC image.

ts

50 µm

sty

50 µm

ms

Fig. 1.95: Neogossea voigti (Paucitubulatina, Neogosseidae) from a ditch close to Leer, Germany. Horizontal view. Note that the animal was nodding with its head. Abbreviations: ct, club-shaped tentacle; ph, pharynx (with distinct bulbs); ts, terminal spines. DIC image.

124 

 1 Gastrotricha

N. pauciseta (Daday, 1905), N. sexiseta Krivanek & Krivanek, 1959, and N. voigti (Daday, 1905). 1.6.2.2.6 [Family Dichaeturidae Remane, 1927] Genus Dichaetura Lauterborn, 1913 (Fig. 1.96) The head region is without appendages or head plates. Some bristles or cilia and few spines appear to be present. Most characteristic is the presence of two pairs of adhesive tubes on the furca, a condition otherwise only reported for Diuronotus. However, Todaro et  al. (2005) point out that the second adhesive tube may also be a lateral extension instead of a tube, which was confirmed in the description of the fourth species, D. filispina, by Suzuki et  al. (2013). A reinvestigation of the other species would be very helpful. Only very few scales are present. Four species were found in few places in Europe and in Japan: D. capricornia (Metschnikoff, 1865), D. filispina Suzuki, Maeda & Furuya, 2013, D. piscator (Murray, 1913), and D. surreyi Martin, 1990. 1.6.2.2.7 Family Proichthydidae Remane 1927 Only two species in two genera have been described from this family. The head is well separated from the trunk. Tentacles, plates, or scales are absent. A terminal furca with slightly curved brances is present. Genus Proichthydium Cordero, 1918 Cilia have only been described as a ring around the head, and no further cilia or cuticular structures have been reported (Cordero 1918). One species, P. coronatum Cordero, 1918 is known from freshwater near Montevideo, Uruguay. Genus Proichthydioides Sudzuki, 1971 In the head region, several cilia, some of considerable length, are present. Two longitudinal bands of cilia extend to the base of the furca. One species, P. remanei Sudzuki, 1971, has been described from Japan.

1.6.2.3 Gastrotricha incertae sedis Genus Marinellina Ruttner-Kolisko, 1955 Ruttner-Kolisko (1955) described one species, Marinellina flagellata, from an Austrian river, Ybbs. It has a slight constriction behind a head region and caudal feet with two adhesive tubes per foot. One further pair of adhesive tubes is present in the head region. There are several single long cilia all over the body, but a ventral ciliation appears to be lacking. The pharynx is large, and the intestine is divided into a smaller anterior and a larger posterior part. The specimens described may be juvenile. Because of the number and distribution of adhesive tubes, Marinellina is usually treated as a freshwater macrodasyid gastrotrich.

1.7 Biogeography Gastrotrichs have a worldwide distribution (see Artois et  al. 2011), they have been found on all continents exclusive Antarctica and in marine environments from the shore to the deep sea. Knowledge on gastrotrich diversity and biogeography has, however, still to be regarded as fragmentary. Europe and North and South America have been sampled more intensively than, for example, Africa, Southeast Asia, or Australia (see Balsamo et al. 2008 for more information). Some examples for local summaries of the gastrotrich fauna are Kisielewski (1991) for Brazilian freshwater, Vanamala Naidu & Chandrasekahara Rao (2004) for Indian freshwater and marine regions, and Schmidt (1974) for the marine region around Galapagos. Currently, freshwater species are described from Japan (e.g., Suzuki et  al. 2013) and marine species from South Korean waters (see Lee et al. 2013 and references therein); some investigations deal with the Caribbean fauna (e.g., Hummon 2010, Hochberg et  al. 2014) or discover the Eastern Mediterranean and the Red Sea (Hummon 2011) or South Africa (Todaro et al. 2011b, 2013).

20 µm

Fig. 1.96: Dichaetura filispina (Paucitubulatina, Dichaeturidae) from a rice paddy close to Mano, Japan. Ventral view. (DIC image was kindly provided by Takahito Suzuki, Osaka.)

1.8 Ecology 

In freshwater species as well as in marine species, some species appear to have a wide distribution, being found in disjunct regions. Other species have been found with a very limited distribution. The standard question behind a wide distribution range is whether the observed populations really belong to one single species or whether they represent several cryptic species. In some cases, a closer look or advanced observation technology reveal differences. Leasi & Todaro (2009) investigated specimens of Xenotrichula intermedia from populations in the Mediterranean (Adriatic Sea) and in the Arabian Gulf (Kuwait). Externally, the specimens could not be separated, but a phalloidin staining of actin revealed clearly different patterns of the musculature. The standard tool to aid the recognition of species boundaries is the “barcoding”, a comparison of gene sequences, preferably the cytochrome oxidase I (COI) gene. Such investigations were to date conducted only rarely with gastrotrichs. Todaro et  al. (1996) made some pioneer work with the above mentioned Xenotrichula intermedia and compared three populations, from the Mediterranean (Italy), Southwestern Atlantic (Virginia), and the Gulf of Mexico (Florida). Morphologically, only the pharynx length differed slightly between populations, but genetically (RFLPs, restriction fragment length polymorphism and partial COI gene sequence), four populations were strongly separated from each other. Therefore, four cryptic species seem to be present in these three sites. Recently, Kieneke et al. (2012) discovered evidence for two cryptic genetic species within the morphospecies Turbanella hyalina when studying mitochondrial COI sequences of specimens from different European coastal sites. In general, geographically separated populations even at a rather regional spatial scale showed a deep genetic substructure as in the species T. cornuta (Kieneke et al. 2012).

1.8 Ecology Gastrotrichs are important components of aquatic habitats and can reach considerable number. Densities are usually extrapolated from small samples to larger volumes and have to be taken with some care. For example, Nesteruk (1996) gives values between 495,000 and 2,600,000 specimens per m2 for freshwater standing waters. Muschiol & Traunspurger (2009) found densities of 0.67 × 106 specimens of Chaetonotus sp. per m2 in a lake on the Galapagos Islands. In the marine environment, gastrotrichs form an important part of the meiofaunal community, but their density depends on the grain size. On sandy beaches, Schmidt & Teuchert (1969) counted almost 1000

 125

specimens/50 cm-3. In muddy sediments in the Adriatic Sea (Mediterranean), Leasi & Todaro (2010) found a low diversity and density of gastrotrichs, ranging between 0 and 97.7 specimens per 10 cm2. Unusual high densities are known, for example, in the marine species Turbanella hyalina (Hummon 1976). Such high population densities may be correlated with a high bacterial abundance due to discharge of untreated sewage water close to the habitat (Hummon & Hummon 1993). In the food web, gastrotrichs have a low trophic level, feeding on bacteria, small algae, or detritus (e.g., Bennett 1979, Balsamo & Todaro 2002, Todaro & Hummon 2008). Gastrotrichs themselves are probably food of predators of about the same size class or may serve, together with other meiobenthic organisms, as nutrition resource for large, sediment-consuming animals of the infaunal macrobenthos (e.g., Giere 2009). Because of the latter connection, gastrotrichs and other meiobenthic animals are considered to be an important link between the microbial loop and higher trophic levels (Balsamo & Todaro 2002). Measured in abundance (individuals per area), Gastrotricha may rank second to fourth in numerical dominance among all micrometazoans (e.g., Hummon 1976). Based on his studies of gastrotrich communities of two Scottish beaches, Hummon (1976) was able to estimate the mean dry weight biomass of Gastrotricha in those biotopes. Values ranged between 48 and 274 mg/m2 while the highest station values ranged between 0.6 and 3.1 g/m2. All marine and many freshwater gastrotrichs are associated with the sediment and live either in the epibenthic layer or in the sediment in the pore system (interstitial system). A number of freshwater species is associated with aquatic vegetation and moves among plants. Some species are capable of short swimming periods, such as Heterolepidoderma sp. (Bancetti & Ricci 1998), but only species of Dasydytidae and Neogosseidae are swimming permanently and can hence be regarded as belonging to the planktonic community. Among freshwater, most chaetonotoid gastrotrichs prefer slow or no water motion. After Ricci & Balsamo (2000), from 250 freshwater species only 35 are found in running (lotic) waters. A number of species appears to be quite tolerant to low oxygen content (Schwank 1990) and a number of species is found in moors and tolerates pH values down to 4 (Kisielewski 1981, Schwank 1990). Kisielewski (1981) found the species diversity and abundance of specimens highest in eutropicated peat bogs. Hummon & Hummon (1979) were able to cultivate Lepidodermella squammata in acid mine water with different contents of carbonate. Survival and reproduction were positively correlated with the carbonate content.

126 

 1 Gastrotricha

Freshwater habitats are often fragile and ephemeral and several freshwater gastrotrichs have the ability to build resting eggs (Ricci & Balsamo 2000, see also chapters Gametes and Reproductive Biology). Kisielewski (1981) found different potentials of species to reconstitute their population after dry periods, after longer periods, only two species, Lepidodermella squamata and Chaetonotus maximus, were able to recover their population. In marine habitats, gastrotrichs are almost always associated with the interstitial system, this is the system of pores among sediment grains. The majority of species has been found in the littoral region to the shallow sublittoral (see, e.g., Todaro et al. 2000b), but species occur down to deep sea (Kieneke & Zekely 2007, Kieneke 2010 and references therein). Beaches have been investigated most intensively for distribution patterns of gastrotrichs. Schmidt & Teuchert (1969) observed on an intertidal beach on the island of Sylt (North Sea) that each species inhabits a special position on the beach in relation to the low and high water lines. Comparable patterns have been found for different species of marine Paucitubulatina inhabiting different zones of beach slopes around the island Sylt, North Sea (Mock 1979). On other beaches, some species appear to be spatially restricted, while others are broadly distributed. For example, Hummon (1972) found on a beach on San Juan Archipelago (Washington) only three species; two of them, Chaetonotus testiculophorus and Turbanella cornuta, were found (in different densities) throughout the beach. Similarly, Schmidt (1974) found on Galapagos that Macrodasys pacificus is distributed throughout most of the beach, while other species are more restricted in their distribution. Most specimens occur in the upper few centimetres (Schmidt & Teuchert 1969) and seem to prefer well-oxygenated sediment, but some species are tolerant for low oxygen content and occur in deeper layers of the sediment (see chapter Physiology). There may be migrations in the sediment during different life stages, and this has, for example, been observed by Hochberg (1999) for Turbanella mustela on a Californian beach. Size classes of this species showed a preference for particular sediment layers, with adults occurring more deeply than juveniles. In the sediment, gastrotrichs are characteristically gregarious and occur in small patches (Boaden 1985). For some species, wave action appears to be an important factor and they occur in regions with high wave energy and high sediment turnover. Xenotrichulids prefer these regions (Ruppert 1979). On a beach close to Venice (Adriatic Sea), Schrom (1966) was able to make a rough correlation between the number of adhesive tubes and wave action and sediment type. In coarse sediment and wave action, Macrodasys caudatus, a large species with numerous adhesive tubes, was most abundant. Turbanella cornuta, a smaller

species with also a large number of adhesive tubes, inhabited medium-exposed sediments, and Paraturbanella dohrni, a species with much fewer adhesive tubes, inhabited tidal ponds with fine sediment. Also, Hummon (1975) concluded that hydrodynamic cycles, seasonal, lunar-phasic tidal, semidiurnal tidal, or localized, can affect dramatic changes in the distribution of Gastrotricha in a marine beach.

Acknowledgments We are very thankful to the Gastrotricha community, especially to our colleagues Rick Hochberg and Birgen Holger Rothe, for their collaboration in several projects. Maria Balsamo, Loretta Guidi, Tobias Kanneby, Takahito Suzuki, and Andreas Hejnol were so kind to send us pictures of representatives we were lacking. Many thanks also to Corinna Schulze for her help and to Pedro Martínez Arbizu for giving AK the opportunity to work on this book chapter.

Literature Ahlrichs, W. H. (1995): Ultrastruktur und Phylogenie von Seison nebaliae (Grube 1859) und Seison markiertes Wort nicht kursiv (Claus 1876) – Hypothesen zu phylogenetischen Verwandtschaftsverhältnissen innerhalb der Bilateria. Cuvillier, Göttingen. Amato, A. J. & Weiss, M. J. (1982): Developmental flexibility in the cuticular pattern of a cell-constant organism, Lepidodermella squammata (Gastrotricha). Trans. Am. Microsc. Soc. 101: 229–240. Artois, T., Fontaneto, D., Hummon, W. D., McInnes, S. J., Todaro, M. A., Sørensen, M. V. & Zullini, A. (2011): Ubiquity of microscopic animals? Evidence from the morphological approach in species identification. In: Fontaneto, D. (ed.) Biogeography of microscopic organisms. Is everything small everywhere? Pp. 244–283. Syst. Assoc. Spec. Vol. 79, Cambridge University Press, Cambridge. Atherton, S. (2014): Urodasys poculostylis sp. nov., a new styletbearing gastrotrich (Macrodasyida) from Capron Shoal, Florida. Mar. Biol. Res. 10: 530–536. Atherton, S. & Hochberg R. (2012): Tetranchyroderma bronchostylus sp. nov., the first known gastrotrich (Gastrotricha) with a sclerotic canal in the caudal organ. Mar. Biol. Res. 8: 885–892. Ax, P. (1966): Die Bedeutung der interstitiellen Sandfauna für allgemeine Probleme der Systematik, Ökologie und Biologie. Veröff. Inst. Meeresforsch. Bremerhaven, Spec. Vol. 2: 15–66. Ax, P. (2003): Multicellular Animals.Vol. III. Springer, Berlin. Balsamo, M. (1992): Hermaphroditism and parthenogenesis in lower Bilateria: Gnathostomulida and Gastrotricha. In: Dallai, R. (ed.) Sex origin and evolution. Selected Symposia and Monographs UZI 6: 309–327. Balsamo, M. & Todaro, M. A. (1987): Aspidiophorus polystictos, a new marine species (Gastrotricha, Chaetonotida) and its life cycle. Boll. Zool. 54: 147–153.

Literature 

Balsamo, M. & Todaro, M. A. (1988): Life history traits of two Chaetonotids (Gastrotricha) under different experimental conditions. Invertebr. Reprod. Dev. 14: 161–176. Balsamo, M. & Todaro, M. A. (2002): Gastrotricha. In: Rundle, S. D., Robertson, A. L. & Schmid-Araya, J. M. (eds.) Freshwater meiofauna: biology and ecology, pp. 45–61. Backhuys Publishers, Leiden. Balsamo, M., Fregni, M. & Ferraguti M. (1999): Gastrotricha. In: Adiyodi, K. G. & Adiyodi R. G. (eds.) Reproductive biology of invertebrates. Vol. IX: progress in male gamete ultrastructure and phylogeny. J. Wiley & Sons Ltd., Chichester, pp. 171–191. Balsamo, M., Ferraguti, M., Guidi, L., Todaro, M. A. & Tongiorgi, P. (2002): Reproductive system and spermatozoa of Paraturbanella teissieri (Gastrotricha, Macrodasyida): implications for sperm transfer modality in Turbanellidae. Zoomorphology 121: 235–241. Balsamo, M., Guidi, L., Pierboni, L., Marotta, R., Todaro, M. A. & Ferraguti, M. (2007): Living without mitochondria: spermatozoa and spermatogenesis in two species of Urodasys (Gastrotricha, Macrodasyida) from dysoxic sediments. Invertebr. Biol. 126: 1–9. Balsamo, M., d’Hondt, J.-L., Kisielewski, J. & Pierboni, L. (2008): Global diversity of gastrotrichs (Gastrotricha) in fresh waters. Hydrobiologia 595: 85–91. Balsamo, M., d’Hondt, J.-L., Pierboni, L. & Grilli, P. (2009): Taxonomic and nomenclatural notes on freshwater Gastrotricha. Zootaxa 2158: 1–19. Balsamo, M., Guidi, L., Ferraguti, M., Pierboni, L. & Kristensen, R. M. (2010a): Diuronotus aspetos (Gastrotricha): new morphological data and description of the spermatozoon. Helgoland Mar. Res. 64: 27–34. Balsamo, M., Guidi, L., Grilli, P. & d’Hondt, J.-L. (2010b): The phylogenetic position and the intraphyletic relationships of Gastrotricha. Bull. Soc. Zool. France 135: 79–91. Balsamo, M., Grilli, P., Guidi, L. & d’Hondt, J.-L. (2014): Gastrotricha – biology, ecology and systematics. Families Dasydytidae, Dichaeturidae, Neogosseidae, Proichthyiidae. Vol. 24 in Dumont, H.J.F. (ed.) Identification guides to the plankton and benthos of inland waters. Backhuys, Leiden. Banchetti, R. & Ricci, N. (1998): The bahavior of Heterolepidoderma sp. (Gastrotricha). Zool. Sci. 15: 131–137. Bartolomaeus, T. & Ax, P. (1992): Protonephridia and Metanephridia – their relation within the Bilateria. Z. Zool. Syst. Evolutionsforsch. 30: 21–45. Bennett, L. W. (1979): Experimental analysis of the trophic ecology of Lepidodermella squamata (Gastrotricha: Chaetonotida) in mixed culture. Trans. Am. Microsc. Soc. 98: 254–260. Blake, C. H. (1933): Nomenclatorial notes on Gastrotricha. Science 77: 606. Bleidorn C. (2007): The role of character loss in phylogenetic reconstruction as exemplified for the Annelida. J. Zool. Syst. Evol. Res. 45: 299–307. Boaden, P. J. S. (1960): Three new gastrotrichs from the Swedish west coast. Cah. Biol. Mar. 1: 397–406. Boaden, P. J. S. (1963): Marine Gastrotricha from the interstitial fauna of some North Wales beaches. Proc. Zool. Soc. Lond. 140: 485–501. Boaden, P. J. S. (1965): Two new interstitial Gastrotricha of the family Thaumastodermatidae. Pubbl. Staz. Zool. Napoli 34: 219–225. Boaden, P. J. S. (1974): Three new thiobiotic Gastrotricha. Cah. Biol. Mar. 15: 367–378.

 127

Boaden, P. J. S. (1985): Why is a gastrotrich? In: Conway Morris, S., George, J. D., Gibson, R. & Platt, H. M. (eds.) The origins and relationships of lower invertebrates. Syst. Assoc. Spec. Vol. 28: 248–260. Boone, M., Bert, W., Claeys, M., Houthoofd, W. & Artois, T. (2011): Spermatogenesis and the structure of the testes in Nemertodermatida. Zoomorphology 130: 273–282. Brandenburg, J. (1962): Elektronenmikroskopische Untersuchung des Terminalaparates von Chaetonotus sp. (Gastrotrichen) als ersten Beispiels einer Cyrtocyte bei Askhelminten. Z. Zellforsch. 57: 136–144. Brunson, R. B. (1949): The life history and ecology of two North America gastrotrichs. Trans. Am. Microsc. Soc. 68: 1–20. Brunson, R. B. (1950): An introduction to the taxonomy of Gastrotricha with a study of eighteen species from Michigan. Trans. Am. Microsc. Soc. 69: 325–352. Bütschli, O. (1876): Untersuchungen über freilebende Nematoden und die Gattung Chaetonotus. Z. Wiss. Zool. 26: 363–413. Cavalier-Smith, T. (1998): A revised six-kingdom system of life. Biol. Rev. 73: 203–266. Chandrasekahara Rao, G. C. (1993): Littoral meiofauna of little Andaman. Rec. Zool. Surv. India 155: 1–120. Chandrasekahara Rao, G. C. & Clausen, C. (1970): Planodasys marginalis gen. et sp. nov. and Planodasyidae fam. nov. (Gastrotricha Macrodasyoidea). Sarsia 42: 73–82. Claparéde, M. E. (1867): Type d’un nouveau genre de gastrotriches. Ann. Sci. Nat. Zool. 8: 16–23. Clausen, C. (1965): New interstitial species of the family Thaumastodermaidae (Gastrotricha, Macrodasyoidea). Sarsia 21: 23–36. Clausen, C. (1968): Crasiella diplura gen. et sp.n. (Gastrotricha, Macrodasyoidea). Sarsia 33: 59–64. Clausen, C. (2004a): A new species of Acanthodasys (Gastrotricha: Thaumastodermatidae) from the west coast of Norway. Sarsia 89: 137–141. Clausen, C. (2004b): Gastrotricha from the Faroe Bank. Sarsia 89: 423–458. Colacino, J. M. & Kraus, D. W. (1984): Hemoglobin-containing cells of Neodasys (Gastrotricha, Chaetonotoida) – II. respiratory significance. Comp. Biochem. Physiol. 79A: 363–369. Cordero, E. H. (1918): Notes sur les Gastrotriches. Phys. Rev. Soc. Arg. Sci. Nat. 4: 241–244. D’Hondt, J.-L. (1971): Gastrotricha. Oceanogr. Mar. Biol. Ann. Rev. 9: 141–192. D’Hondt, J.-L. (1968): Contribution a la connaissance des gastrotriches intercotidaux du Golfe de Gascogne. Cah. Biol. Mar. 9: 387–404. D’Hondt, J.-L. (1974): Note sur Psammodasys cambriensis (Boaden, 1963) et Crasiella oceanica n.sp. (Gastrotriches Macrodasyoides). Bull. Soc. Zool. France 99: 675–680. D’Hondt, J.-L., Balsamo, M. & Grilli, P. (2010): Un cas d’école: l’espèce type du genre Heterolepidoderma Remane (Gastrotriches, Chaetonotidae). Bull. Mens. Soc. Limn. Lyon 79: 91–94. De Beauchamp, P. M. (1929): Le developpement des Gastrotriches. (Note prelimaire). Bull. Soc. Zool. France 54: 549–558. Dunn, C. W., Hejnol, A., Matus, D. Q., Pang, K., Browne, W. E., Smith, S. A., Seaver, E., Rouse, G. W., Obst, M., Edgecombe, G. D., Sørensen, M. V., Haddock, S. H. D., Schmidt-Rhaesa, A., Okusu, A., Kristensen, R. M., Wheeler, W. C., Martindale, M. Q. & Giribet, G. (2008): Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452: 745–750.

128 

 1 Gastrotricha

Eakin, R. M. (1972): Structure of invertebrate photoreceptors. In: Dartnall, H. J. A. (ed.) Photochemistry of vision. Handbook of sensory physiology, pp. 625–684. Springer, Berlin. Ehlers, U., Ahlrichs, W., Lemburg, C. & Schmidt-Rhaesa, A. (1996): Phylogenetic systematization of the Nemathelminthes (Aschelminthes). Verh. Dtsch. Zool. Ges. 89: 8. Ehrenberg, C.-G. (1830): Organisation, Systematik und geographisches Verhältnis der Infusionsthierchen. F. Dümmler, Berlin. Ehrenberg, C.-G. (1838): Die Infusionsthierchen als vollkommene Organismen. Ein Blick in das tiefere organische Leben der Natur. Verlag Leopold Voss, Leipzig. Evans, W. A. (1994): Morphological variability in warm-temperate and subtropical populations of Macrodasys (Gastrotricha: Macrodasyida: Macrodasyidae) with the description of seven new species. Proc. Biol. Soc. Washington 107: 239–255. Evans, W. A. & Hummon, W. D. (1991): A new genus and species of Gastrotricha from the Atlantic coast of Florida, U.S.A. Trans. Am. Microsc. Soc. 110: 321–327. Ferraguti, M. & Balsamo, M. (1994): Sperm morphology and anatomy of the genital organs in Mesodasys laticaudatus Remane, 1951 (Gastrotricha: Macrodasyida). J. Submicrosc. Cytol. Pathol. 26: 21–28. Ferraguti, M. & Balsamo, M. (1995): Comparative spermatology of Gastrotricha. In: Jamieson, B. G. M., Ausio, J. & Justine, J.-L. (eds.) Advances of spermatozoal phylogeny and taxonomy. Mém. Mus. Natl. Hist. Nat. 166: 105–117. Ferraguti, M. & Balsamo, M., Fregni, E. (1995): The spermatozoa of three species of the Xenotrichulidae (Gastrotricha, Chaetonotida): the two “dünne Nebengeisseln” of spermatozoa in Heteroxenotrichula squamosa are peculiar para-acrosomal bodies. Zoomorphology 115: 151–159. Fischer, U. (1994): Ultrastructure of spermatogenesis and spermatozoa of Cephalodasys maximus (Gastrotricha, Macrodasyida). Zoomorphology 114: 213–225. Fischer, U. (1996): Ultrastructure of penetrated spermatozoa, ovary, and oogenesis of Dactylopodola baltica (Gastrotricha, Macrodasyida). Invertebr. Reprod. Dev. 29: 71–78. Forneris, L. (1966): Über zwei Chaetonotus-Arten aus Hamburger Stadtgewässern (Gastrotricha). Abh. Verh. Naturwiss. Ver. Hamburg 10: 93–96. Fregni, E., Balsamo, M. & Ferraguti, M. (1999): Morphology of the reproductive system and spermatozoa of Mesodasys adenotubulatus (Gastrotricha: Macrodasyida). Mar. Biol. 135: 515–520. Garraffoni, A. R. S., Araujo, T. Q., Lourenço, A. P., Balsamo, M. (2010): New data on freshwater psammic Gastrotricha from Brazil. ZooKeys 60: 1–12. Gagné, G. D. (1977): Dolichodasys elongatus n.g., n.sp., a new macrodasyid gastrotrich from New England. Trans. Am. Microsc. Soc. 96: 19–27. Gagné, G. D. (1980a): Ultrastructure of the anterior sensory devices in the phylum Gastrotricha. Am. Zool. 20: 749. Gagné, G. D. (1980b): Ultrastructure of the sensory palps of Tetranchyroderma papii (Gastrotricha, Macrodasyida). Zoomorphologie 95: 115–125. Giere, O. (2009): Meiobenthology. The microscopic motile fauna of aquatic sediments. Springer, Berlin, Heidelberg. Gilbert, S. F. & Singer, S. R. (2006): Developmental Biology. 8th edition. Sinauer, Sunderland.

Giribet, G. (2002): Current advances in the phylogenetic reconstruction of metazoan evolution. A new paradigm for the Cambrian explosion? Mol. Phyl. Evol. 24: 345–357. Giribet, G., Distel, D. L., Polz, M., Sterrer, W. & Wheeler, W. C. (2000): Triploblastic relationships with emphasis on the Acoelomates and the position of Gnathostomulida, Cycliophora, Plathelminthes, and Chaetognatha: a combined approach of 18S rDNA sequences and morphology. Syst. Biol. 49: 539–562. Giribet, G., Dunn, C. W., Edgecombe, G. D. & Rouse, G. W. (2007): A modern look at the animal tree of life. Zootaxa 1668: 61–79. Guidi, L., Marotta, R., Pierboni, L., Ferraguti, M., Todaro, M. A. & Balsamo, M. (2003a): Comparative sperm ultrastructure of Neodasys ciritus and Musellifer delamarei, two species considered to be basal among Chaetonotida (Gastrotricha). Zoomorphology 122: 135–143. Guidi, L., Ferraguti, M., Pierboni, L. & Balsamo, M. (2003b): Spermiogenesis and spermatozoa in Acanthodasys aculeatus (Gastrotricha, Macrodasyida): an ultrastructural study. Acta Zool. 84: 77–85. Guidi, L., Pierboni, L., Ferraguti, M., Todaro, M. A. & Balsamo, M. (2004): Spermatology of the genus Lepidodasys Remane, 1926 (Gastrotricha, Macrodasyida): Towards a revision of the family Lepidodasyidae Remane, 1927. Acta Zool. 85: 211–221. Guidi, L., Ferraguti, M., Todaro, M. A., Pierboni, L. & Balsamo, M. (2009): Unusual spermatozoa and reproductive modalities of Xenodasys eknomios (Gastrotricha: Xenodasyidae). Ital. J. Zool. 76: 165–172. Guidi, L., Todaro, M. A., Ferraguti, M. Balsamo, M. (2011): Reproductive system of the genus Crasiella (Gastrotricha, Macrodasyida). Helgoland Mar. Res. 65: 175–185. Guidi, L., Todaro, M. A., Ferraguti, M. & Balsamo, M. (2014): Reproductive system and spermatozoa ultrastructure support the phylogenetic proximity of Megadasys and Crasiella (Gastrotricha, Macrodasyida). Contrib. Zool. 83: 119–131. Hejnol, A., Obst, M., Stamatakis, A., Ott, M., Rouse, G. W., Edgecombe, G. D., Martinez, P., Baguña, J., Bailly, X., Jondelius, U., Wiens, M., Müller, W. E. G., Seaver, E., Wheeler, W. C., Martindale, M. Q., Giribet, G. & Dunn, C. W. (2009): Assessing the root of bilaterian animals with scalable phylogenomic methods. Proc. R. Soc. B 276: 4261–4270. Hochberg, R. (1998): Postembryonic growth and morphological variability in Turbanella mustela (Gastrotricha, Macrodasyida). J. Morphol. 237: 117–126. Hochberg, R. (1999): Spatiotemporal size-class distribution of Turbanella mustela (Gastrotricha: Macrodasyida) on a northern californian beach and its effect on tidal suspension. Pacific Sci. 53: 50–60. Hochberg, R. (2001): A special form of sensory cilia in Lepidodermella squamata (Gastrotricha: Chaetonotida). Ophelia 55: 137–139. Hochberg, R. (2002): Two new species of Pseudostomella (Gastrotricha: Macrodasyida) with scaled triancres from the east coast of Australia. NZ J. Mar. Freshwater Res. 36: 571–579. Hochberg, R. (2003): Two new species of Dactylopodola (Gastrotricha, Macrodasyida) from the islands off the

Literature 

Queensland Coast, Australia. Meiofauna Mar. 12: 37–45. Hochberg, R. (2005): Musculature of the primitive gastrotrich Neodasys (Chaetonotida): functional adaptations to the interstitial environment and phylogenetic significance. Mar. Biol. 146: 315–323. Hochberg, R. (2007): Comparative immunohistochemistry of the cerebral ganglion in Gastrotricha: an analysis of FMRFamidelike immunoreactivity in Neodasys cirritus (Chaetonotida), Xenodasys riedli and Turbanella cf. hyalina (Macrodasyida). Zoomorphology 126: 245–264. Hochberg, R. (2008): Ultrastructure of feathered triancres in the Thaumastodermatidae and the description of a new species of Tetranchyroderma (Gastrotricha : Macrodasyida) from Australia. J. Mar. Biol. Assoc. UK 88: 729–737. Hochberg, R. (2010a): Two new species of Oregodasys (Gastrotricha: Macrodasyida: Thaumastodermatidae) from Carrie Bow Cay, Belize with ultrastructural observations of the epidermal glandular system. Zootaxa 2660: 1–17. Hochberg, R. (2010b): The epidermal glands of gastrotrichs: ultrastructural insights and hypotheses of function. Integr. Comp. Biol. 50: e76. Hochberg, R. (2014): Crasiella fonseci, a new species of Gastrotricha (Macrodasyida, Planodasyidae) from São Paulo, Brazil. Mar. Biodiv. 44: 237–242. Hochberg, R. & Atherton, S. (2011): A new species of Lepidodasys (Gastrotricha, Macrodasyida) from Panama with a description of its peptidergic nervous system using CLSM, anti-FMRFamide and anti-SCPB. Zool. Anz. 250: 111–122. Hochberg, R. & Litvaitis, M. K. (2000): Phylogeny of Gastrotricha: a morphology-based framework of gastrotrich relationships. Biol. Bull. 198: 299–305. Hochberg, R. & Litvaitis, M. K. (2001a): A muscular double helix in Gastrotricha. Zool. Anz. 240: 61–68. Hochberg, R. & Litvaitis, M. (2001b): The muscular system of Dactylopodola baltica and other macrodasyidan gastrotrichs in a functional and phylogenetic perspective. Zool. Scr. 30: 325–336. Hochberg, R. & Litvaitis, M. (2001c): Functional morphology of muscles in Tetranchyroderma papii (Gastrotricha). Zoomorphology 121: 37–43. Hochberg, R. & Litvaitis, M. (2001d): The musculature of Draculiciteria tesselata (Chaetonotida, Paucitubulatina): implications for the evolution of dorsoventral muscles in Gastrotricha. Hydrobiologia 452: 155–161. Hochberg, R. & Litvaitis, M. (2001e): Macrodasyida (Gastrotricha): a cladistics analysis of morphology. Invertebr. Biol. 120: 124–135. Hochberg, R. & Litvaitis, M. K. (2003a): Organization of muscles in Chaetonotida Paucitubulatina (Gastrotricha). Meiofauna Mar. 12: 47–58. Hochberg, R. & Litvaitis, M. K. (2003b): Ultrastructural and immunocytochemical observations of the nervous system of three macrodasyidan gastrotrichs. Acta Zool. 84: 171–178. Hochberg, R., Atherton, S. and Gross, V. (2013): A new species of sublittoral marine gastrotrich, Lepidodasys ligni sp. n. (Macrodasyida, Lepidodasyidae), from the Atlantic coast of Florida. Zookeys 289: 1–12. Hochberg, R., Atherton, S. & Kieneke, A. (2014): Marine Gastrotricha of Little Cayman Island with the description of one new species

 129

and an initial assessment of meiofaunal diversity. Mar. Biodiv. 44: 89–113. Hummon, M. R. (1984a): Reproduction and sexual development in a freshwater gastrotrich. 1. Oogenesis of parthenogenetic eggs (Gastrotricha). Zoomorphology 104: 33–41. Hummon, M. R. (1984b): Reproduction and sexual development in a freshwater gastrotrich. 2. Kinetics and fine structure of postparthenogenetic sperm formation. Cell Tiss. Res. 236: 619–628. Hummon, M. R. (1984c): Reproduction and sexual development in a freshwater gastrotrich 3. Postparthenogenetic development of primary oocytes and the X-body. Cell Tiss. Res. 236: 629–636. Hummon, M. R. (1986): Reproduction and sexual development in a fresh-water gastrotrich. 4. Life History Traits and the possibility of sexual reproduction. Trans. Am. Microsc. Soc. 105: 97–109. Hummon, M. R. & Hummon, W. D. (1979): Reduction of fitness of the gastrotrich Lepidodermella squammata by dilute acid mine water and amelioration of the effect by carbonates. Int. J. Invertebr. Reprod. 1: 297–306. Hummon, M. R. & Hummon, W. D. (1983b): Gastrotricha. In: Adiyodi, K. G. & Adiyodi R. G. (eds.) Reproductive biology of invertebrates. Vol. 2: Spermatogenesis and sperm function, pp. 195–205. Wiley & Sons, Chichester. Hummon, M. R. & Hummon, W. D. (1992): Gastrotricha. In: Adiyodi, K. G. & Adiyodi R. G. (eds.) Reproductive biology of invertebrates. Vol. 5: Sexual differentiation and behaviour, pp. 137–146. Wiley & Sons, Chichester. Hummon, M. R. & Hummon, W. D. (1993): Gastrotricha. In: Adiyodi, K. G. & Adiyodi R. G. (eds.) Reproductive biology of invertebrates. Vol. 6: asexual propagation and reproductive strategies, pp. 265–277. Wiley & Sons, Chichester. Hummon, W. D. (1969): Musellifer sublitoralis, a new genus and species of Gastrotricha from the San Juan Archipelago, Washington. Trans. Am. Microsc. Soc. 88: 282–286. Hummon, W. D. (1972): Dispersion of Gastrotricha in a marine beach of the San Juan Archipelago, Washington. Mar. Biol. 16: 349–355. Hummon, W. D. (1974): Gastrotricha from Beaufort, North Carolina, U.S.A. Cah. Biol. Mar. 15: 431–446. Hummon, W. D. (1975): Habitat suitability and the ideal free distribution of Gastrotricha in a cyclic environment. In: Barnes, H. (ed.) Proc. 9th Eur. Mar. Biol. Symp., Aberdeen University Press, Aberdeen: 495–525. Hummon, W. D. (1976): Seasonal changes in secondary production, faunal similarity and biological accommodation, related to stability among Gastrotricha of two semi-enclosed Scottish beaches. In: Personne, G. & Jaspers, E. (eds.) Proc. 10th Eur. Mar. Biol. Symp., Universa Press, Wetteren: pp. 309–336. Hummon, W. D. (1982): Gastrotricha, In: S.P. Parker, ed. Synopsis and classification of living organisms, vol. 1. McGraw-Hill, New York. Hummon, W. D. (2008a): Gastrotricha of the North Atlantic Ocean: 1. Twenty four new and two redescribed species of Macrodasyida. Meiofauna Mar. 16: 117–174. Hummon, W. D. (2008b): Brackish-water Gastrotricha of the Polish Baltic coast. Meiofauna Mar. 16: 109–116. Hummon, W. D. (2010): Marine Gastrotricha of the Caribbean Sea: a review and new descriptions. Bull. Mar. Sci. 86: 661–708.

130 

 1 Gastrotricha

Hummon, W. D. (2011): Marine Gastrotricha of the Near East: I. Fourteen new species of Macrodasyida and a redescription of Dactylopodola agadasys Hochberg, 2003. ZooKeys 94: 1–59. Hummon, W. D. & Hummon, M. R. (1983a): Gastrotricha. In: Adiyodi, K. G. & Adiyodi R. G. (eds.) Reproductive biology of invertebrates. Vol. 1: Oogenesis, oviposition, and oosorption, pp. 211–221. Wiley & Sons, Chichester. Hummon, W. D. & Hummon, M. R. (1988): Gastrotricha. In: Adiyodi, K. G. & Adiyodi, R. G. (eds.) Reproductive biology of invertebrates. Vol. 3: Accessory sex glands, pp. 81–85. Wiley & Sons, Chichester. Hummon, W. D. & Hummon, M. R. (1989): Gastrotricha. In: Adiyodi, K. G. & Adiyodi R. G. (eds.) Reproductive biology of invertebrates. Vol. 4: fertilization, development, and parental care, pp. 201–206. Wiley & Sons, Chichester. Hummon, W. D. & Todaro, M. A. (2010): Analytic taxonomy and notes on marine, brackish-water and estuarine Gastrotricha. Zootaxa 2392: 1–32. Hummon, W. D., Todaro, A. & Tongiorgi, P. (1993): Italian marine Gastrotricha: II. One new genus and ten new species of Macrodasyida. Boll. Zool. 60: 109–127. Hummon, W. D., Todaro, A. & Evans, W. A. (2005): Video database for described species of marine Gastrotricha. Meiofauna Mar. 14: 23–26. Hyman, L. H. (1951): The invertebrates. Vol. 3: Acanthocephala, Aschelminthes, and Entoprocta: the pseudocoelomate Bilateria. McGraw-Hill, New York. Kånneby, T., Todaro, A. & Jondelius, U. (2009): On a new species and records of Ichthydium Ehrenberg, 1830 (Gastrotricha: Chaetonotida) from Sweden with a key to the genus. Zootaxa 2278: 26–46. Kånneby, T., Todaro, M. A. & Jondelius, U. (2012): A phylogenetic approach to species delimination in freshwater Gastrotricha from Sweden. Hydrobiologia 683: 185–202. Kånneby, T., Todaro, M. A. & Jondelius, U. (2013): Phylogeny of Chaetonotidae and other Paucitubulatina (Gastrotricha: Chaetonotida) and the colonization of aquatic ecosystems. Zool. Scr. 42: 88–105. Kånneby, T., Atherton, S. & Hochberg, R. (2014): Two new species of Musellifer (Gastrotricha: Chaetonotida) from Florida and Tobago and the systematic placement of the genus within Paucitubulatina. Mar. Biol. Res. 10: 983–995. Kieneke, A. (2010): A new species of Thaumastoderma (Gastrotricha: Macrodasyida) from the Antarctic deep sea with a phylogenetic analysis of the whole genus. J. Mar. Biol. Assoc. UK 90: 575–584. Kieneke, A. & Hochberg, R. (2012): Ultrastructural observations of the protonephridia of Polymerurus nodicaudus (Gastrotricha: Paucitubulatina). Acta Zool. 93: 115–124. Kieneke, A. & Ostmann, A. (2012): Structure, function and evolution of somatic musculature in Dasydytidae (Paucitubulatina, Gastrotricha). Zoomorphology 131: 95–114. Kieneke, A & Riemann, O. (2007): Eine interessante Teilgruppe der Süßwassergastrotrichen (Bauchhärlinge): Neogosseidae und Dasydytidae. Mikrokosmos 96: 139–146. Kieneke, A. & Zekely, J. (2007): Desmodasys abyssalis sp. nov. – first record of a deep sea gastrotrich from hydrothermal vents. Mar. Biodiv. Rec. 1: e88. Kieneke, A., Martínez Arbizu, P. & Ahlrichs, W. H. (2007): Ultrastructure of the protonephridial system in Neodasys

chaetonotoideus (Gastrotricha: Chaetonotida) and in the ground pattern of Gastrotricha. J. Morphol. 268: 602–613. Kieneke, A., Riemann, O. & Ahlrichs, W. H. (2008a): Novel implications for the basal internal relationships of Gastrotricha revealed by an analysis of morphological characters. Zool. Scr. 37: 429–460. Kieneke, A., Martínez Arbizu, P. & Riemann, O. (2008b): Body musculature of Stylochaeta scirtetica Brunson, 1950 and Dasydytes (Setodytes) tongiorgii (Balsamo, 1982) (Gastrotricha: Dasydytidae): a functional approach. Zool. Anz. 247: 147–158. Kieneke, A., Ahlrichs, W. H., Martínez Arbizu, P. & Bartolomaeus, T. (2008c): Ultrastructure of protonephridia in Xenotrichula carolinensis syltensis and Chaetonotus maximus (Gastrotricha: Chaetonotida): comparative evaluation of the gastrotrich excretory organs. Zoomorphology 127: 1–20. Kieneke, A., Martínez Arbizu, P. & Ahlrichs, W. H. (2008d): Anatomy and ultrastructure of the reproductive organs in Dactylopodola typhle (Gastrotricha: Macrodasyida) and their possible functions in sperm transfer. Invertebr. Biol.127: 12–32. Kieneke, A., Ahlrichs, W. H. & Martínez Arbizu, P. (2009): Morphology and function of reproductive organs in Neodasys chaetonotoideus (Gastrotricha: Neodasys) with a phylogenetic assessment of the reproductive system in Gastrotricha. Zool. Scr. 38: 289–311. Kieneke, A., Martínez Arbizu, P. M. & Fontaneto, D. (2012): Spatially structured populations with a low level of cryptic diversity in European marine Gastrotricha. Mol. Ecol. 21: 1239–1254. Kieneke, A., Rothe, B. H. & Schmidt-Rhaesa, A. (2013a): Record and description of Anandrodasys agadasys (Gastrotricha: Redudasyidae) from Lee Stocking Island (Bahamas), with remarks on populations from different geographic areas. Meiofauna Mar. 20: 39–48. Kieneke, A., Narkus, S., Hochberg, R. & Schmidt-Rhaesa, A. (2013b): Diplodasys rothei n. sp. (Gastrotricha, Macrodasyida), a new marine gastrotrich species from the Bahamas. Meiofauna Mar. 20: 49–61. Kisielewska, G. (1981): Hermaphroditism in freshwater gastrotrichs in natural conditions. Bull. Acad. Pol. Sci., Ser. Sci. Biol. 29: 167–172. Kisielewski, J. (1981): Gastrotricha from raised and transitional peat bogs in Poland. Monogr. Fauna Polski 11: 5–143. Kisielewski, J. (1987a): Two new interesting genera of Gastrotricha (Macrodasyida and Chaetonotida) from the Brazilian freshwater psammon. Hydrobiologia 153: 23–30. Kisielewski, J. (1987b): New records of marine Gastrotricha from the French coasts of Manche and Atlantic. I. Macrodasyida, with descriptions of seven new species. Bull. Mus. Natl. Hist. Nat. Paris, 4e Ser. 9: 837–877. Kisielewski, J. (1991): Inland water Gastrotricha from Brazil. Ann. Zool. Pol. Akad. Nauk Inst. Zool. 43, Suppl. 2: 1–168. Kisielewski, J. (1997): On the subgeneric division of the genus Chaetonotus Ehrenberg. Ann. Zool. 46: 145–151. Kolicka, M., Kisielewski, J., Nesteruk, T. & Zawierucha, K. (2013): Gastrotricha from the Poznań Palm House—one new subgenus and three new species of freshwater Chaetonotida (Gastrotricha). Zootaxa 3717: 231–279. Kraus, W. D. & Colacino, J. M. (1984): The oxygen consumption rates of three gastrotrichs. Comp. Biochem. Physiol. 79A: 691–693.

Literature 

Kraus, D. W., Travis, P. B., Colacino, J. M. & Ruppert, E. E. (1981): Occurrence of hemoglobin in Gastrotricha. Am. Zool. 21: 950. Kreutz, M. & Foissner, W. (2006): The Sphagnum ponds of Simmelried in Germany: a biodiversity hot-spot for microscopic organisms. Protozool. Monogr. 3: 1–267. Kristensen, R. M. & Hay-Schmidt A. (1989): The protonephridia of the Arctic kinorhynch Echinoderes aquilonius (Cyclorhagida, Echinoderidae). Acta Zool. 70: 13–27. Laverack, M. S. (1974): Structure and function of chemoreceptor cells. In: Grant, P. T. & Mackie, A. M. (eds.) Chemoreception in marine organisms, pp. 1–48. Academic Press, New York. Leasi, F. & Todaro, A. (2008): The muscular system of Musellifer delamarei (Renaud-Mornant, 1968) and other chaetonotidans with implications for the phylogeny ans systematization of the Paucitubulatina (Gastrotricha). Biol. J. Linn. Soc. 94: 379–398. Leasi, F. & Todaro, M. A. (2009): Meiofaunal cryptic species revealed by confocal microscopy: the case of Xenotrichula intermedia (Gastrotricha). Mar. Biol. 156: 1335–1346. Leasi, F. & Todaro, M. A. (2010): The gastrotrich community of a north Adriatic Sea site, with a redescription of Musellifer profundus (Chaetonotida: Muselliferidae). J. Mar. Biol. Assoc. UK 90: 645–653. Leasi, F., Rothe, B. H., Schmidt-Rhaesa, A. & Todaro, M. A. (2006): The musculature of three species of gastrotrichs surveyed with confocal laser scanning microscopy (CLSM). Acta Zool. 87: 171–180. Lee, J. M. & Chang, C. Y. (2002): Pseudostomella gastrotrichs (Macrodasyida, Thaumastodermatidae) from South Korea, with a brief review of the genus. Korean J. Biol. Sci. 6: 207–213. Lee, J. M. & Chang, C. Y. (2003): Two new marine gastrotrichs of the genus Ptychostomella (Macrodasyida, Thaumastodermatidae) from South Korea. Zool. Sci. 20: 481–489. Lee, J. M. & Chang, C. Y. (2011): Two new species of the genus Lepidodasys (Gastrotricha, Lepidodasyidae) from Tsushima Island, Japan. J. Nat. Hist. 45: 855–867. Lee, J. & Chang, C. Y. (2012): Crasiella clauseni, a new gastrotrich species (Macrodasyida, Planodasyidae) from Jeju Island, South Korea. Anim. Syst. Evol. Divers. 28: 42–47. Lee, J., Kim, D. & Chang, C. Y. (2013): Description of three new Tetranchyroderma gastrotrichs (Macrodasyida: Thaumastodermatidae) from South Korea. Zootaxa 3709: 483–493. Lee, J., Chang, C. Y. & Kim, D. (2014): Dendrodasys duplus, a new gastrotrich species (Macrodasyida: Dactylopodolidae) from South Korea. Anim. Syst. Evol. Divers. 30: 103–107. Levy, D. P. (1984): Obligate post-parthenogenic hermaphroditism and other evidence for sexuality in the life cycles of freshwater Gastrotricha. Ph.D. thesis, Rutgers University, New Brunswick, NJ. Levy, D. P. & Weiss, M. J. (1980): Sperm in the life cycle of the freshwater gastrotrich Lepidodermella squammata. Am. Zool. 20: 749. Liesenjohann, T., Neuhaus, B. & Schmidt-Rhaesa, A. (2006): Head sensory organs of Dactylopodola baltica (Macrodasyida, Gastrotricha): a combination of transmission electron micros­ copi­cal and immunocyto chemical techniques. J. Morphol. 267: 897–908. Ludwig, H. (1875): Über die Ordnung Gastrotricha Metschn. Z. Wiss. Zool. 26: 193–226. Malakhov, V. V. (1994): Nematodes – structure, development, classification, and phylogeny. Smithsonian Institution Press, Washington, DC.

 131

Manylov, O. G. (1995): Regeneration in Gastrotricha – I. Light microscopical observations on the regeneration in Turbanella sp. Acta Zool. 76: 1–6. Manylov, O. G., Vladychenskaya, N. S., Milyutina, I. A., Kedrova, O. S., Korokhov, N. P., Dvoryanchikov, G. A., Aleshin, V. V. & Petrov, N. B. (2004): Analysis of 18S rRNA gene sequences suggests significant molecular differences between Macrodasyida and Chaetonotida (Gastrotricha). Mol. Phyl. Evol. 30: 850–854. Marotta, R., Guidi, L., Pierboni, L., Ferraguti, M., Todaro M. A. & Balsamo, M. (2005): Sperm ultrastructure of Macrodasys caudatus (Gastrotricha: Macrodasyida) and a sperm-based phylogenetic analysis of Gastrotricha. Meiofauna Mar. 14: 9–21. Marotta, R., Todaro, A. & Ferraguti, M. (2008): The unique gravireceptor organs of Pleurodasys helgolandicus (Gastrotricha: Macrodasyida). Zoomorphology 127: 111–119. Mečnikow, E. (1865): Ueber einige wenig bekannte niedere Thierformen. Z. Wiss. Zool. 15: 450–458. Mock, H. (1979): Chaetonotoidea (Gastrotricha) der Nordseeinsel Sylt. Mikrofauna Meeresbodens 78: 1–107. Muschiol, D. & Traunspurger, W. (2009): Life at the extreme: meiofauna from three unexplored lakes in the caldera of the Cerro Azul volcano, Galápagos Islands, Ecuador. Aquat. Ecol. 43: 235–248. Nesteruk, T. (1996): Density and biomass of Gastrotricha in sediments of different types of standing waters. Hydrobiologia 324: 205–208. Neuhaus, B. (1987): Ultrastructure of the protonephridia in Dactylopodola baltica and Mesodasys laticaudatus (Macrodasyida): implications for the ground pattern of the Gastrotricha. Microfauna Mar. 3: 419–438. Neuhaus, B., Kristensen, R. M. & Lemburg, C. (1996): Ultrastructure of the cuticle of the Nemathelminthes and electron microscopical localization of chitin. Verh. Dtsch. Zool. Ges. 89.1: 221. Paps, J. & Riutort, M. (2012): Molecular phylogeny of the phylum Gastrotricha: new data brings together molecules and morphology. Mol. Phyl. Evol. 63: 208–212. Paps, J., Baguñà, J. & Riutort, M. (2009): Bilaterian phylogeny: a broad sampling of 13 nuclear genes provides a new Lophotrochozoa phylogeny and supports paraphyletic basal acoelomorpha. Mol. Biol. Evol. 26: 2397–2406. Petrov, N. B., Pegova, A. N., Manylov, O. G., Vladychenskaya, N. S., Mugue, N. S. & Aleshin, V. V. (2007): Molecular phylogeny of Gastrotricha on the basis of a comparison of the 18S rRNA genes: rejection of the hypothesis of a relationship between Gastrotricha and Nematoda. Mol. Biol. 41: 445–452. Pierboni, L. & Kristensen, R. M. (2007): Ultrastructural observations on the spermatozoa of a species of Lepidodasys (Gastrotricha, Macrodasyida). Helgoland Mar. Res. 61: 217–220. Pierboni, L., Marotta, R., Guidi, L., Todaro, M. A., Ferraguti, M. & Balsamo, M. (2003): I singolari spermatozoi di Urodasys (Gastrotricha, Macrodasyida): uno studio ultrastrutturale. 64 Congresso UZI, Varese, p. 182. Pierboni, L., Guidi, L., Marotta, R. & Todaro, M. A. (2004): Primi dati spermatologici sul genre Xenodasys (Gastrotricha, Macrodasyida, Dactylopodolidae) e loro

132 

 1 Gastrotricha

implicazioni filigenetiche. In: Atti 68 Congresso UZI. Giardini Naxos, Catania. Raikova, O. I., Flyatchinskaya, L. P. & Justine, J. L. (1998): Acoel spermatozoa: ultrastructure and immunocytochemistry of tubulin. Hydrobiologia 383: 207–214. Reisinger, E. (1978): Strukturanalyse der Bewegungsformen bei Gastrotrichen. Zool. Jb. Anat. 99: 23–24. Remane, A. (1924): Neue aberrante Gastrotrichen I. Macrodasys buddenbrocki nov. gen. nov. spec. Zool. Anz. 61: 289–297. Remane, A. (1925a): Neue aberrante Gastrotrichen II: Turbanella cornuta nov. spec. und T. hyalina M. Schultze 1853. Zool. Anz. 64: 309–314. Remane, A. (1925b): Organisation und systematische Stellung der aberranten Gastrotrichen. Verh. Dtsch. Zool. Ges. 1925: 121–128. Remane, A. (1926a): Morphologie und Verwandtschaftsbeziehungen der aberranten Gastrotrichen. Z. Morph. Ökol. Tiere 5: 625–754. Remane, A. (1926b): Marine Gastrotrichen aus der Ordnung Chaetonotoidea. Zool. Anz. 64: 243–252. Remane, A. (1927a): Neue Gastrotricha Macrodasyoidea. Zool. Jb. Syst. 54: 203–242. Remane, A. (1927b): Xenotrichula velox nov. gen. nov. spec., ein chaetonotides Gastrotrich mit männlichen Geschlechtsorganen. Zool. Anz. 71: 289–294. Remane, A. (1927c): Gastrotricha. In: Grimpe, G. (ed.) Die Tierwelt der Nord- und Ostsee, pp. 1–56. Akademische Verlagsgesellschaft, Leipzig. Remane, A. (1929): Gastrotricha. In Krumbach, T. (ed.) Handbuch der Zoologie, vol. 2, pp. 121–186. De Gruyter, Berlin. [volume occurred partitioned between 1929 and 1933]. Remane, A. (1934): Die Gastrotrichen des Küstengrundwassers von Schilksee. Schr. Naturwiss. Ver. Schleswig-Holstein 20: 473–478. Remane, A. (1936): Gastrotricha. In: Bronn, H. G. (ed.) Klassen und Ordnungen des Tierreiches. Gastrotricha und Kinoryncha. Vol. 4: Vermes II. Abteilung Askelminthes, Trochhelminthes, pp. 1–242. Akademische Verlagsgesellschaft, Leipzig. Remane, A. (1961): Neodasys uchidai nov. spec., eine zweite Neodasys-Art (Gastrotricha Chaetonotoidea). Kieler Meeresforsch. 17: 85–89. Ricci, C. & Balsamo, M. (2000): The biology and ecology of lotic rotifers and gastrotrichs. Freshwater Biol. 44: 15–28. Rieger, R. M. (1976): Monociliated epidermal cells in Gastrotricha: significance for concepts of early metazoan evolution. Z. Zool. Syst. Evolutionsforsch. 14: 198–226. Rieger, G. E. & Rieger, R. M. (1977): Comparative fine structure study of the gastrotrich cuticle and aspects of cuticle evolution within the Aschelminthes. Z. Zool. Syst. Evolutionsforsch. 15: 81–124. Rieger, G. E. & Rieger, R. M. (1980): Fine structure and formation of eggshells in marine Gastrotricha. Zoomorphology 96: 215–229. Rieger, R. M., Ruppert, E. E., Rieger, G. E. & Schoepfer-Sterrer, C. (1974): On the fine structure of gastrotrichs with description of Chordodasys antennatus sp. n. Zool. Scr. 3: 219–237. Roszczak, R. (1971): Metadasydytes quadrimaculatus g. n. sp. n. (Gastrotricha). Bull. Acad. Pol. Sci. B 19: 65–66. Rothe, B. H. & Schmidt-Rhaesa, A. (2008): Variation in the nervous system in three species of the genus Turbanella (Gastrotricha, Macrodasyida). Meiofauna Mar. 16: 175–184. Rothe, B. H. & Schmidt-Rhaesa, A. (2009): Architecture of the nervous system in two Dactylopodola species (Gastrotricha, Macrodasyida). Zoomorphology 128: 227–246.

Rothe, B. H. & Schmidt-Rhaesa, A. (2010): Oregodasys cirratus, a new species of Gastrotricha (Macrodasyida) from Tenerife (Canary Islands), with a description of the muscular and nervous system. Meiofauna Mar. 18: 49–66. Rothe, B. H., Schmidt-Rhaesa, A. & Kieneke, A. (2011a): The nervous system of Neodasys chaetonotoideus (Gastrotricha: Neodasys) revealed by combining confocal laserscanning and transmission electron microscopy – evolutionary comparison of neuroanatomy within the Gastrotricha and basal Protostomia. Zoomorphology 130: 51–84. Rothe, B. H., Kieneke, A. & Schmidt-Rhaesa, A. (2011b): The nervous system of Xenotrichula intermedia and X. velox (Gastrotricha: Paucitubulatina) by means of immunohistochemistry (IHC) and TEM. Meiofauna Mar. 19: 71–88. Ruppert, E. E. (1970). On Pseudostomella Swedmark 1956 with descriptions of P. plumosa nov. spec., P. cataphracta nov. spec. and a form of P. roscovita Swedmark 1956 from the west Atlantic coast. Cah. Biol. Mar. 11: 121–143. Ruppert, E. E. (1977): Zoogeography and speciation in marine Gastrotricha. Mikrofauna Meeresbodens 61: 231–251. Ruppert, E. E. (1978a): The reproductive system of gastrotrichs. II. Insemination in Macrodasys: a unique mode of sperm transfer in Metazoa. Zoomorphologie 89: 207–228. Ruppert, E. E. (1978b): The reproductive system of gastrotrichs. III. Genital organs of Thaumastodermatinae subfam. n. and Diplodasyinae subfam. n. with discussion of reproduction in Macrodasyida. Zool. Scr. 7: 93–114. Ruppert, E. E. (1979): Morphology and systematics of the Xenotrichulidae (Gastrotricha, Chaetonotida). Mikrofauna Meeresbodens 76: 1–56. Ruppert, E. E. (1982): Comparative ultrastructure of the gastrotrich pharynx and the evolution of myoepithelial foreguts in Aschelminthes. Zoomorphology 99: 181–220. Ruppert, E. E. (1988): Gastrotricha. In: Higgins, R. P. & Thiel, H. (eds.) Introduction to the study of meiofauna, pp. 302–311. Smithsonian Institution Press, Washington, DC. Ruppert, E. E. (1991): Gastrotricha. In Harrison, F. W. & Ruppert, E. E. (eds.) Microscopic anatomy of invertebrates, pp. 41–109. Wiley & Sons, New York. Ruppert, E. E. & Shaw, K. (1977): The reproductive system of gastrotrichs. I. Introduction with morphological data for two new Dolichodasys species. Zool. Scr. 6: 185–195. Ruppert, E. E. & Travis, P. B. (1983): Hemoglobin-containing cells of Neodasys (Gastrotricha, Chaetonotoida). I. Morphology and ultrastructure. J. Morphol. 175: 57–64. Ruttner-Kolisko, A. (1955): Rheomorpha neiswestnovae und Marinellina flagellata, zwei phylogenetisch interessante Wurmtypen aus dem Süßwasserpsammon. Österr. Zool. Z. 6: 55–69. Sacks, M. (1955): Observations on the embryology of an aquatic gastrotrich, Lepidodermella squammata (Dujardin, 1841). J. Morphol. 96: 473–495. Schmidt, P. (1974): Interstitielle Fauna von Galapagos IV. Gastrotricha. Mikrofauna Meeresbodens 26: 1–76. Schmidt, P. & Teuchert, G. (1969): Quantitative Untersuchungen zur Ökologie der Gastrotrichen im Gezeiten-Sandstrand der Insel Sylt. Mar. Biol. 4: 4–23. Schmidt-Rhaesa, A. (2002): Two dimensions of biodiversity research exemplified by Nematomorpha and Gastrotricha. Integr. Comp. Biol. 42: 633–640.

Literature 

Schmidt-Rhaesa, A. (2007): The evolution of organ systems. Oxford University Press, Oxford. Schmidt-Rhaesa, A. (2013): Gastrotricha, Cycloneuralia and Gnathifera: general history and phylogeny. In: Schmidt-Rhaesa, A. (ed.) Handbook of Zoology. Gastrotricha, Cycloneuralia and Gnathifera. Volume 1: nematomorpha, Priapulida, Kinorhyncha and Loricifera, pp. 1–10. De Gruyter, Berlin. Schmidt-Rhaesa, A., Bartolomaeus, T., Lemburg, C., Ehlers, U. & Garey, J. R. (1998): The position of the Arthropoda in the phylogenetic system. J. Morphol. 238: 263–285. Schöpfer-Sterrer, C. (1969): Chordodasys riedli gen. nov., spec. nov. a macrodasyid gastrotrich with a chordoid organ. Cah. Biol. Mar. 10: 391–404. Schoepfer-Sterrer, C. (1974): Five new species of Urodasys and remarks on the terminology of the genital organs in Macrodasyidae (Gastrotricha). Cah. Biol. Mar. 15: 229–254. Schrom, H. (1966): Verteilung einiger Gastrotrichen im oberen Eulitoral eines nordadriatischen Sandstrandes. Veröff. Inst. Meeresforsch. Bremerhaven, Spec. Vol. 2: 95–103. Schrom, H. (1972): Nordadriatische Gastrotrichen. Helgoländer Wiss. Meeresunt. 23: 286–351. Schultze, M. (1853): Über Chaetonotus und Ichthydium (Ehrb.) und eine neue verwandte Gattung Turbanella. Müller’s Arch. Anat. Physiol. 6: 241–254. Schwank, P. (1990): Gastrotricha. In: Schwoerbel, J. & Zwick, P. (eds.) Süsswasserfauna von Mitteleuropa. Vol. 3 Gastrotricha und Nemertini, pp. 1–252. Gustav Fischer Verlag, Stuttgart. Strand, E. (1929): Zoological and palaeontological nomenclatorical notes. Arb. Syst. Zool. Inst. Lettl. Univ. Riga. 29: 1–29. Strayer, D. L. & Hummon, W. D. (1991): Gastrotricha. In: Thorp, J. H. & Covich, A. P. (eds.) Ecology and classification of North American freshwater invertebrates, pp. 173–185. Academic Press, New York. Suzuki, T., Maeda, M. & Furuya, H. (2013): Two new Japanese species of Gastrotricha (Chaetonotida, Chaetonotidae, Lepidodermella and Dichaeturidae, Dichaetura), with comments on the diversity of gastrotrichs in rice paddies. Zootaxa 3691: 229–239. Swedmark, B. (1955): Developement d’un gastrotriche macrodasyoide, Macrodasys affinis Remane. Comp. Rend. Acad. Sci. 240: 1812–1814. Swedmark, B. (1956) Nouveaux gastrotriches Macrodasyides de la region de Roscoff. Arch. Zool. Exp. Gen. 94: 43–57. Swedmark, B. (1967): Trois nouveaux gastrotriches macrodasyoides de la faune interstitielle marine des sables de Roscoff. Cah. Biol. Mar. 8: 323–330. Teuchert, G. (1967): Zum Protonephridialsystem mariner Gastrotrichen der Ordnung Macrodasyoidea. Mar. Biol. 1: 110–112. Teuchert, G. (1968): Zur Fortpflanzung und Entwicklung der Macrodasyoidea (Gastrotricha). Z. Morph. Tiere 63: 343–418. Teuchert, G. (1973): Die Feinstruktur des Protonephridialsystems von Turbanella cornuta Remane, einem marinen Gastrotrich der Ordnung Macrodasyoidea. Z. Zellforsch. 136: 277–289. Teuchert, G. (1974): Aufbau und Feinstruktur der Muskelsysteme von Turbanella cornuta Remane (Gastrotricha, Macrodasyoidea). Mikrofauna Meeresbodens 39: 223–246.

 133

Teuchert, G. (1975a): Organisation und Fortpflanzung von Turbanella cornuta (Gastrotricha). Film C 1176. Institut für den wissenschaftlichen Film, Göttingen. Teuchert, G. (1975b): Differenzierung von Spermien bei dem marinen Gastrotrich Turbanella cornuta (Ordnung Macrodasyoidea). Verh. Anat. Ges. 69: 743–748. Teuchert, G. (1976a): Sinneseinrichtungen bei Turbanella cornuta Remane (Gastrotricha). Zoomorphologie 83, 193–207. Teuchert, G. (1976b): Elektronenmikroskopische Untersuchung über die Spermatogenese und Spermatohistogenese von Turbanella cornuta Remane (Gastrotricha). J. Ultrastruct. Res. 56: 1–14. Teuchert, G. (1977a): The ultrastructure of the marine gastrotrich Turbanella cornuta Remane (Macrodasyoidea) and its functional and phylogenetical importance. Zoomorphologie 88: 189–246. Teuchert, G. (1977b): Leibeshöhlenverhältnisse von dem marinen Gastrotrich Turbanella cornuta Remane (Ordnung Macrodasyoidea) und eine phylogenetische Bewertung. Zool. Jb. Anat. 97: 586–596. Teuchert, G. (1978): Strukturanalyse von Bewegungsformen bei Gastrotrichen. Zool. Jb. Anat. 99: 12–22. Teuchert, G. & Lappe, A. (1980): Zum sogenannten “Pseudocoel” der Nemathelminthes – Ein Vergleich der Leibeshöhlen von mehreren gastrotrichen. Zool. Jb. Anat. 103: 424–438. Todaro, M. A. (2002): An interesting new gastrotrich from littoral meiobenthos (Long Beach Island, USA), with a key to species of Tetranchyroderma (Gastrotricha, Macrodasyida). J. Mar. Biol. Assoc. UK 82: 555–564. Todaro, M. A. (2012): A new marine gastrotrich from the State of São Paulo (Brazil), with a key to species of Pseudostomella (Gastrotricha, Thaumastodermatidae). ZooKeys 223: 39–51. Todaro, M. A. (2013): A new non-naked species of Ptychostomella (Gastrotricha) from Brazil. ZooKeys 289: 13–24. Todaro, A. & Hummon, W. D. (2008) An overview and a dichotomous key to genera of the phylum Gastrotricha. Meiofauna Mar. 16: 3–20. Todaro, M. A. & Leasi, F. (2013): A new eye-bearing Macrodasys (Gastrotricha: Macrodasyida) from Jamaica. Meiofauna Mar. 20: 33–38. Todaro, M. A., Fleeger, J. W., Hu, Y. P., Hrincevich, A. W. & Foltz, D. W. (1996): Are meiofauna species cosmopolitan? Morphological and molecular analysis of Xenotrichula intermedia (Gastrotricha: Chaetonotida). Mar. Biol. 125: 735–742. Todaro, M. A., Balsamo, M., Guidi, L. & Bernhard, J. M. (2000a): Peculiarità ultrastrutturali di un gastrotrico microaerofilo. In: Atti 61° Congresso UZI. S. Benedetto del Tronto, p. 130. Todaro, M. A., Hummon, W. D., Balsamo, M. Fregni, E. & Tongiorgi, P. (2000b): Inventario dei Gastrotrichi Marini Italiani: Una Checklist Annotata. Atti Soc. Tosc. Sci. Nat. Mem. Ser. B 107: 75–137. Todaro, M. A., Littlewood, D. T. J., Balsamo, M., Herniou, E. A., Casanelli, S., Manicardi, G., Wirz, A. & Tongiorgi, P. (2003): The interrelationships of the Gastrotricha using nuclear small rRNA subunit sequence data, with an interpretation based on morphology. Zool. Anz. 242: 145–156. Todaro, M. A., Balsamo, M. & Kristensen, R. M. (2005): A new genus of marine chaetonotids (Gastrotricha), with a description of two new species from Greenland and Denmark. J. Mar. Biol. Assoc. UK 85: 1391–1400.

134 

 1 Gastrotricha

Todaro, M. A., Telford, M. J., Lockyer, A. E. & Littlewood, D. T. J. (2006a): Interrelationships of Gastrotricha and their place among the Metazoa inferred from 18S rRNA genes. Zool. Scr. 35: 251–259. Todaro, M. A., Guidi, L., Leasi, F. & Tongiorgi, P. (2006b): Morphology of Xenodasys (Gastrotricha): the first species from the Mediterranean Sea and the establishment of Chordodasiopsis gen. nov. and Xenodasyidae fam. nov. J. Mar. Biol. Assoc. UK 86: 1005–1015. Todaro, M. A., Dal Zotto, M., Maiorova, A. S. & Adrianov, A. V. (2009): A new species of Aspidiophorus (Gastrotricha, Chaetonotida) from the Russian Far East with a key to marine species of the genus. Mar. Biol. Res. 5: 297–303. Todaro, M. A., Kånneby, T., Dal Zotto, M. & Jondelius, U. (2011a): Phylogeny of Thaumastodermatidae (Gastrotricha: Macrodasyida) inferred from nuclear and mitochondrial sequence data. PLoS One 6: e17892. Todaro, M. A., Dal Zotto, M., Perissinotto, R. & Bownes, S. J. (2011b): First records of Gastrotricha from South Africa, with description of a new species of Halichaetonotus (Chaetonotida, Chaetonotidae). Zookeys 142: 1–13. Todaro, M. A., Guidi, L., Ferraguti, M., Balsamo, M. (2012a): A fresh look at Dinodasys mirabilis (Gastrotricha, Macrodasyida), with focus on the reproductive apparatus and sperm ultrastructure. Zoomorphology 131: 115–125. Todaro, M. A., Dal Zotto, M., Jondelius, U., Hochberg, R., Hummon, W. D., Kånneby, T. & Rocha, C. E. F. (2012b): Gastrotricha: a marine sister for a freshwater puzzle. PLoS One 7(2): e31740. Todaro, M. A., Perissinotto, R. & Bownes, S. J. (2013): Neogosseidae (Gastrotricha, Chaetonotida) from the iSimangaliso Wetland Park, KwaZulu-Natal, South Africa. ZooKeys 315: 77–94. Travis, P. B. (1983): Ultrastructural study of body wall organization and Y-cell composition in the Gastrotricha. Z. Zool. Syst. Evolutionsforsch. 21: 52–68. Tyler, S. (1988): The role of function in determination of homology and convergence – examples from invertebrate adhesive organs. Fortschr. Zool. Progr. Zool. 36: 331–347.

Tyler, S., Melanson, L. A. & Rieger, R. M. (1980): Adhesive organs of the Gastrotricha II. The organs of Neodasys. Zoomorphologie 95: 17–26. Tyler, S. & Rieger, G. E. (1980): Adhesive organs of the Gastrotricha. I. Duo-gland organs. Zoomorphologie 95: 1–15. Valbonesi, A. & Luporini, P. (1984): Researches on the coast of Somalia. Gastrotricha Macrodasyoidea. Ital. J. Zool. Suppl. 19: 1–34. Vanamala Naidu, K. & Chandrasekahara Rao, G. (2004): The fauna of India and the adjacent countries: Gastrotricha. Zool. Surv. India 1–169. Von und zu Gilsa, A., Kieneke, A., Hochberg, R. & Schmidt-Rhaesa, A. (2014): Two new species of the genus Dactylopodola (Gastrotricha, Marcodasyida) from the Bahamas, with an updated key to the genus. Cah. Biol. Mar. 55: 333–345. Wägele, J. W. (2005): Foundations of phylogenetic systematics. Verlag Dr. Friedrich Pfeil, München. Wagner, F. (1893): Der Organismus der Gastrotrichen. Biol. Zentralbl. 13: 223–238. Weiss, M. J. (2001): Widespread hermaphroditism in freshwater gastrotrichs. Invertebr. Biol.120: 308–341. Weiss, M. J. & Levy, D. P. (1979): Sperm in “parthenogenetic” freshwater gastrotrichs. Science 205: 302–303. Wiedermann, A. (1995): Zur Ultrastruktur des Nervensystems bei Cephalodasys maximus (Macrodasyida, Gastrotricha). Microfauna Mar. 10: 173–233. Wieser, W. (1957): Gastrotricha Macrodasyoidea from the intertidal of Puget Sound. Trans. Am. Microsc. Soc. 76: 372–381. Wilke, U. (1954): Mediterrane Gastrotrichen. Zool. Jb. Syst. Ökol. Geogr. Tiere 82: 497–550. Wirz, A., Pucciarelli, S., Miceli, C., Tongiorgi, P. & Balsamo, M. (1999): Novelty in phylogeny of Gastrotricha: evidence from 18s rRNA gene. Mol. Phyl. Evol. 13: 314–318. Zelinka, C. (1889): Die Gastrotrichen. Z. Wiss. Zool. 49: 209–384. Zrzavý, J. (2003): Gastrotricha and metazoan phylogeny. Zool. Scr. 32: 61–81. Zrzavy, J., Mihulka, S., Kepka, P., Bezdek, A. & Tietz, D. (1998): Phylogeny of the metazoa based on morphological and 18S ribosomal DNA evidence. Cladistics 14: 249–285.

Wolfgang Sterrer and Martin V. Sørensen

2 Phylum Gnathostomulida 2.1 Introduction and history of research (Fig. 2.1)1 Gnathostomulida (“lesser jaw worms”; Kiefermäulchen) is a small phylum of free-living, microscopic, unsegmented, entirely monociliated marine worms. The first gnathostomulid – a millimeter-long worm with snapping jaws – was discovered in 1928, in the fine sand of Kiel Bight (Germany), by Adolf Remane (Kiel), pioneer of interstitial sand fauna research (Ax 1956). Remane eventually passed his scarce material on to Josef Meixner (Graz) who, naming it Remanella n. g. paradoxa n. sp., provided a drawing (Fig. 2.1) and brief diagnosis within a new family, Remanellidae. This he appended, with a question mark, to Turbellaria-Macrostomida in the manuscript for “Turbellaria (Strudelwürmer) II. System”, which was to be published in Tierwelt der Nord- und Ostsee in 1938. The outbreak of World War II prevented the publication of this chapter, and only page proofs were privately circulated (Meixner 1938). In 1946, Josef Meixner died in a prisoner-of-war camp. From 1951 on, Peter Ax, a student of Remane to whom much of Meixner’s material was entrusted, began finding the elusive creatures again, first at Kiel Bight, then on the island of Sylt (North Sea). The discovery, in September 1954, of what turned out to be another genus of the same worm group, this time in a brackish coastal pond near Banyulssur-Mer (Mediterranean), prompted Ax to publish “The Gnathostomulida – an enigmatic worm group from marine sand” (Ax 1956), name Remane’s original find Gnathostomula paradoxa (Remanella having become unavailable), and describe the Mediterranean species as Gnathostomaria lutheri. The taxon Gnathostomulida was presented as a new order of Turbellaria-Archoophora. Three more species of Gnathostomula were described over the next few years, one each from the Maldives (Gerlach 1958), the Barents Sea (Mamkaev 1961), and the Caribbean (Kirsteuer 1964). In 1964, as a beginning graduate student looking for Turbellaria in subtidal, detritus-rich sand off a small beach near Kristineberg (Swedish west coast), the first author happened upon G. paradoxa, and then in short order upon a dozen new species of Gnathostomulida, in three new genera (Sterrer 1966a, b, 1969). Subsequent finds in the Mediterranean (Sterrer 1965) made it clear that gnathostomulids had

seemed so elusive only because marine interstitial fauna research had traditionally focused on clean sand – whereas Gnathostomulida, as a rule, inhabit sand with a substantial admixture of organic detritus (Sterrer 1971a, b). Since then, there has been a steady trickle of new species discoveries, from the NW Atlantic (Kirsteuer 1969a, b, 1970, Riedl 1970a, b, 1971a, b, Farris 1973, 1977, Sterrer 1970a, c, 1973a, 1976, 1992, 1998, 2004, 2011, Sterrer & Farris 1975), the NE Atlantic (Kristensen & Nørrevang 1977, 1978, Sterrer 1997), the Red Sea (Riedl 1966), Mediterranean (Sterrer 1970b, 1991d), the central and south Pacific (Sterrer 1991a, b, c, 2001a, 2006), and NE Pacific (Ehlers & Ehlers 1973, Sterrer & Sørensen 2006). Another two dozen species, mostly from the Pacific and Mediterranean, are awaiting description (Sterrer, unpublished data). To date, 100 species have been described in 26 genera, 12 families, 2 suborders, and 2 orders (Tab. 2.1). Although it was once predicted “that the total number of gnathostomulid species will exceed 1,000” (Sterrer 1972), this estimate, barring a large number of as yet unrecognized cryptic species, should probably now be revised down to “a few hundred”. Even though much of the world’s oceans (especially deeper sandy bottoms, and polar regions) have yet to be prospected for Gnathostomulida, the unexpectedly large number of cosmopolitan morpho-species makes the discovery of many more species less likely. Originally ranked among Turbellaria, the taxon Gnathostomulida was elevated by Riedl (1969) to the rank of phylum, for which Sterrer (1972) proposed the currently accepted system (Sørensen et  al. 2006). Some authors continued to argue for strong affinities between gnathostomulids and platyhelminths (Ax 1985, 1989, 1995, Meglitsch & Schram 1991), and a sister-group relationship was supported by several phylogenetic analyses based on morphological data (Eernisse et  al. 1992, Schram & Ellis 1994, Zrzavý et al. 1998, Peterson & Eernisse 2001). However, as pointed out by Jenner (2004), most of these analyses were either based on a restricted taxon sampling that did not allow test of alternative hypotheses, or they neglected characters that would support alternative positions of the gnathostomulids. Rieger (1976) signaled similarities in the ultrastructure of the monociliated epidermal cells in gastrotrichs and gnathostomulids, and after further studies of the gnathostomulid epidermis,

1  We dedicate this chapter to Volker Lammert (1955–1998), who in his brief life contributed more to gnathostomulid knowledge than he had time to publish.

136 

 2 Phylum Gnathostomulida

Fig. 2.1: The original line art sketch, showing the first gnathostomulid ever recorded. The animal was found by Adolf Remane who handed it over to Josef Meixner. Meixner produced this drawing and assigned the name Remanella paradoxa, but this initial work of Meixner was never published. Abbreviations: bm, bursa mouthpiece; bs, bursa copulatrix; co, copulatory organ; da, intestine; o, ovary; ph, pharynx; te, testis. (Original courtesy of Tor G. Karling.)

Rieger & Mainitz (1977) argued that the phylum should be situated in a “position between Platyhelminthes and Aschelminthes”. The conflicting views that lie behind this somewhat imprecise statement were also reflected in the review of Sterrer et  al. (1985), who concluded that gnathostomulids possessed traits that pointed toward platyhelminths, nemertodermatids, some groups of microscopic annelids, gastrotrichs, and rotifers. A potential homology between the pharyngeal hard parts in gnathostomulids and rotifers was first suggested by Reisinger (1961), and this hypothesis was sustained when transmission electron microscopic (TEM) examinations revealed an identical ultrastructural composition of the hard parts in the two groups (Ahlrichs 1995a, b, Rieger & Tyler 1995). The presence of pharyngeal hard parts with a unique ultrastructure was suggested as a synapomorphy for Gnathostomulida and Rotifera (including the acanthocephalans), and the two groups were united in a clade named Gnathifera (Ahlrichs 1995a). This putative relationship obtained further support with the discovery of the third gnathiferan phylum, Micrognathozoa (Kristensen & Funch 2000), and gnathiferan monophyly has subsequently been confirmed in several studies and cladistic analyses based on morphological data (Haszprunar 1996, Sørensen et al. 2000, Nielsen 2001, Sørensen 2003, Zrzavý 2003, Halanych 2004). Whereas the morphological approach appears to have reached an increasing consensus about the phylogenetic position of gnathostomulids, analyses of molecular sequence data still fail to present unambiguous results. Some analyses of 18S rRNA suggest a sister-group relationship between gnathostomulids and chaetognaths (Littlewood et  al. 1998, Glenner et  al. 2004), but this result may very well be a long-branch artifact (Jenner 2004). As a preliminary result, Peterson & Eernisse (2001) found support for an acoel-gnathostomulid relationship, but also due to long-branch problems, the gnathostomulid terminal was excluded from the final analyses. A gastrotrich-gnathostomulid relationship was supported by analysis of 18S rRNA by Zrzavý et al. (1998) and of combined analysis of the loci 18S rRNA, 28S rRNA, and histone 3 (Giribet et al. 2004). However, with the histone 3 sequence omitted from the latter study, support was found for a monophyletic group consisting of the gnathiferan taxa and Cycliophora (Giribet et al. 2004). An identical clade has been recovered from analysis of 18S rRNA sequences alone and 18S rRNA sequences combined with a morphological data set (Giribet et al. 2000). The most recent phylogenomic analysis of expressed sequence tags from a

Filospermoidea

Order

Suborder

Author

Genus

Sterrer, 1998 Sterrer, 1966 Sterrer, 1969

Sterrer, 1969 Sterrer, 1969

Sterrer, 1966

belizensis filum gubbarnorum

lunulifera rosea

ruberrima

Sterrer, 1966

simplex

Sterrer, 1991a Sterrer, 1991a

arcus bastillae

Sterrer, 1998

Sterrer, 1991b

rufa

rubromaculata Sterrer, 1969

Sterrer, 1991b

Pterognathia filum Sterrer, 1966 Pterognathia gubbarnorum Sterrer, 1966; Haplognathia lyra Sterrer, 1970

Fiji (S Pacific)

Kristineberg (NE Atlantic) Hawaii (NE Pacific) Kristineberg Pterognathia (NE Atlantic) simplex Sterrer, 1966 Florida (NW Atlantic) Fiji (S Pacific)

Kristineberg Pterognathia (NE Atlantic) grandis Kirsteuer, 1969

Kristineberg (NE Atlantic) Kristineberg Haplognathia (NE Atlantic) rosacea Sterrer, 1970

Belize (NW Atlantic) Kristineberg (NE Atlantic) Kristineberg (NE Atlantic)

Hawaii (NE Pacific)

Species author Type locality Synonyms and reference

asymmetrica

Species

Pterognathiidae Sterrer, 1972 Cosmognathia aquila

Haplognathiidae Sterrer, 1972 Haplognathia

Family

Tab. 2.1: Classification, species, and global distribution of Gnathostomulida.

Sterrer 1998

Sterrer (unpublished data) Müller & Ax 1971

Müller & Ax 1971

Sterrer 1998 and unpublished data Müller & Ax 1971, Sterrer 1998, 2001, 2006, and unpublished data Sterrer 1998, 2001, 2006, and unpublished data

Müller & Ax 1971, Sterrer 1998, 2001, 2006, and unpublished data

Müller & Ax 1971

Sterrer 1998, 2001, 2006, and unpublished data

References

NW Atlantic, NE Sterrer 1998, Pacific, NE Australia Sterrer 2001 NE Australia Sterrer 2001 (Continued)

NW Atlantic

NE Atlantic

Red Sea

NW Atlantic, NE Pacific NW Atlantic, S Pacific, NE Australia, New Zealand, NE Pacific NW Atlantic, Mediterranean, S Pacific, NE Australia, New Zealand, NE Pacific

NW Atlantic, Mediterranean, NE Australia, New Zealand, NE Pacific

NE Atlantic

NW Atlantic, SW Pacific, NE Australia, New Zealand, NE Pacific

Distribution

 2.1 Introduction and history of research 

 137

Suborder

Family

Author

Pterognathia

Genus

Bursovaginoidea Conophoralia Austrognathiidae Sterrer, 1971 Triplignathia

Order

Tab. 2.1: ( Continued )

Sterrer, 1991b Sterrer, 1969 Sterrer, 2006 Sterrer, 1998

Sterrer, 2001 Sterrer, 1969 Sterrer, 1966 Sterrer, 2006 Sterrer, 1991c

Sterrer, 1991a

hawaiiensis meixneri portobello pygmaea

sica sorex swedmarki tuatara ugera

vilii

Sterrer, 1998

Sterrer, 1970

ctenifera

bathycola

Sterrer, 1991a

crocodilus

Sterrer, 1991d

Sterrer, 1969

atrox

adriatica

Sterrer, 1998

alcicornis

Sterrer, 1991b

manubrium

Fiji (S Pacific) Dubrovnik (Mediterranean) North Carolina (NW Atlantic)

North Carolina (NW Atlantic) Hawaii (NE Pacific) Kristineberg (NE Atlantic) New Zealand (S Pacific) North Carolina (NW Atlantic) Lizard Island (NE Australia) Kristineberg (NE Atlantic) Kristineberg (NE Atlantic) New Zealand (S Pacific) Tahiti (S Pacific)

Hawaii (NE Pacific) Belize (NW Atlantic) Kristineberg (NE Atlantic) Fiji (S Pacific)

Species author Type locality Synonyms and reference

Species

NW Atlantic, New Zealand, Red Sea, NE Pacific

NE Atlantic, NW Atlantic NW Atlantic

New Zealand

NW Atlantic, NE Atlantic, Red Sea S Pacific, NE Australia

NW Atlantic, NE Australia, Red Sea

Distribution

Sterrer 1998, 2006, and unpublished data

Müller & Ax 1971, Sterrer 1998 Sterrer 1998, Müller & Ax 1971

Sterrer 2006

Sterrer 1998 and unpublished data Sterrer 1998, Sterrer 2001

Sterrer 1998 and unpublished data

References

138   2 Phylum Gnathostomulida

Order

Suborder

Family

Author

Species Sterrer, 2001

Species author Type locality Synonyms and reference

Riedl, 1966

riedli f. marisrubri singatokae

Sterrer, 2011 Sterrer, 1971 Sterrer, 1991a Sterrer, 1970

Sterrer, 1998 Sterrer, 1991c

boadeni homunculus kirsteueri

medusifera mooreensis

Ehlers & Ehlers, 1973 Sterrer, 2006

barbadensis

Austrognatha- atraclava ria australis

Rovinj (Mediterranean) Al-Ghardaqa (Red Sea) Fiji (S Pacific) Galapagos (E Pacific) New Zealand (S Pacific) Barbados (NW Atlantic) N Ireland (NE Atlantic) Fiji (S Pacific) North Carolina (NW Atlantic) Panama (NW Atlantic) Tahiti (S Pacific)

Sterrer, 1965

riedli

Sterrer, 1991a

Fiji (S Pacific)

novaezelandiae Sterrer, 1991a

Lizard Island (NE Australia) christianae Farris, 1977 North Carolina (NW Atlantic) clavigera Sterrer, 1997 Canaries (NE Atlantic) hymanae Kirsteuer, 1970 Barbados (NW Atlantic) macroconifera Sterrer, 1991c Tahiti (S Pacific) microconulifera Farris, 1977 Bermuda (NW Atlantic) nannulifera Sterrer, 1991a Fiji (S Pacific)

Austrognathia australiensis

Genus

NW Atlantic

(Continued)

Sterrer 1998

Sterrer 1998, 2001

Sterrer 1998 and unpublished data Sterrer 2001

NW Atlantic, Red Sea Papua New Guinea

Papua New Guinea

Sterrer 1998, 2001, and unpublished data

References

NW Atlantic, NE Australia, E Pacific

Distribution

 2.1 Introduction and history of research 

 139

Order

nominata

beckeri

strunki

stirialis

sterreri

Sterrer, 1991a

pecten

Rastrognathiidae Kristensen & Nørrevang, 1977

Sterrer, 1991a

vitiensis

Bermuda (NW Atlantic) Fiji (S Pacific) Kristensen & Helsingør Nørrevang, 1977 (E Atlantic)

Sterrer, 1976

rikerae

Rastrognathia macrostoma

Tenuignathia

Labidognathia longicollis

remanei

eastwardiae

Sterrer, 1972 Mesognatharia bahamensis

Sterrer, 1992 Clausognathia suicauda

Clausognathiidae Mesognathariidae

lutheri

Fiji (S Pacific) (Kirsteuer, Bimini 1969) (NW Atlantic) Sterrer, 1998 Florida (NW Atlantic) Farris, 1973 Florida (NW Atlantic) Sterrer, 1971 Florida (NW Atlantic) Sterrer, 2001 Lizard Island Agnathiella sp. (NE Australia) Sterrer, 1998 Sterrer, 1997 Canaries (NE Atlantic) Ax, 1956 Southern France (Mediterranean) Sterrer, 1992 Belize (NW Atlantic) Kirsteuer, 1969 Bimini (NW Atlantic) Sterrer, 1998 North Carolina (NW Atlantic) Sterrer, 1966 Kristineberg (NE Atlantic) Riedl, 1970a North Carolina (NW Atlantic)

Species author Type locality Synonyms and reference

Species

Paragnathiella trifoliceps

Sterrer, 1972 Agnathiella

Genus

Sterrer, 1972 Gnathostomaria

Agnathiellidae

Scleroperalia

Author

Gnathostomariidae

Family

Suborder

Tab. 2.1: ( Continued)

NW Atlantic, NE Atlantic, NE Australia, SW and NE Pacific NW Atlantic, NE Australia, Red Sea

NW Atlantic, Red Sea

NW Atlantic

NW Atlantic, South Africa

NW Atlantic, Fiji (S Pacific) Red Sea

SW and NE Pacific

NW Atlantic

NW Atlantic

NW Atlantic

Distribution

Sterrer 1998, 2001, and unpublished data

Sterrer 1998, 2001, and unpublished data

Sterrer 1998 and unpublished data

Sterrer 1998

Sterrer (unpublished data) Sterrer 1998 and unpublished data

Sterrer 1998 and unpublished data Sterrer 1998

Sterrer 1998

Sterrer 1998

Sterrer 1998

References

140   2 Phylum Gnathostomulida

Order

Suborder

Author

Genus anonyma

Species Sterrer, 1998

Species author Type locality Synonyms and reference

dracula

armata axi

arabica

Gnathostomula algreti

Chirognathia

Corculognathia apennata

Galapagos (E Pacific) Bamfield (NE Pacific) Tahiti (S Pacific) Riedl, 1971b Al-Ghardaqa (Red Sea) Riedl, 1971b Massachusetts (NW Atlantic) Kirsteuer, 1964 Venezuela (NW Atlantic)

Ehlers & Ehlers, 1973 Sterrer & Sørensen, 2006 Sterrer, 1991c

Belize (NW Atlantic) Onychognathi- Sterrer, 1972 Onychognathia bractearotunda Ehlers & Ehlers, Galapagos idae 1973 (E Pacific) filifera Riedl, 1971 Florida (NW Atlantic) rhombocephala Sterrer, 1998 Belize (NW Atlantic) Vampyrognathia horribilis Sterrer, 1998 Florida (NW Atlantic) minor Sterrer, 1998 North Carolina (NW Atlantic) varanus Sterrer, 2001 Lizard Island (NE Australia) Valvognathia pogonostoma Kristensen & Helsingør Nørrevang, 1978 (E Atlantic) Nanognathia exigua Sterrer, 1973 North Carolina (NW Atlantic) Goannagnathia susannae Sterrer, 2001 Lizard Island (NE Australia) Problognathiidae Sterrer & Problognathia minima Sterrer & Farris, Bermuda Farris, 1975 1975 (NW Atlantic) Gnathostomulidae Sterrer, 1972 Semaeognasterreri Riedl, 1970b North thia Carolina (NW Atlantic) Ratugnathia makuluvae Sterrer, 1991a Fiji (S Pacific)

Paucidentulidae Sterrer, 1998 Paucidentula

Family

Sterrer 1998

NW Atlantic

NW Atlantic

NE Pacific

NW Atlantic, South Africa

Sterrer 1998

NW Atlantic

2.1 Introduction and history of research 

(Continued)

Sterrer 1998

Sterrer 1998 and unpublished data

Sterrer 1998

Sterrer (unpublished data)

Red Sea, NE Pacific

Sterrer 1998

Sterrer 1998

Sterrer 1998

References

NW Atlantic

Distribution



 141

Suborder

2 suborders

Order

2 orders

Tab. 2.1: ( Continued)

12 families

Family

Author

26 genera

Genus

100 species

raji salotae uncinata

peregrina

paradoxa

nigrostoma

murmanica

microstyla

mediterranea

maorica mediocristata

maldivarum

karlingi

jenneri

costata

Riedl, 1971b

brunidens North Carolina (NW Atlantic) Ehlers & Ehlers, Galapagos 1973 (E Pacific) Riedl, 1971b North Carolina (NW Atlantic) Riedl, 1971b Oregon (NE Pacific) Gerlach, 1958 Maldives (Indian Ocean) Sterrer, 1991a Fiji (S Pacific) Riedl, 1971b North Carolina (NW Atlantic) Sterrer, 1970b Rovinj (Mediterranean) Riedl, 1971b North Carolina (NW Atlantic) Mamkaev, 1961 Barents Sea (NE Atlantic) Riedl, 1971b North Carolina (NW Atlantic) Ax, 1956 Sylt (NE Atlantic) Kirsteuer, 1969 Bimini Gnathostomula (NW Atlantic) tuckeri Farris, 1977 Sterrer, 1991a Fiji (S Pacific) Sterrer, 1991a Fiji (S Pacific) Sterrer, 1998 North Carolina (NW Atlantic)

Species author Type locality Synonyms and reference

Species

New Zealand

NW Atlantic

NE Atlantic

NE Australia

Distribution

Sterrer 1998

Karling 1962, Müller & Ax 1971 Farris 1977, Sterrer 1998

Sterrer 1998

Sterrer 2001

Sterrer 1998

References

142   2 Phylum Gnathostomulida

2.2 Morphology 

broad range of metazoan terminals places the gnathostomulids in a clade with rotifers, acoels, and myzostomids (Dunn et al. 2008), but the clade is poorly supported and rather dubious. Hence, at present, the best-founded hypothesis supports Gnathostomulida within Gnathifera, as a sister group to Micrognathozoa and Rotifera (including acanthocephalans).

2.2 Morphology 2.2.1 General and external morphology (Figs. 2.2–2.6) Adult gnathostomulids (Fig. 2.2) range in length from 230 µm (Problognathia minima) to 3600 µm (Haplognathia belizensis); an as yet undescribed filospermoid species from Belize is more than 12 mm long. Animals are wormshaped, cylindrical, with a diameter of 40–100 µm. Most

 143

are colorless-opaque to whitish or yellowish, but a few species (Fig. 2.3 A, B, D, G) are uniformly or patchily crimson or pink due to pigment in the periphery of the integument. The pre-oral region (rostrum) is elongated and pointed in Filospermoidea (Figs. 2.2 A, 2.3 A–J, and 2.6 A) but typically more rounded and head-like, bearing sensory cirri (Fig. 2.6 D, E), in Bursovaginoidea (Figs. 2.4, 2.5, and 2.6 B–H). Posteriorly, the body is either rounded or tapering to a more or less pronounced tail (Figs. 2.2 B and 2.4 A, B, D, F). Gnathostomulida are entirely monociliated (Fig. 2.7); the single-layered epidermis may also contain mucous and adhesive gland cells (Fig. 2.8) and various sensory cells including compound sensory cirri on the rostrum (Fig. 2.12). The nervous system (Fig. 2.11), located intraepithelially, consists of a frontal and a buccal ganglion, longitudinal nerve fibers, and cross-connections in the region of the male organ and tail. The musculature (Figs. 2.9 and 2.10) is invariably cross-striated and

Fig. 2.2: Habitus of the three gnathostomulid main lineages: (A) Filospermoidea, represented by Haplognathia rosea. (B) Scleroperalia, represented by Gnathostomula peregrina. (C) Conophoralia, represented by Austrognathia microconulifera. (Line art, after Sterrer 1986.)

144 

 2 Phylum Gnathostomulida

Fig. 2.3: Filospermoid gnathostomulid species. (A) Haplognathia ruberrima from the Red Sea. (B) H. rosea from Denmark (scale bar not available). (C) H. filum from Denmark (scale bar not available). D, H. ruberrima, juvenile specimen from the Red Sea (scale bar not available). (E) Cosmognathia manubrium from the Red Sea. (F) C. arcus from Belize. G. H. rufa from the Red Sea (scale bar not available). (H) H. lunulifera from Denmark. (I) H. simplex from Denmark. (J) Pterognathia ugera from the Red Sea. (LM micrographs of live specimens.)

located between the epidermis and the gut, with circular muscles outside of bundles of longitudinal muscles. The mouth is subterminal, ventral, leading to a muscular pharynx (Figs. 2.13–2.19) armed with cuticular structures (Figs. 2.20–2.35) in the form of an unpaired basal plate in the lower lip area, paired, pincer-like jaws, and (in some taxa) an unpaired jugum in the upper lip area. The intestine is straight, and there is no permanent anus

(Fig. 2.36). (Except for the mouth, all body openings in many species – anus, male and female gonopore – appear to be tissue connections). There are paired groups of unconnected protonephridia (Fig. 2.37). Body cavities and a parenchyma are lacking. Gnathostomulida are simultaneous hermaphrodites; fertilization is by copulation and development is direct (Fig. 2.47). A simple, unpaired ovary lies dorsally

2.2 Morphology 

 145

Fig. 2.4: Scleroperalian gnathostomulid species. (A) Gnathostomula paradoxa from Denmark. (B) Onychognathia rhombocephala from Florida. (C) Clausognathia suicauda from the Red Sea (scale bar not available). (D) Vampyrognathia varanus from the Red Sea. (E) Mesognatharia remanei from Denmark. (F) Semaeognathia sterreri from South Africa. (LM micrographs of live specimens.)

in the anterior body half, between the epidermis and the gut, with eggs maturing posteriorly. In Bursovaginoidea (Fig. 2.2 B, C), there is a dorsal vagina and bursa for sperm storage behind the ovary. Paired testes (or an unpaired testis) extend dorsolaterally in the posterior body half, converging on a male copulatory organ, which, in most Scleroperalia, contains a stylet. The male pore is subterminal and ventral. Sperm shape is diverse (Fig. 2.42), ranging from flagellate-filiform (Filospermoidea) (Fig. 2.44 A) to aflagellate and cone-shaped (Conophoralia) (Figs. 2.46) or droplet-shaped (Scleroperalia) (Figs. 2.44 B–D and 2.45).

2.2.2 Integument (Figs. 2.7 and 2.8) The epidermis of Gnathostomulida is single-layered and rather uniform throughout the phylum. It consists of irregularly polygonal cells (Fig. 2.7 C) 2–5 µm in diameter, each of which carries a single cilium 20–25 µm in length, with a regular 9+2 axoneme (Rieger & Mainitz 1977, Lammert 1986b). Each cilium originates in a centrally located, invaginated pit, 0.46 µm in diameter, and just as deep, surrounded by a circlet of 8 microvilli, which continue into the pit as eight ridges. The rest of the cell surface is covered with regularly arranged short (0.1–0.5 µm) microvilli

146 

 2 Phylum Gnathostomulida

Fig. 2.5: Conophoralian gnathostomulid species. (A) Austrognathia microconulifera from Bermuda. (B) A. microconulifera, juvenile specimen from Bermuda. (C) Austrognathia sp. from Hong Kong. (D) Austrognathia sp. from Florida. (LM micrographs of live specimens.)

(Fig. 2.7 A), at a density of 20–50/µm2. Microvilli have an electron-dense core filament that may be branched. Core filaments are connected to each other by means of a delicate network that lies immediately under the cell surface membrane. Usually covered with a surface coating assumed to be glycoprotein, microvilli are generally longer near the mouth and reach considerable length (4–6 µm) inside the buccal cavity. The basal body of the cilium is connected to a system of two rootlets, which are more or less aligned with the longitudinal body axis, in a straight line from the (rostral) cell surface to the (caudal) cell base (Fig. 2.7 B). The rostral rootlet shows a central fibrous portion that does not appear striated in longitudinal sections; its ventral, striated portion continues into the (striated) caudal rootlet. Behind, and perpendicular to the basal body, lies an accessory centriole whose distal end points to the left side of the animal. This centriole has a fibrillous connection (centriole appendix) to the caudal rootlet.

Immediately below the cell surface, the cytoplasm always contains vesicles or electron-dense inclusions, which are interpreted as the pigment-containing vesicles (Fig. 2.7 C) in red Haplognathia species. The cell nucleus is round to oval, 4  ×  1.5 µm (in Haplognathia rosea) to 2  ×  1 µm (in Gnathostomula mediterranea); a nucleolus has been found in all species studied. The endoplasmic reticulum is only weakly developed in the epidermal cells. Mitochondria are elongated, sausage-shaped (diameter 300–700 nm), and mostly concentrated in the distal part of the cell, with cristae generally weakly developed. The epidermis rests on a 200- to 400-nm thick basal lamina that is a distinct, homogeneous layer that is neither continuous with any intercellular matrix nor has any fibrous components (Rieger & Mainitz 1977). Epidermal cells originate near the basal lamina, as small cells with a condensed nucleus and adjacent diplosome, a poorly developed endoplasmic reticulum, and few mitochondria (Lammert 1986b). Ciliogenesis begins within a vesicle; as

2.2 Morphology 

 147

Fig. 2.6: Rostra of various gnathostomulid species. (A) Haplognathia gubbarnorum from Denmark. (B) Mesognatharia remanei from Denmark. (C) Agnathiella beckeri from Japan (scale bar not available). (D) Vampyrognathia horribilis from Florida. (E) Onychognathia rhombocephala from Florida. (F) Rastrognathia macrostoma from Denmark. (G) Gnathostomula paradoxa from Denmark. (H) Austrognathia sp. from Florida. (LM micrographs of live specimens.)

Fig. 2.7: Epidermal structures. (A) Epidermis in Haplognathia ruberrima. (SEM micrograph courtesy of Pierre-Yves Pascal.) (B) Transverse section through epidermal cell of Haplognathia simplex. (TEM micrograph courtesy of Reinhard M. Rieger and Gunde Rieger.) (C) The monociliated epidermis of Haplognathia sp. (LM micrograph of live specimen.) Cilia emerge in the center of the circular yellow areas. Abbreviations: ac, accessory centriole; cb, cell borders; ci, cilium; cr, caudal ciliary rootlet; n, nucleus; rr, rostral ciliary rootlet; sv, surface vesicle.

148 

 2 Phylum Gnathostomulida

the cell elongates and reaches the epidermal surface, the vesicle membrane merges with the apical cell membrane, forming the ciliary pit. Microtubules are arranged at the perpendicular centriole, forming the 9  ×  2 + 2 axoneme of the cilium. Degeneration of epidermal cells proceeds through loss of cilium and microvilli. The cell becomes ovoid and the nucleus translucent, until the cell is shed into the medium. Filospermoidea lack epidermal gland cells, whereas all members of at least the families Gnathostomulidae and Austrognathiidae have long, sausage-like mucous cells (Rieger & Mainitz 1977, Lammert 1989), which convey a “striped” appearance to the epidermis in members of these families. In Austrognathiidae, mucous cells are highly branched, forming a network in the epidermis. In Semaeognathia sterreri, mucous cells measure at least 20  ×  2 µm. Mucous cells are the only epidermal cells that lack both diplosome and cilium (Lammert 1986b). Rieger & Mainitz (1977) considered it possible that mucous cells are holocrine, being continuously produced from regular epidermal cells, whereas Lammert (1986b) argues for apocrine secretion (Fig. 2.8 B). Emergent gland cells, hugging the basal lamina, are characterized by an extensive rough endoplasmic reticulum (RER) that connects nucleus and Golgi complex on either end of the cell. Translucent primary vesicles arise from the cisternae, then move to the Golgi complexes where secondary vesicles up to 750 nm diameter are produced, containing the mucous secretion. Mature gland cells are packed with ovoid mucous vesicles to 7 µm long; mucus is freed by rupture of the apical cell membrane.

Two types of adhesive gland cells have been located in Gnathostomula paradoxa (Lammert 1989). The “ciliated” type (Fig. 2.8 A), found only along the ventral body surface, bears a single, short, 9  ×  2 + 2 cilium with an accessory centriole but lacks ciliary rootlets, and there is a circlet of 11 (instead of 8) microvilli. Cilium and microvilli are of equal length (1.6 µm), inside a proximal cavity filled with secretion, whereas the cytoplasm of the gland neck contains two types of vesicles. Distally, the cell terminates in a secretory pore. Given that Gnathostomula (and other Bursovaginoidea) have been observed to have a weak ability to adhere with their ventral/caudal surfaces only, the “ciliated” gland type is thought to facilitate adhesion in locomotion. Gland cells of the “diplosome” type are limited to the posterior fifth of the ventral body surface, around the male pore, in Gnathostomula (Lammert 1989). They are cylindrical (4 µm in diameter, 3 µm high), lack both cilium and microvilli but have an apical surface enlarged by protrusions with electron-dense caps. Given their location, they are suspected of having a role in adhesion during copulation. Spindle-shaped rhabdoid glands (2  ×  0.7 µm) are only found in Austrognathiidae, where they often form bundles in the tail region, possibly having an adhesive function. None of the gnathostomulid adhesive systems studied to date is considered homologous with duo-gland systems as described for Platyhelminthes and Gastrotricha (Tyler 1976).

Fig. 2.8: Epidermal structures. (A) The gland neck of a ciliary adhesive cell and (B) epidermal mucous gland cells of Gnathostomula paradoxa. Abbreviations: ac, accessory centriole; bb, basal body; bl, basal lamina; ci, cilium; ec, epidermal cells; mv, microvilli; cp, ciliary pit; ps, prosecretory vesicle; sev, secretory vesicle. (Line art courtesy of Volker Lammert.)

2.2 Morphology 

2.2.3 Musculature (Figs. 2.9 and 2.10) With the exception of smooth muscle fibers surrounding the testis in Austrognathia sp. (Lanfranchi & Falleni 1998), all musculature is cross-striated. It consists of individual cells in Filospermoidea, but at least in some Bursovaginoidea, it shows a tendency toward syncytial

Fig. 2.9: Depth-coded CLSM stacks showing phalloidin stained musculature in Gnathostomula peregrina. (A) Dorsal view. (B) Ventral view. Abbreviations: bu, bursa; dim(1–7), diagonal muscles 1–7; dlm, dorsolateral muscles; dom (1–5), dorsal longitudinal muscles 1–5; mo, mouth; pbu, prebursa; ph, pharynx; pss, proximal stylet sack; st, penis stylet; vlm, ventrolateral muscles; vm(1–4), ventral longitudinal muscles 1–4. (CLSM micrographs courtesy of Monika Müller.)

 149

organization (Lammert 1986b). Body-wall musculature is simple and situated between the epidermal basal lamina and the gut epithelium, with weak circular and diagonal fibers running outside of 3 pairs (Haplognathia) to 11 pairs (Gnathostomula) of stronger longitudinal bundles (Sterrer 1969, Tyler & Hooge 2001, Müller & Sterrer 2004). The musculature of pharynx and male copulatory organ is more complex and still insufficiently understood. According to Müller & Sterrer (2004), the body-wall musculature of Gnathostomula peregrina (Fig. 2.9) consists of 11 pairs (4 ventral, 1 ventrolateral, 1 dorsolateral, and 5 dorsal) of longitudinal muscles; anterior to the pharynx, the ventral and ventrolateral muscles anchor in a rostral chiasma. There are two types of diagonal muscles (thin fibers throughout the body, plus slightly thicker fibers, of which 7 pairs occur ventrally and 2 pairs dorsally; the ventral pairs originate ventrolaterally, then run caudoventrally to form a caudal chiasma with the corresponding fibers of the other side. Evenly spaced thin circular fibers densely gird the posterior half of the body, continuing less prominently into the anterior half. Thin fibers connect the pharynx with the ventral and dorsal longitudinal muscles. The difference in circular fiber density between posterior (high) and anterior (low density) body half is even more pronounced in Gnathostomula armata, an arrangement that Tyler & Hooge (2001) relate to the worm’s ability to navigate its interstitial environment. Contraction of the circular muscles of the hind body would force the anterior half into a restrictive space whereupon the longitudinal musculature would pull the rest of the body through. The body musculature of Filospermoidea is made up of outer circular fibers and a paired band of lateral longitudinal fibers (Sterrer 1969), but all are so sparsely provided with myofilaments that no “muscle tube” results (Lammert 1986b). Anterior to the mouth, the longitudinal muscles split into 5 pairs, which, often in close proximity with nerves, traverse the frontal ganglion from behind and end near the tip of the rostrum. Postpharyngeally, the lateral muscle band splits into smaller fibers, as does another muscle, the jaw retractor, which here joins the lateralis (Fig. 2.13). A post-pharyngeal cross section, therefore, shows 2 ventral and 4 ventrolateral fibers of the lateralis and 6 lateral plus 2 dorsolateral fibers of the retractor, per body side (Sterrer 1969). A dorsal musculature is lacking. Longitudinal fibers of either body side cross over and join caudally, possibly in conjunction with the muscles of the male copulatory organ. This simple organization of muscles explains all motions and changes of shape observed in Filospermoidea, especially the remarkable mobility of the rostrum,

150 

 2 Phylum Gnathostomulida

Fig. 2.10: CLSM maximum projections of phalloidin-stained musculature. (A) Anterior end of Gnathostomula paradoxa. (B) Posterior end of Gnathostomula paradoxa. (C) Anterior end of Haplognathia filum. (D) Detail of trunk musculature around the bursa of Valvognathia pogonostoma. (E) Detail of trunk musculature around the bursa of Gnathostomula paradoxa. (F) Detail of trunk musculature around the bursa of Valvognathia pogonostoma. Abbreviations: bu, bursa; cr, circular body musculature; lo, longitudinal body musculature; ph, pharynx; ps, penis stylet; pss, proximal stylet sack; sm, stylet musculature.

which seems to have both an exploratory and sensory function (Lammert 1986b), and the coiling of the body, a result of asymmetric contraction of longitudinal fibers (Sterrer 1969).

Myocytes are rather uniform within the phylum (Rieger & Mainitz 1977). They are elongated, with nucleus, adjacent diplosome, and other organelles located in an eccentric, sac-like protrusion (Lammert

2.2 Morphology 

 151

1986b). Myofilaments are cross-striated. In Bursovaginoidea, the Z-bands are composed of isolated dots in a plane band, whereas in Filospermoidea, the Z-elements are arranged in continuous filaments, resembling hypercontractile muscles found in some insects. Ectodermally derived muscle cells have been identified within the pharynx of Filospermoidea (Lammert 1986b), and while sharing most features with mesodermal muscles, they lie outside the basal lamina and their buccal cavity surfaces are covered by microvilli.

2.2.4 Nervous system (Fig. 2.11) Located intraepithelially, the nervous system consists of a frontal and a buccal ganglion, longitudinal nerves and a commissure each in the regions of the male organ and tail. It has been studied in Rastrognathia macrostoma (Kristensen & Nørrevang 1977); Gnathostomula paradoxa, Haplognathia rosea, H. simplex, Pterognathia meixneri (Lammert 1986b); and G. peregrina (Müller & Sterrer 2004). The G. peregrina nervous system (Fig. 2.11 A–C) consists of (1) a central (nervous) system with a frontal ganglion (brain), 1 pair of ventral longitudinal nerves ending in a terminal commissure, and 1 median ventral nerve originating from a commissure behind the pharynx; (2) 8–10 unipolar perikarya above and up to 10 bipolar perikarya in front of the brain; (3) a peripheral (nervous) system with 5 (1 unpaired, 2 paired) dorsal longitudinal nerves and 2–4 accompanying perikarya; and (4) a stomatogastric (nervous) system with 1 pair of buccal nerves that link the frontal with the buccal ganglion, with 6–8 perikarya dorsocaudally in the pharynx bulbus (Müller & Sterrer 2004). Lammert (1986b) reports an additional commissure between the ventral longitudinal nerves at the level anterior to the penis for both G. paradoxa and P. meixneri (Fig. 2.11 D, E). Kristensen & Nørrevang (1977) report 3 pairs of longitudinal nerves, and a buccal ganglion (Fig. 2.19C), in R. macrostoma. The nervous system of P. meixneri differs in the following. While the Gnathostomula frontal ganglion is in the shape of a flat cap whose basal lamina hugs the underlying rostral musculature, bordered laterally and frontally by perikarya, the frontal ganglion of Filospermoidea is elongated, rod-shaped (Fig. 2–10E), and in cross section seems enveloped along its entire length by basal lamina, giving the appearance of a subepithelial location. However, this basal lamina is wide open frontally, where nervous tissue is continuous with epidermis, suggesting that the entire ganglion is sunk-in (“eingesenkt”, Lammert 1986b) rather than subepidermal. Filospermoidea also have at least 3 paired and 2 unpaired

Fig. 2.11: The gnathostomulid nervous system visualized (A) with CLSM and (B–E) as schematic line art. (A) Depth-coded staining of FMRFamidergic nervous system in Gnathostomula peregrina. Arrow heads indicate fibers branching off from the ventral nerves (vn) to form the ventromedian nerve cord (vmn). (B, C) Reconstruction of FMRF-amidergic nervous system in G. peregrina, shown in (B) ventral and (C) dorsal views. (D) Reconstruction of nervous system in Gnathostomula paradoxa, based on serial sections. (E) Reconstruction of nervous system in Haplognathia simplex, based on serial sections. Abbreviations: bn, buccal nerves; bg, buccal ganglion; bpk, bipolar perikarya; cc, circumesophageal connectives; dln, dorsolateral longitudinal nerve cord; dmn, dorsomedian longitudinal nerve cord; fg, frontal ganglion; he, head; ln, lateral longitudinal nerve cord; mn, ventral main nerves; pc, penial neuronal connection; pk, perikarya; ta, tail; tc, terminal commissure; tr, trunk; vmn, ventromedian nerve cord; vn, ventral nerves. (CLSM micrograph courtesy of Monika Müller; B, C, line art after Müller and Sterrer 2004; D, E, line art after Lammert 1986.)

nerves that run forward, often tightly parallel with muscles, from the frontal ganglion (Lammert 1986b). In H. rosea and P. meixneri, all ganglia and nerves are intraepithelial, but nerves sometimes “stray” for short stretches below the basal lamina, where they presumably synapse with muscles (Lammert 1986b). Based on the presence of specific neurovesicles, Lammert (1986b) distinguishes at least six different types of neurons in the frontal ganglion of P. meixneri. A diplosome is present next to the nucleus in the perikarya of

152 

 2 Phylum Gnathostomulida

both the frontal and the buccal ganglia. Typical neuromuscular synapses have only rarely been documented (see Lammert 1986b, figure 16B).

2.2.5 Sensory structures (Fig. 2.12) The most conspicuous sensory organs, in light microscopy (LM), are found in Bursovaginoidea (Fig. 2.12 A), where the rostrum is set with 1–2 pairs of single, stiff cilia (apicalia) and 3–5 pairs of long (to 60 µm) compound ciliary cirri (frontalia, ventralia, dorsalia, lateralia and, in some species, postlateralia). Although these cirri usually appear stiff, they may also join the beat of epidermal cilia in unison. A row of single, stiff cilia (occipitalia) arranged longitudinally in the dorsal midline is present in all Gnathostomulida (Sterrer 1970a). “Ciliary pits”, which have been identified with LM in all lower Scleroperalia and in Conophoralia but seem to be lacking in Onychognathiidae and Gnathostomulidae, have yet to be studied with TEM. Situated rostrolaterally on the rostrum, between dorsalia and lateralia, they are distinguished by 1–2 bundles of cilia that originate deep in the epidermis, then bend parallel to the surface as they emerge. Those of the posterior bundle bend dorsally, dorsorostrally, and dorsocaudally, whereas those of the anterior bundle bend dorsorostrally only (Sterrer 1976, Tenuignathia rikerae). Some species also have an additional, unpaired pit situated at the very tip of the rostrum, whose cilia point ventrally. Lammert (1986a) using TEM identified at least 15 different single-cell receptors (Fig. 2.12 B–O) in Gnathostomulida. Arranged bilaterally in pairs or groups (rarely medially), they are concentrated in the epidermis of the rostrum, the buccal cavity, and around the mouth and male pore. All receptors are monociliary (with a 9  ×  2 + 2 axoneme, except for the pompom receptor, with 9  ×  2 + 0, and the perpendicular receptor, with 9 ×  2 + 1) and possess an accessory centriole next to the basal body. Those that lack a cilium have a diplosome made up of basal body and accessory centriole, or at least its rudiments. Ciliary rootlets, where present, ­correspond with those of epidermal cells; only the tri-radices receptor has three rootlets. Receptors differ mostly in (1) number, length, and orientation of ciliary rootlets; (2) dimensions of the ciliary pit; (3) number and length of circumciliary microvilli and villi of the cell surface; (4) length and structure of the cilium; (5) vesicle types; and (6) shape of receptors and their position in the epithelium. There are unipolar and

bipolar primary receptors as well as secondary receptors. Among the more prominent are: –– Bipolar primary receptors 1. Compound ciliary cirri: In Gnathostomula, each cirrus is composed of at least 10 cells, each (Fig. 2.12 E) bearing a cilium up to 50 µm in length. The cell is elongated, up to 10 µm, apically bearing 8 microvilli that surround the cilium. An accessory centriole is present, and the 10-µm-long, single, caudal rootlet reaches to the bottom of the cell, into the efferent axon. Compound ciliary cirri are typical of Bursovaginoidea but lack in Filospermoidea. 2. Receptors of the pre-oral sensory field (all species) are most similar to those of the cirri, but with shorter cilia (10 µm) and rootlet. 3. Pompom ciliary receptors (“Puschel” cilium receptors) Found only in the rostrum of Filospermoidea, up to 30 per body side – an 9  ×  2 + 0 cilium branches out, 1.5 µm from the base, into microtubuli, each within a cytoplasmic capitulum up to 1 µm long (Fig. 2.12 H). Accompanying microvilli, only 7 in number, also extend into distal capitula. Rootlets are lacking. Dendrites below the receptors run caudally to the frontal ganglion. 4. Receptors of the male pore (P. meixneri) also lack rootlets (Fig. 2.12 F). 5. Receptors of the mouth opening (P. meixneri), with rootlet rudiments only, are arranged in bilateral groups of at least 5 each (Fig. 2.12 G), on either side of the mouth (see Lammert 1986a, figure 43A, B) 6. Receptors with one vertical rootlet (P. meixneri) occur in all body regions (Fig. 2.12 I). 7. Buccal cavity receptors (all species), distinguished by very long (to 2.1 µm) microvilli, have only one (the caudal) rootlet. 8. Receptors with two vertical rootlets (P. meixneri) that run parallel to each other (Fig. 2.12 J). 9. Receptors with lamellar body (P. meixneri). Four such receptors, seemingly bulbous, concentriclamellar dendritic processes, lie under non-ciliated epidermal cells. 10. Free nerve endings, in the form of paired dendritic processes without ciliary structures, are found dorsally to the median edge of the jaws in P. meixneri and H. rosea (see Lammert 1986a, figure 23A, B). –– Unipolar primary receptors 1. Stereovilli receptor. Found only in the anterior end of G. paradoxa, this type has 8 circumciliary microvilli with distinct microfilamental structure (stereocilia).

2.2 Morphology 

 153

Fig. 2.12: Gnathostomulid sensory structures. (A) Line art showing rostral sensoria in Agnathiella beckeri (ventral view). (B) TEM section through spiral ciliary receptor of Gnathostomula paradoxa. (C) Schematic line art reconstruction of spiral ciliary receptor of G. paradoxa; inset shows the position of spiral ciliary receptors (scr) on the rostrum. (D) Epidermal cell. (E) Receptor from compound rostral cirri (G. paradoxa). (F) Receptor from near the male genital pore (Pterognathia meixneri). (G) Receptor with rudimentary rootlets from mouth opening (P. meixneri). (H) Pompom ciliary receptor (P. meixneri). (I) Receptor with one vertical rootlet (P. meixneri). (J) Receptor with two vertical rootlets (P. meixneri). (K) Receptor from occipitalia (G. paradoxa). (L) Tri-radices receptor (G. paradoxa). (M) Club-cilium receptor (G. paradoxa). (N) Balloon receptor (P. meixneri). (O) Perpendicular receptor (P. meixneri). Abbreviations: ac, accessory centriole; bb, basal body; c, cavity; cs, cavity sheath; cp, ciliary pits; d, dorsalia; f, frontalia; l, lateralia; mv, microvilli; n, nucleus; p, postlateralia; ph, pharynx; rc, rolled-up cilium; scr, spiral ciliary receptor; spr, sulcus pre-pharyngealis; v, ventralia. (TEM photo courtesy of Volker Lammert; A, line art after Sterrer 1971c; C–O, line art after Lammert 1986.)

154 

 2 Phylum Gnathostomulida

2. Club-cilium receptors (P. meixneri). Located in the rostral epidermis, above the neuropilem, this receptor has one rootlet and a short (to 4 µm) club-like cilium surrounded by two circlets of microvilli (Fig. 2.12 M). 3. Occipitalia (G. paradoxa) are arranged singly, at ~2-µm distance, along the dorsal midline of the rostrum (Fig. 2.12 K). Occipitalia are distinguished by a long rootlet (2.5 µm or longer) and centriole appendix (1.2 µm). Up to 13 occipitalia have been reported for H. rosea (Sterrer 1970a), and a group of up to 6 occipitalia, most angled caudally and some with a characteristic bend at half-length, have been encountered in most Bursovaginoidea (Sterrer 1970a, 1973a, 1976). 4. Tri-radices receptors (G. paradoxa), found laterally to the frontal ganglion, have 3 rootlets, 2 vertical, and 1 running horizontal-caudally (Fig. 2.12 L). The anterior vertical rootlet is interpreted as additional to the two-rootlet pattern of epidermal cells (Lammert 1986a). 5. Spiral ciliary receptors occur in pairs, only 15 µm apart from each other, on either side of the anterior tip of the rostrum, between apicalia and rostralia, in G. paradoxa (Fig. 2.12 B, C). Similar structures were found in Rastrognathia macrostoma (Nørrevang & Kristensen 1977) and Valvognathia pogonostoma (Nørrevang & Kristensen 1978), as well as in H. simplex and H. rosea (Lammert 1984). The dumbbell-shaped cell contains the nucleus, free ribosomes, and few mitochondria in one half, whereas the other half bears a single cilium rolled up 7–12 times in a cavity surrounded by a cytoplasmic sheath. The cavity may be rendered doughnut-shaped by a microvillus (H. simplex, G. paradoxa) or a solid cytoplasmic axis (H. rosea). The cilium of the spiral ciliary receptor in H. simplex (and H. filum) shows a somewhat dissipated arrangement of microtubules whereas H. rosea and G. paradoxa have the typical 9 + 2 pattern. Cilia, basal bodies, and accessory centrioles of neighboring spiral ciliary receptors stand rectangular to each other, and ciliary rootlets are lacking. The structure and rectangular configuration of spiral ciliary receptors suggest a similarity with the labyrinth of Cyclostomata and thus a function as a three-dimensional positioning organ (Lammert 1984). –– Secondary receptors 1. Balloon receptors (P. meixneri). Located in the epidermis above the penis, a rudimentary basal body

supports an electron-optically empty vacuole that seems to correspond to a ciliary pit, albeit missing circumciliary microvilli (Fig. 2.12 N). 2. Perpendicular receptors (P. meixneri). Located in the rostrum, the short (0.7 µm), 9  ×  2 + 1 cilium of this receptor runs parallel to the apical cell surface, under a partial roof of microvilli, with its basal ciliary structures offset at a 90° angle to equivalent structures in the epidermis (Fig. 2.12 O).

2.2.6 Intestinal system (Figs. 2.13–2.36) Located ventrally, always well behind the tip of the rostrum, the mouth opens into a spacious buccal cavity, which is ventrocaudally bordered by the muscular pharynx. The esophagus rises above the pharynx, then widens posteriorly into the intestine. In the lip region, a receptor-rich epidermis gives way to the single-layered but massive glandular epithelium of the buccal cavity. Buccal cells are voluminous (to 22 µm), non-ciliated but with a diplosome and large numbers of different vesicles, and are densely set with long (2.5 µm), glycocalyx-studded microvilli, which more or less fill the lumen of the buccal cavity. The intestine behind the pharynx is rather uniform, single-layered, lacking cilia throughout, but with a microvilli-studded lumen that ends, at least in Haplognathia, in a subterminal dorsal tissue connection (Fig. 2.36) that has been interpreted as a functional anus (Knauss 1979, but see Lammert 1986b). In LM, the dorsal gut cells of some species (e.g., Tenuignathia rikerae, Clausognathia suicauda; Fig. 2.4 C) seem particularly succulent or vacuolized (Sterrer 1976, 1992). Intracellular bacterial symbionts, as frequently found in thiobiotic meiofauna (Leisch et al. 2011) have never been identified in Gnathostomulida (Pascal et al. 2014).

2.2.6.1 Pharynx Located ventrally between the buccal cavity and the intestine, the muscular pharynx, with its cuticular armaments, provides a most distinctive character set for the phylum. All pharynx musculature is cross-striated and organized so as to effect the movement of basal plate and jaws. The filospermoid pharynx (Figs. 2.13–2.15), as analyzed by Sterrer (1969) in Haplognathia simplex (Fig. 2.13), consists of a jaw adductor (connecting the rostral apophyses ventrally), a jaw abductor (or diductor, connecting both jaw tips and running behind the symphysis), a jaw levator (connecting the jaw tips via a loop around the dorsorostral

2.2 Morphology 

 155

Fig. 2.14: Longitudinal section through the pharynx of Haplognathia simplex. Abbreviations: ja, jaws; mc, mouth cavity; mv, microvilli; pm, pharyngeal musculature. (TEM micrograph courtesy of Volker Lammert.)

Fig. 2.13: Pharynx of Haplognathia simplex, visualized with (A) LM and (B, C) line art, showing the pharyngeal musculature. All illustrations are shown to same scale. (A, B) Lateral view. (C) Dorsal view. Abbreviations: ab, jaw abductor; bp-ca, basal plate caudalis; bp-ro, basal plate rostralis; co, jaw constrictor; de, jaw depressor; dt, digestive tract; ja, jaws; la, trunk lateralis; la-co, cross-connection between trunk laterales; le, jaw levator; re, jaw retractor. (After Sterrer 1969.)

buccal cavity, a constrictor (crossing the jaws dorsally and running a loop rostrally around the mouth), and a retractor (inserting at the jaw tips then joining the lateral body musculature). Basal plate muscles: a bp caudalis (connecting the caudal sides of the basal plate via a loop behind the jaw symphysis) and a bp rostralis (connecting the rostral sides of the basal plate via a loop rostrally around the mouth). The pharynx of Pterognathia swedmarki (Sterrer 1969) is somewhat more compact, with the constrictor broadened to enclose a bulb around the jaw symphysis.

Considerably more complex, the scleroperalian pharynx (Figs. 2.16–2.18) in LM appears capsular and often tripartite. It has been studied (Herlyn & Ehlers 1997, Tyler & Hooge 2001) and reviewed in detail by Sørensen at al. (2003), who describe the musculature of Gnathostomula armata as follows (Fig. 2.17). Two loop-shaped unpaired muscles and 2 paired muscles control the basal plate. The bp dorsalis forms an almost closed ring and attaches to the tectum caudalis, from where it runs laterally, then rostrally until the terminals bend inward rostrally of the basal plate. The other unpaired muscle, the bp rostralis, attaches caudally to the alae laterals and runs dorsorostrally until it forms a loop rostrally to the basal plate. The paired bp retractors attach on the caudal lobes of the alae laterals from where they run caudally and attach ventrally to the pharyngeal wall. At least one group of unpaired and four groups of paired muscles attach to the jaws. The caudal-most muscles (abductors) are large and loop-shaped, located posterior to the cauda, and extending rostrally to each involucrum where they anchor. The caudal part of the jaws is located inside the abductor loops. Most caudally, a pair of inclinators attach on the cauda and extend dorsally to anchor in the dorsal wall of the pharynx. The strongest muscles, the diductors, are located rostral to the inclinators. They appear to attach in the involucrum and run dorsally to their anchor points in the pharyngeal wall. Another pair of muscles, the levators, also attach on the dorsal side of the apophyses from where they run dorsally to anchor in the dorsal pharyngeal wall. In addition to the muscles that attach to hard parts, there is one large, loop-

156 

 2 Phylum Gnathostomulida

Fig. 2.15: Pharyngeal hard parts and musculature in Haplognathia simplex, visualized with combined LM and CLSM (A, C, D, E) and only CLSM of phalloidin stained musculature (B, D, F, H). (A, B) Optic section through dorsal part of pharynx. (C, D) Optic section through the dorsomedian part of pharynx. (E, F) Optic section through the ventromedian part of pharynx. (G, H) Optic section through ventral part of pharynx. Abbreviations: ab, jaw abductor; bp, basal plate; bp-ca, basal plate caudalis; bp-ro, basal plate rostralis; co, jaw constrictor; de, dentarium; ja, jaws; la, trunk lateralis; ra, rostral apophyse; re, jaw retractor; sy, symphysis.

2.2 Morphology 

 157

Fig. 2.16: Pharyngeal hard parts and musculature in Rastrognathia macrostoma, visualized with SEM (A, B), CLSM of phalloidin stained musculature (C, E, G, I), and combined LM and CLSM (D, F, and H). (A) Jaws, dorsal view. (B) Jaws, ventral view. (C). Maximum projection of CLSM images through pharynx. (D, E) Optic section through dorsal part of pharynx. (F, G) Optic section through median part of pharynx. (H, I) optic section through ventral part of pharynx. Abbreviations: a.ad, addental apophysis; ap-ph ab; apophysis-pharyngeal abductor; ap-ps ab; apophysis-pseudofulcrum abductor; de, dentarium; ps, pseudofulcrum; ps-ph, pseudofulcrum-pharyngeal abductor.

158 

 2 Phylum Gnathostomulida

Fig. 2.17: Reconstruction of pharyngeal musculature in Gnathostomula peregrina. (A) Pharynx and pharyngeal hard parts in dorsal view. (B) Basal plate in dorsal view. (C) Pharynx and pharyngeal hard parts in lateral view. Abbreviations: ab, jaw abductors; bp, basal plate; bp-do, basal plate dorsalis; bp-re, basal plate retractor; bp-ro, basal plate rostralis; co, jaw constrictors; di, jaw diductors; in, jaw inclinators; ja, jaw; le, jaw levator; mo, mouth; pb, pharynx bulb. (Line art after Sørensen et al. 2003.)

Closing the jaws is a rapid but passive movement that happens when the diductors relax.

2.2.6.2 Jaws

Fig. 2.18: Horizontal section through the symphysis (sy) of the jaws of Haplognathia rosea. Note the translucent rods (ro) with electron-dense cores that make up the lamella symphyses (ls). (TEM micrograph courtesy of Volker Lammert.)

shaped constrictor whose terminals are located medially, caudal to the jaws. From this point, they run laterally and then bifurcate and continue rostrally, perhaps forming two loops rostral to the basal plate. The constrictors as well as the ends of the diductors contribute to the wall of the pharyngeal bulb. From live observations and structural analyses, the following feeding scenario can be invoked. Acting as a scraper that dislodges food (such as filamentous bacteria) from the substrate, the basal plate is moved forward by contraction of the bp rostralis, then tilted downward and out of the mouth by the bp dorsalis so that the (dorsal) teeth point ventrally. Withdrawal is by relaxation of the bp dorsalis, coordinated with contraction of the bp retractors. Loosened food is then embraced by the jaws, which are moved forward and out of the mouth by the constrictor while being opened (and tilted ventrally) by the diductors.

The pharyngeal hard parts of gnathostomulids consist of forceps-like jaws, an unpaired, ventral basal plate and, in some species, an unpaired, dorsal jugum. Jaws are present in most species except in Agnathiella (Sterrer 1971c), whereas a basal plate is lacking in several species across the phylum. Overviews of the gnathostomulid hard parts and their morphology are given by Riedl & Rieger (1972), Sterrer (1972), and Sørensen & Sterrer (2002). The pharyngeal hard parts (Figs. 2.20–2.35) are cuticular structures that are formed by the ventral and ventrolateral pharyngeal epidermis, and in some instances, epidermal cells situated in compartments inside the jaws (Kristensen & Nørrevang 1977, Sørensen 2000). Some parts of the jaws consist of a homogenous material, whereas others are composed of densely packed rod-like structures (Figs. 2.18 and 2.22 B, E). When viewed in cross section with TEM (Fig. 2.18), the rods appear to consist of a lucent material with an electron dense core (Rieger & Tyler 1995, Herlyn & Ehlers 1997). An identical substructure is found in the scleropilar rods in jaws of Micrognathozoa and Rotifera (Ahlrichs 1995b, Kristensen & Funch 2000), and they are considered homologous with the gnathostomulid jaws. During organogenesis, jaws and basal plate are laid down in situ by surrounding cells, rather than from a growth region (Sterrer et  al. 1985, Lammert 1991). Unlike the jaw apparatuses in Micrognathozoa and Rotifera, the gnathostomulid jaws are not made up from multiple individual units (sclerites), but consist of one major, forceps-like element only. However, this single unit can be subdivided

2.2 Morphology 

Fig. 2.19: Pharyngeal region of Rastrognathia macrostoma. (A, B) Oblique longitudinal sections through pharynx and jaws, dorsal is up and anterior is right. (C) Cross section through buccal ganglion, posterior to jaws. Abbreviations: a.ad, addental apophysis; de, dentes; j.mu, jaw musculature; ls, lamella symphysis; mo, mouth opening; mv, microvilli; n.bg, nuclei of nervous cells in buccal ganglion; n.br, nuclei of nervous cells in brain; pf, pseudofulcrum. (TEM sections courtesy of Reinhardt M. Kristensen.)

 159

into different morphologically or functionally defined main regions (Riedl & Rieger 1972, Sørensen & Sterrer 2002). The articularium constitutes the main pincers of the jaws and is formed by two paired structures, the lamellae symphyses, that fuse caudally at the symphysis. Rostrally, the tooth-bearing parts of the lamellae symphyses make up the dentarium. Paired lamellae may attached to the lateral sides of the lamellae symphyses and extend laterocaudally, forming 1 or 2 pairs of apophyses that anchor the jaws in the pharyngeal wall or form muscle attachment sites. These elements are referred to as the suspensorium. Anchoring apophyses may also be situated around the caudal symphysis of the articularium, and these are also regarded a part of the suspensorium. An apical specialization of the suspensorium may be present lateral to the dentarium. Here, the lamellae of the suspensorial apophyses may wrap around themselves, forming a pair of cone-shaped structures. Such elements are referred to as the involucrum. The rod-like jaw composition is most conspicuous in the lamellae symphyses, whereas the involucrum and suspensorium consist of the more homogenous material. Across the gnathostomulid taxa, three different types of jaws can be recognized: “the compact type”, “the open lamellar type”, and “the fused lamellar type” (Sterrer 1972). The compact jaw type is relatively simple and is found in species of Haplognathia (Figs. 2.20 A, 2.21 A–D, 2.22 B–I, and 2.48). The articularium is well developed, and the lamellae symphyses are equipped with 1 or 2 pairs of strong apophyses. Caudally, the lamellae symphyses fuse in a symphysis that may be rounded, disc-shaped, or laterally expanded. The dentarium carries teeth that are either acicular or digitiform and arranged in clusters or distinct rows. An involucrum is not present. Jaws of the compact jaw type are embedded in the pharyngeal wall, which prevents protrusion of the jaws through the mouth. Variations of the disparate open lamellar type are found among a variety of taxa, including species of the filospermoid genera Pterognathia and Cosmognathia and all Scleroperalia except Gnathostomulidae. The most characteristic features of the open lamellar type are the formation suspensorial apophyses that serve as muscle attachment sites and the jaws being embedded in the pharyngeal wall more posteriorly, which allow the jaws to move more freely and enable protrusion through the mouth. The jaws of two filospermoid genera, Pterognathia (Fig. 2.23 G–I) and Cosmognathia (Figs. 2.20 B, 2.21 E–F, and 2.23 C–D), are stout, with a prominent suspensorium made up by well-developed rostral and caudal apophyses composed of a homogenous but non-flexible material.

160 

 2 Phylum Gnathostomulida

Fig. 2.20: Jaws and basal plate of (A, B) Haplognathia, representing the compact jaw type and (C, D) Cosmognathia, representing the filospermoid version of the open lamellar jaw type. (A) Basal plate, ventral view. (B) Jaws, dorsal view. (C) Basal plate, ventral view. (D) Jaws, dorsal view. Abbreviations: ca, caudal apophyse; de, dentarium; dnt, denticles; lp, lateral process; ls, lamella symphyses; mp, medial process; mt, median teeth row; ra, rostral apophyse; sy, symphysis; vt, ventral teeth. (Line art.)

Fig. 2.21: Jaws (A–F) and basal plates (B, E, F) of filospermoid gnathostomulid species. (A) Haplognathia simplex from Denmark. (B) H. asymmetrica from California. (C) H. rosea from the Red Sea. (D) H. rufa from the Red Sea. (E) Cosmognathia arcus from Belize. (F) C. manubrium from the Red Sea. (LM micrographs of prepared jaws, A, and live specimens, B–F.)

2.2 Morphology 

 161

Fig. 2.22: Basal plates (A) and jaws (B–I) of selected species of Haplognathiidae: Haplognathia. All jaws belong to the compact type. (A) Haplognathia simplex from Denmark ventral view. (B) Detail showing cauda of H. rosea from Denmark. (C) Detail showing dentarium of H. simplex from Denmark. (D) H. simplex from Denmark, dorsal view. (E) H. rosea from Denmark, ventral view. (F) H. gubbarnorum from Denmark, ventral view. (G) H. asymmetrica from Panama, dorsal view. (H) H. ruberrima from Denmark, ventral view. (I) H. lunulifera from Denmark, dorsal view. (SEM micrographs.)

162 

 2 Phylum Gnathostomulida

Fig. 2.23: Basal plates (A, B, E, F) and jaws (C, D, G–I) of selected species of Pterognathiidae: Cosmognathia and Pterognathia. All jaws belong to the open lamellar type. (A) Cosmognathia manubrium, ventral view. (B) C. aquila, frontal view. (C) C. manubrium, ventral view. (D) C. aquila, ventral view. (E) Pterognathia swedmarki, dorsal view. F., P. swedmarki, ventral view. (G) P. crocodilus, ventral view. (H) Detail showing dentarium of P. swedmarki. I. detail showing dentarium of P. atrox. (SEM micrographs.)

2.2 Morphology 

 163

Fig. 2.24: Jaws and basal plate of (A, B) Gnathostomaria and (C, D) Onychognathia, both representing the bursovaginoid version of the open lamellar jaw type. (A) Basal plate, ventral view. (B) Jaws, dorsal view. (C) Basal plate, ventral view. (D) Jaws, dorsal view. Abbreviations: cd, cauda; de, dentarium; dn, dentes; dnt, denticles; dt.t, dorsoterminal tooth; fv, fenestra ventralis; ls, lamella symphyses; sl, shoulder lamella; sy, symphysis; vt, ventral teeth. (Line art.)

Fig. 2.25: Jaws of scleroperalian gnathostomulid species. (A) Tenuignathia rikerae from the Red Sea. (B) Mesognatharia remanei from Denmark. (C) Vampyrognathia varanus from the Red Sea. (D) Rastrognathia macrostoma from Denmark. (LM micrographs of prepared jaws, B, and live specimens, A, C, D.)

164 

 2 Phylum Gnathostomulida

Fig. 2.26: Jaws of selected species of Mesognathariidae: Tenuignathia, Mesognatharia, and Labidognathia. All jaws belong to the open lamellar type. (A) Tenuignathia rikerae, lateral view. (B) Mesognatharia remanei, dorsal view. (C) Labidognathia longicollis, ventral view. (D) Detail showing dentarium of T. rikerae, lateral view. (E) Detail showing dentarium of M. remanei, dorsal view. (F) Detail showing dentarium of L. longicollis, ventral view. (SEM micrographs.)

Fig. 2.27: Basal plate (A) and jaws (B, D) of Gnathostomariidae: Gnathostomaria lutheri. The jaws belong to the open lamellar type. (A) Ventral view. (B) Detail showing dentarium in dorsal view. (C) Ventral view; note the basal plate behind the jaws. (D) Dorsal view. (SEM micrographs.)

2.2 Morphology 

 165

Fig. 2.28: Jaws of selected species of Onychognathiidae: Onychognathia and Vampyrognathia. All jaws belong to the open lamellar type. (A) Onychognathia rhombocephala, dorsal view. (B) O. rhombocephala, ventral view. (C) Undescribed species of Onychognathia from Italy, dorsal view. (D) Vampyrognathia minor, lateral view. (SEM micrographs.)

166 

 2 Phylum Gnathostomulida

Fig. 2.29: Basal plates (A, B) and jaws (C, D) of Onychognathiidae: Valvognathia pogonostoma. The jaws belong to the open lamellar type. (A) Ventral view. (B) Dorsal view. (C) Ventral view. (D) Dorsal view. (SEM micrographs.)

2.2 Morphology 

Fig. 2.30: Jaws of Rastrognathiidae: Rastrognathia macrostoma. The jaws belong to the open lamellar type. (A) Ventral view. (B) Dorsal view. (C) Lateral view. (SEM micrographs.)

 167

The anchoring symphysis at the caudal tip of the lamellae symphyses is usually laterally expanded. The dentarium is horizontally bipartite, hence forming paired dorsal and ventral rows of acicular teeth. An involucrum is not present. The jaws of the genera Valvognathia (Fig. 2.29 C, D), Rastrognathia (Figs. 2.25 D and 2.30), Onychognathia (Figs. 2.24 D and 2.28 A–C), Vampyrognathia (Figs. 2.25 C and 2.28 D), and Nanognathia (all belonging to the family Onychognathiidae) have a less prominent suspensorium. It is most developed laterally near the rostral tips of the lamellae symphyses, where it may form compartments that contain epidermal cells. The suspensorial lamellae also tend to wrap and form structures that resemble a cone-shaped involucrum. Pairs of extremely thin and delicate apophyses or lamellae further extend from the caudal edges of each involucrum. An isolated part of the suspensorium occurs around the symphysis, where an unpaired extension forms a club-shaped, caudally extending cauda. The dentarium always consists of long, well-developed teeth that bend and form a large basket. In species of Nanognathia, Onychognathia, and Valvognathia, the teeth can clearly be divided into a dorsal, a median, and a ventral portion that attach along a U-shaped rim, whereas the teeth in species of Vampyrognathia and Rastrognathia all appear to emerge and spread out from the same point.

Fig. 2.31: Jaws (A–D) and basal plates (E, F) of Problognathiidae: Problognathia minima. The jaws belong to the open lamellar type. (A) Ventral view. (B) Detail showing dentarium in ventral view. (C) Dorsal view. (D) Detail showing dentarium in dorsal view. (E) Ventral view. (F) Detail showing teeth of basal. (SEM micrographs.)

168 

 2 Phylum Gnathostomulida

Fig. 2.32: Jaws and basal plate of (A, B) Gnathostomula, (C, D) Austrognathia, and (E, F) Austrognatharia, all representing the fused lamellar jaw type. (A) Basal plate, dorsal view. (B) Jaws, ventral view. (C) Basal plate, ventral view. (D) Jaws, dorsal view. (E) Basal plate, ventral view. (F) Jaws, ventral view. Abbreviations: ap, apophysis; cd, cauda; ct, caudal tectum; dtr, dorsal tooth row; ll, lateral lobe; lw, lateral wing; ml, median lobe; mtr, median tooth row; rdd, rostrodorsal dentes; rw, rostral wing; rwd, rostroventral denticles; sl, shoulder lamella; tt, terminal tooth; vtr, ventral tooth row. (Line art.)

The suspensorium in jaws of Labidognathia longicollis (Fig. 2.26 B, F), Tenuignathia rikerae (Figs. 2.25 A and 2.26 A, D), and Mesognatharia remanei (Figs. 2.25 B and 2.26 C, E) forms a fully developed involucrum. Each involucrum opens through a caudal window, and extensions from the lateral and dorsal margins of the windows form a pair of large, but rather thin, shoulder lamellae. The third jaw type, the fused lamellar type, is present in species of the families Austrognathiidae, Triplignathia, Austrognathia (Figs. 2.32 C, D, 2.34 C, E, and 2.35 C, D, G), and Austrognatharia (Figs. 2.32 F, 2.34 B, and 2.35 E, F); and Gnathostomulidae, Chirognathia (Fig. 2.33 F), Gnathostomula (Figs. 2.32 B and 2.33 E, G), Ratugnathia, Corculognathia, and Semaeognathia (Fig. 2.33 C, H). This type was previously thought to have evolved only once (Sterrer 1972, Sørensen 2002), but more recent phylogenetic studies based on combined morphological and molecular data indicate that the jaw types developed

independently within the two families (Sørensen et  al. 2006). In the fused lamellar type, the rostral part of the suspensorium always forms an involucrum that serves as a major muscle attachment site, and an apophyse extends caudally from each involucrum. An isolated part of the suspensorium furthermore forms a cauda around the symphysis of the articularium. In species of Austrognathiidae, the cauda is unpaired and clubshaped, whereas it forms paired caudal extensions in species of Gnathostomulidae. The dentarium is located on the inferior surface of the involucrum and forms two or three rows of teeth. A single tooth is often much more prominent than the others and is referred to as the terminal tooth. In species of Austrognathiidae, the articularium and suspensorium is extremely delicate and feebly visible in scanning electron microscopy (SEM) preparations, whereas these regions are well-developed in species of Gnathostomulidae.

2.2 Morphology 

 169

Fig. 2.33: Basal plates (A, B) and jaws (C, H) of selected species of Gnathostomulidae: Chirognathia, Gnathostomula, and Semaeognathia. All jaws belong to the closed lamellar type. (A) Chirognathia dracula, dorsal view. (B) Gnathostomula armata, dorsal view. (C) Semaeognathia sterreri, ventral view. (D) Chirognathia dracula, ventral view. (E) Gnathostomula armata, ventral view. (F) Detail showing dentarium of Chirognathia dracula. (G) Detail showing dentarium of Gnathostomula armata. (H) Detail showing dentarium of Semaeognathia sterreri. Abbreviations: dt, dorsal tooth row; mt, medial tooth row; vt, ventral tooth row. (SEM micrographs.)

2.2.6.3 Basal plate and jugum The basal plate is an unpaired element situated ventral and slightly rostral to the jaws, near the posterior rim of the mouth opening. In species of Filospermoidea, the basal plate is either a rather simple structure without clear regionalization or a transverse rod with a medial group of teeth. In Haplognathia (Figs. 2.20 A, 2.21 B, 2.22 A, and 2.48), it is, if present at all, often a small triangular, oval, diamond- or droplet-shaped element, lopsided in at least one species (H. asymmetrica, Fig. 2.21 B), and its functionality remains questionable in

some species. In species of Cosmognathia (Figs. 2.20 C, 2.21 E, F, and 2.23 A, B) and Pterognathia (Fig. 2.23 E, F), the basal plate is larger and forms a transverse rod. Median clusters of acicular teeth suggest that it plays an active role during feeding. Among species of Bursovaginoidea, the basal plate is a flattened, somewhat crescentic structure and appears to be differentiated into a clear five partitions. In species of Goannagnathia, Vampyrognathia, Nanognathia, Onychognathia, Valvognathia (Fig. 2.29 A, B), Problognathia (Fig. 2.31 E, F), and Gnathostomaria (Fig. 2.24 A and 2.27 A), the five partitions are often evident as five more or less equally sized plates that are separated by

170 

 2 Phylum Gnathostomulida

Fig. 2.34: Jaws (A–C, E) and basal plates (A–D) of conophoralian gnathostomulid species. (A) Austrognatharia barbadensis from Barbados. (B) Austrognatharia sp. from the Red Sea (Israel), scale not available. (C) Austrognathia hymanae from Barbados. (D, E) Austrognathia riedli forma marisrubri from the Red Sea, scale not available. (LM micrographs of live specimens.)

longitudinal or diagonal fibulae. The anterior margin of the medial plate, and sometimes the margins of the paramedial plates as well, may carry small teeth or denticles. In Labidognathia longicollis, the basal plate has been extended in the longitudinal direction and appears longer than wide. In species of Tenuignathia and Mesognatharia, the basal plate is more rounded and thinner and appears to be partly reduced. The basal plate in species of Triplignathia, Austrognathia (Figs. 2.34 C, D and 2.35 A, H) and Austrognatharia (Figs. 2.34 A, B and 2.35 B) is extremely delicate and is best observed with phase contrast LM, as it will appear as a thin membrane only in SEM. Only the teeth are well developed. Among these species, the lateral plates are expanded and form a pair of laterocaudally projecting

wings. Furthermore, the teeth have changed position and occur on the caudal margins of the medial and ­paramedial plates. In species of Gnathostomulidae, the denticulation is also well developed and forms a median group of welldefined teeth on the rostral margin of the medial plate (Fig. 2.33 A, B). Ventral to this tooth group, the margins of the median and paramedian plates furthermore carry a row of densely set denticles. Within this family, further differentiation of the plates can be observed. Whereas the medial plate retains its shape, the paramedial ones extend rostrally, forming rostral wings. The rostral wings are only slightly developed in Chirognathia dracula, but are very prominent in species of Gnathostomula, Semaeognathia, ­Corculognathia, and Ratugnathia. In these latter

2.2 Morphology 

 171

Fig. 2.35: Basal plates (A, B, H) and jaws (C–G) of selected species of Austrognathiidae: Austrognathia and Austrognatharia. All jaws belong to the closed lamellar type. Note that all jaw parts, except the dentaria, are so delicate that they hardly can be visualized by the SEM. (A) Austrognathia christianae, ventral view. (B) Austrognatharia moorensis, ventral view. (C) Right and (D) left dentarium of Austrognathia christianae, ventral view. (E) Right and (F) left dentarium of Austrognatharia moorensis, ventral view. (G) Austrognathia christianae, ventral view. (H) Detail showing basal plate teeth of Austrognathia riedli. Abbreviations: dt, dorsal tooth row; tt, terminal tooth; vt, ventral tooth row. (SEM micrographs.)

four genera, the lateral plates are also expanded, forming lateral wings. Examinations of the pharyngeal musculature in Gnathostomula armata show that the lateral wings act as muscle attachment sites for several muscles, whereas the equally developed rostral wings do not have any attached musculature (Sørensen et al. 2003). A basal plate is missing in species of Clausognathia, Paucidentula, Paragnathiella, and Rastrognathia. Another unpaired, pharyngeal structure, the jugum, is present in species of Gnathostomula, Semaeognathia, Corculognathia, and Ratugnathia only. The element is a rigid, but not sclerotized, crescentic structure that is situated rostrally to the mouth opening. Unlike the other pharyngeal hard parts, the jugum dissolves during sodium hypochlorite treatment, which suggests that it

has a different biochemical composition. The jugum has no associated musculature, and it has most probably a supportive function.

2.2.7 Body cavities and connective tissue Gnathostomulida lacks a coelom, nor are there cavities or mesenchyme between organ system (Mainitz 1979, Rieger 1985). Body musculature and intestinal epithelium throughout most of the body are in immediate contact with the basal lamina of the overlying epidermis. Potential mesenchyme cell types were identified in the caudal intestinal epithelium in Filospermoidea (Lammert 1986b).

172 

 2 Phylum Gnathostomulida canal cell communicate with the cilium duct where the microvilli end, the canal cell nucleus is tightly wrapped around the distal cilium duct, and the nephroporus cell lacks a central canal, so that protonephridial fluid must pass through its cytoplasm.

2.2.9 Reproductive organs (Figs. 2.38–2.41) 2.2.9.1 Female organs

Fig. 2.36: Cross section through Haplognathia gubbarnorum in the region of the anal tissue connection Arrows indicate the interruption of the basal lamina). Abbreviations: gt, gut, sd sperm duct, sdg sperm duct gland. (TEM micrograph courtesy of Elizabeth B. Knauss.)

2.2.8 Excretory system (Fig. 2.37) The excretory system of Gnathostomulida consists of up to 10 pairs of unconnected protonephridia, which are serially arranged, probably in groups, lateral to the pharynx, bursa, and penis (Graebner 1968a, Kristensen & Nørrevang 1977, Lammert 1985). There is no common canal system. Each protonephridium is made up of three cells: a ciliated terminal cell, a canal cell, and a nephroporus cell (Fig. 2.37). Although the former two are located subepidermally, the nephroporus cell emerges into the epidermis. As analyzed in Gnathostomula paradoxa (and Haplognathia rosea) by Lammert (1985), the terminal cell is cup-like, with nucleus, dictyosome, and most mitochondria concentrated in the apical half while the cytoplasm of the other half forms a porous filtration area. Originating in the terminal cell, an 11-µm-long, 9+2 cilium (with a caudal rootlet but lacking a rostral rootlet) protrudes into the protonephridial duct, and thus into the canal cell, accompanied along its entire length by eight microvilli. The voluminous canal cell, packed with mitochondria and riddled with a highly connected lacunar system, protrudes through the basal lamina into the epidermis, forming an unbranched outlet canal. This protrusion is in turn enveloped, doughnut-like, by the nephroporus cell. Both canal cell and nephroporus cell have a diplosome but lack a cilium. The protonephridium of Haplognathia rosea differs mainly in that the microvilli of the terminal cell are much shorter than the cilium, the lacunae of the

Remarkably uniform throughout the phylum, an unpaired, elongated pear-shaped ovary lies dorsally between the gut and the epidermis, extending from behind the pharynx to just behind midbody region (Mainitz 1983). The ovary is not enveloped by a tunica; it usually consists of 4–6 oocytes whose dimensions increase posteriorly, with the last considerably larger (50–60 µm) than the others, taking up the entire width of the body (Falleni 1993). In Bursovaginoidea, the single mature egg is located immediately in front of the bursa, and in the diminutive Problognathia minima, the egg wraps around and extends behind the bursa system (Sterrer & Farris 1975). Located immediately behind the ovary, the bursa in Conophoralia is a simple soft pouch posteriorly connected to a dorsal vagina; both ovary and vagina may be ­facultative (Sterrer et  al. 1985). The bursa often contains one or two (“Siamese twin”) conuli of characteristic appearance: shorter than conuli from the testis and with a hollow and coarse-granular cone body. By contrast, Scleroperalia have a bursa system that consists of an anterior, round or miter-shaped bursa joined posteriorly by a more or less globular prebursa. In Gnathostomulidae, the bursa is made up of three groups of concentrically layered cells that meet in three longitudinal ridges (cristae) (Fig. 2.38 A), tapering anteriorly to a nozzle (mouthpiece) (Fig. 2.38 B) that points toward the mature egg. Bursa, cristae, and mouthpiece appear hard due to strongly developed desmosomes (Mainitz 1983). The mouthpiece surrounds a narrow fertilization canal that usually contains elongate sperm, and the bursa is often tightly packed with sperm. The prebursa (Fig. 2.38 C and 2.40 A), derived from female tissue, seems to receive rather than store sperm and sometimes contains structures that may be remnants of male stylets (Sterrer 1973). The scleroperalian vagina, located dorsally between bursa and prebursa, is temporary in most species and in TEM can be identified as a tissue connection (Mainitz 1983). It is likely that this site serves for both fertilization (hypodermic impregnation) and oviposition.

2.2 Morphology 

 173

Fig. 2.37: Protonephridium of Gnathostomula paradoxa. (A) Complete protonephridium in longitudinal section. (B) Cross section through terminal cell with filtration area. (C) Cross section through median part of canal cell. (D) Cross section through terminal part of canal cell. Abbreviations: ac, accessory centriole; bb, basal body; bl, basal lamina; CC, canal cell; ci, cilium; cd, cilium duct; cr, ciliary rootlet; crs, structures resembling ciliary rootlets; di, diplosome; fi, filtration area; mi, mitochondria; mv, microvilli; n, nucleus; NP, nephridiopore cell; oc, outlet canal; TC, terminal cell; tl, tubules of lacunar system. (Schematic reconstruction after Lammert 1985.)

Fig. 2.38: Scleroperalian female bursa morphology. (A) Cross section through bursa of Gnathostomula jenneri. (B) Longitudinal section through bursa mouthpiece of Gnathostomula mediterranea. (C) Reconstruction of bursa of Labidognathia longicollis. (D) Reconstruction of bursa of G. jenneri. Abbreviations: bmp, bursa mouthpiece; bu, bursa; cc, collar cells; ds, mature dwarf sperm; pb, prebursa. (TEM micrographs, A, B, and schematic reconstruction, C, D, after Mainitz 1983.)

174 

 2 Phylum Gnathostomulida

Fig. 2.39: Reproductive system in Gnathostomula sp. (A, C–E) and Gnathostomula paradoxa (B). (A) Section through proximal stylet sack of the male stylet apparatus. (B) Oblique section through distal part of stylet apparatus. (C) Bursa. (D) Two early dwarf sperm spermatids. (E) Mature dwarf sperm inside bursa. Abbreviations: bl, basal lamina; bu, bursa; ds, mature dwarf sperm; ir, inner rod; m.ss, muscular layer around stylet sack; mi, mitochondria; mp, micropodia; n.spi, nuclei of spermatids; or, outer rod; rc, rod cells; ss, stylet sheath. (TEM micrograph courtesy of Reinhardt M. Kristensen.)

2.2.9.2 Male organs Male located are located in the posterior body region and consist of paired or unpaired testes and vasa deferentia, a penis, and a genital pore. Testes are paired in all Scleroperalia, whereas Conophoralia (and most likely all Pterognathiidae) have an unpaired dorsal testis. Copulatory organs in Filospermoidea were analyzed by TEM (Knauss & Rieger 1979) in Haplognathia rosea (as H. cf. rosacea) and H. gubbarnorum (as H. cf. lyra). H. rosea has paired sperm duct glands but no specialized penis muscles, a penis with only one cell type and without a detectable lumen.

H. gubbarnorum, somewhat more complex, has an H-shaped sperm duct gland, three groups of penis muscles, and a penis with two cell types and a lumen (Fig. 2.26). There is no open gonopore but a tissue connection between the penis and the ventral epidermis, surrounded by a rosette of gland cells (see Sterrer 1969: figures 54, 55, and 291), through which the sperm is assumed to exit during copulation. The poorly known copulatory organ of Conophoralia resembles, at least superficially, that of Haplognathia (Mainitz 1979). In LM, the organ seems tripartite (Fig. 2.40 E), with a proximal part of medium grainy appearance, a

2.2 Morphology 

 175

Fig. 2.40: Scleroperalian female (A, B) and male (C, D) reproductive organs. (A) Bursa of Valvognathia pogonostoma from Denmark. (B) Bursa of Gnathostomula paradoxa from Denmark. (C) Penis apparatus of Valvognathia pogonostoma from Denmark. (D) Penis apparatus of Gnathostomula paradoxa from Denmark. Abbreviations: bu, bursa; ds, mature dwarf sperm; pb, prebursa; ps, penis stylet; sg, stylet gland. (LM micrographs of live specimens.)

middle part coarsely grained, and the distal part finely grained and appearing as a concentrically structured, sometimes twisted funnel (Sterrer 1997, 2001). Proximally, the organ often contains at least one mature sperm. The scleroperalian male copulatory organ (Figs. 2.39 A, B, 2.40 C, D, and 2.41) is complex but rather uniform (Mainitz 1977, 1979, 1988). An invaginated extension of the body wall, the conical penis consists of a proximal stylet sac and a stylet apparatus. The latter is composed of only two cell layers, epithelial and muscular. The core of the penis is a lumen-less “stylet” consisting of 8 (in Labidognathia longicollis) or 10 (in Gnathostomula and

Semaeognathia sterreri) radially arranged, microtubuleand crystal-filled rod-like cells whose bodies are located at the proximal end of the stylet. In Gnathostomulidae (but not in Labidognathia), the entire stylet is surrounded by a tube-like, cuticular stylet sheath (Fig. 2.41). A monolayered but massive stylet gland envelops the entire structure. Distally, the stylet points to the male pore in whose vicinity there are ciliary receptors and gland cells. It is assumed that these play a role in copulation when the stylet apparatus is projected to inject sperm into the partner. The observation in Gnathostomula of stylet-like structures in newly filled bursae as well as of empty stylet

176 

 2 Phylum Gnathostomulida

Fig. 2.41: Scleroperalian penis stylet apparatuses. (A) Labidognathia longicollis. (B) Gnathostomula sp. Abbreviations: ac, atrial cells; bl, basal lamina; ec, epithelial cell; ir, inner rod; ml, muscular layer; mo, male opening; or, outer rod; po.ss, proximal opening of stylet sack; pss, proximal stylet sack; rc, rod cell: sg, stylet gland; ss, stylet sheath. (Schematic reconstruction after Mainitz 1979.)

sheaths (Mainitz 1979) suggests that the loss of the stylet may be a regular occurrence during copulation. Two scleroperalians, Gnathostomaria lutheri and Rastrognathia macrostoma, differ from this pattern mostly by having a much longer (100–125 µm) stylet (Sterrer 1998, Kristensen & Nørrevang 1977).

droplets and glycogen particles. There is no exocytosis for the formation of a non-sclerotized eggshell (Falleni 1993). The mature egg of Gnathostomula jenneri shows a cytoplasm packed with evenly distributed dense yolk granules, elongate mitochondria with only a few cristae, ribosomes, and cytoplasmic cisternae. It appears that fertilization occurs after meiotic division (Mainitz 1983).

2.2.10 Gametes

2.2.10.2 Sperm and spermiogenesis (Figs. 2.39 E and 2.42–2.46)

2.2.10.1 Eggs and oogenesis As studied by Falleni (1993) in Austrognathia cf. riedli, oogenesis is not synchronous but progresses posteriorly and has a previtellogenic and a vitellogenic phase. Throughout oogenesis, each oocyte is completely enveloped by the long cytoplasmic processes of flat accessory cells. The previtellogenic phase is characterized by a high nuclear/cytoplasmic ratio and the presence of chromatoid bodies surrounded by mitochondria. Vitellogenic oocytes are characterized by the increase of RER, numerous Golgi complexes, and the accumulation of small (1–2.5 µm) electron-dense yolk globules. Yolk synthesis proceeds both by an endogenous mechanism involving RER and Golgi activity and an exogenous uptake of yolk precursors by endocytosis. Yolk consists of proteins and a small amount of glycoproteins, but mature oocytes also contain lipid

Three types of sperm (Fig. 2.42) can be distinguished: filiform (in Filospermoidea), conulus (in Conophoralia), and dwarf (in Scleroperalia) (Sterrer et al. 1985). Up to 100 µm long, filiform sperm (Figs. 2.42 A, B and 2.44 A) consists of head, middle piece, and a single tail (Sterrer 1966, 1969, Sterrer et  al. 1985, Lammert 1991). Tipped by an acrosomal vesicle, the head is made up of an elongated nucleus that winds, corkscrew-like, around a clear central axis. The middle piece has an envelope of mitochondrial derivatives around the single flagellar axoneme of a 9+2 pattern, which continues into the tail. All Filospermoidea have this type of sperm; in LM, species differ in sperm length and proportions as well as the prominence of head spiralization. Filiform sperm move actively by slow flagellar writhing as well as corkscrewlike drilling.

2.2 Morphology 

All Conophoralia have cone- or mushroom-shaped sperm of the conulus type (Sterrer et  al. 1985). The size and shape of conuli (Figs. 2.42 H, 2.44 E, F, and 2.46) are species-specific, ranging in length from 6 µm (Austrognatharia barbadensis) to 75 µm (Austrognathia sp., Sterrer, unpublished data), and with shapes ranging from squat to button-mushroom- to spindle-like. In LM, conuli have been described as consisting of a cone-shaped body (stalk) topped, at its blunt end, by a capitulum (hat) much like half a car tire, bordered by a frilly cingulum (fringe), all of it embedded in a “matrix” (Sterrer 1972). As analyzed with TEM in Austrognathia sp. by Lanfranchi & Falleni (1998), the capitulum and body are made up by the nucleus, in three degrees of chromatin condensation (Fig. 2.44 F): high and medium (in the capitulum) and low (in the conulus body). The capitulum is capped by a thin cytoplasmic layer with paracristalline palisades, suggesting an acrosomal function. The electron-lucent nuclear region of the sperm body is arranged as concentric lamellae around two longitudinal axes; it is at the transition between capitulum and body that (at least) two centriole-like structures (but no axonemes) can be found, each consisting of 9 solid, single microtubules. The cytoplasm of mature sperm is characterized by an abundant, narrowly tubular labyrinth and bulges at the intersection of capitulum and body where it forms the frilly cingulum rich in mitochondria. The “matrix” of earlier descriptions corresponds to the cytoplasm of sperm (and atrophied trophocyte). Conulus sperm do not actively move. Dwarf sperm (Figs. 2.42 C–G, 2.44 B–D, and 2.45 A–B) is typically 2–3 µm long, round or droplet-shaped, and

 177

lacks a flagellum (Graebner 1968b, 1969a, b, Riedl 1969, Graebner & Adam 1970, Alvestad-Graebner & Adam 1983). It is further distinguished by a basally located nucleus, 1–2 mitochondria, a single dense layer of microtubules immediately under the plasma membrane and “micropodia”, i.e., tubular protuberances of the plasma membrane, 0.18 to 1 µm long. Micropodia are arranged in clusters along one side (Gnathostomula paradoxa), or fringe one end of the elongated sperm (G. mediterranea), or bunch out of a spherical sperm (G. jenneri). Micropodia might either aid sperm motility or have an acrosomal function (AlvestadGraebner & Adam 1983). All Scleroperalia sperm follow this general pattern, with the exception of Gnathostomaria (Sterrer et  al. 2001) whose sperm (“mega dwarf”, Fig. 2.42 C and 2.44 C) are much larger, up to 18 µm, consisting of a spherical body 8 µm in diameter and a dense tuft of microvilli-like micropodia up to 10 µm long. A rudimentary flagellum has been identified above the nucleus in Gnathostomaria sperm (Sterrer et al. 2001, figure 1), but not in typical dwarf sperm. Squeezed out of their follicles, dwarf sperm (Fig. 2.45) may show directionless amoeboid behavior akin to Brownian motion (Alvestad-Graebner & Adam 1983). 2.2.10.2.1 Spermiogenesis On the basis of LM observations in Filospermoidea, Sterrer (1969) failed to note follicles nor an origin or direction of spermatogenesis within the testes; the latter simply narrow posteriorly into sperm ducts filled with mature sperm. In spermatogonia, the nucleus is very large, with a loosely granular chromatin structure. Meiosis apparently proceeds without cytokinesis, resulting in four spermatid

Fig. 2.42: Sperm cells of various gnathostomulid species. (A) Filiform sperm of Pterognathia swedmarki. (B) Filiform sperm of Haplognathia simplex. (C) Giant sperm of Gnathostomaria lutheri. (D) Dwarf sperm of Onychognathia filifera. (E) Dwarf sperm of Gnathostomula microstyla. (F) Dwarf sperm of Agnathiella beckeri. (G) Dwarf sperm of Gnathostomula jenneri. (H) Conulus of Austrognathia riedli. Abbreviations: c.bo, conulus body; c.ci, conulus cingulum; c.ha, conulus hat; c.ma, conulus matrix. (Line art reconstructions from LM observations, after Sterrer 1974.)

178 

 2 Phylum Gnathostomulida

Fig. 2.43: Spermiogenesis of Haplognathia simplex. (A) Section through trunk, showing paired testes, marked with double black and white dashed lines. (B) Section through spermatocyte, showing early stage of spermiogenesis with uncondensed nuclei of four spermatids. (C) Spermatocyte with developing spermatid inside. (D) Same spermatid as in C with developing acrosome next to the Golgi apparatus. (E) Section through spermatocyte, showing later stage of spermiogenesis with condensed nuclei of four spermatids. (F) Section through testis, showing longitudinal sections through mature spermatozoa; note the characteristic corkscrew shape. Abbreviations: ac, developing acrosome; cn.spi, condensed nucleus of spermatids; ep, epithelial cell; f.spa, flagella of spermatozoa; f.spi, flagella of spermatids; ga, Golgi apparatus; m.spi, membrane of spermatid; m.spy, membrane of spermatocyte; mi, mitochondria; mu, muscle; n.spa, nuclei of spermatozoa; n.spy, nucleus of spermatocyte; un.spi, uncondensed nuclei of spermatids. (TEM sections courtesy of Reinhardt M. Kristensen.)

2.2 Morphology 

 179

Fig. 2.44: Gnathostomulid sperm cells. (A) Filiform sperm of Haplognathia rosea, scale not available. (B) Dwarf sperm of Gnathostomula sp. (C) “mega” dwarf sperm of Gnathostomaria lutheri. (D) Dwarf sperm of Gnathostomula cf. mediterranea. (E) Conulus of Austrognathia sp. from Hong Kong. (F) Conulus of Austrognathia sp. from the Mediterranean. Abbreviations: ds, mature dwarf sperm; mi, mitochondrion; mp, micropodia; n, nucleus. (TEM sections courtesy of Reinhard M. Rieger and Gunde Rieger, A, B, and Johannes G. Achatz, C, D, from Graebner 1969b.)

180 

 2 Phylum Gnathostomulida which may have two nuclei, eventually degenerates and the periphery of the spermatid cytoplasm deflates to a tubular labyrinth. Spermiogenesis in Scleroperalia takes place in follicles surrounded by a theca (Alvestad-Graebner & Adam 1983) and proceeds from the periphery toward the follicle lumen. Spermatogonia have a highly active Golgi apparatus, two centrioles, a nucleus with irregularly dispersed chromatin, and a prominent nucleolus. Spermatocytes and spermatids develop in tetrads. In spermatocytes, the Golgi vesicles surround the nucleus in a single array; in spermatids, they come to lie in regularly spaced “outpocketings” of the plasma membrane, which may eventually become the “micropodia” typical of dwarf sperm.

2.3 Reproduction and development 2.3.1 Reproductive biology Fig. 2.45: Bursovaginoid sperm cells. (A) Dwarf sperm of Agnathiella beckeri from South Africa. (B) Dwarf sperm of Vampyrognathia varanus from the Red Sea. (LM micrographs of live specimens.)

nuclei within a single cell (Knauss & Rieger 1979). In filospermoid (Figs. 2.43) spermatids, which are bunched in tetrads, the nucleus condenses then stretches to a spindle, at which time the axoneme appears. The spindle finally coils spirally around a translucent axis. Several mitochondria merge to a single derivative, which eventually surrounds the centriole of the growing axoneme, to form the middle piece of the sperm (Sterrer et al. 1985, Lammert 1991, figure 17a, b). The axoneme is usually much longer in spermatids than in mature sperm. In Conophoralia, spermatogonia hug the inside of the thick basement membrane enclosing the saccular testis (Lanfranchi & Falleni 1998). Production of primary and secondary spermatocytes proceeds toward the testis lumen, where spermatids and sperm eventually congregate. From the secondary spermatocytes onward, a nuclear envelope is no longer visible. Spermatids develop synchronously in tetrads. The spermatid nucleus, initially ovoid, lengthens into the characteristic conulus shape, then differentiates into the three chromatin condensations of capitulum and stalk. Each spermatid is tightly surrounded by a large nurse cell (trophocyte) that provides, via wide cytoplasmic bridges, mitochondria, and nutrients to the swollen tubular labyrinth in the periphery of the spermatid cytoplasm. The nurse cell,

All gnathostomulids are simultaneous hermaphrodites, albeit with maturation of male sometimes preceding that of female gonads. As mating has never been observed directly, we can only postulate mechanisms on the basis of anatomy (Sterrer 1974). Simultaneous cross-fertilization seems possible in Filospermoidea but is unlikely in other taxa. In Filospermoidea, the glandular male opening (see Sterrer 1969, figures 54–55) is pressed against, and likely adheres to, a partner’s epidermis into which the highly mobile sperm actively drill their way, ending up between (or inside?) gut cells throughout the body (see Sterrer 1969, figures 56 and 61); some are even found in the rostrum. It is here that they seem to linger, singly or in bundles, prior to fertilizing an egg. The partner’s sperm look identical to those found in vasa deferentia. In Conophoralia, the muscular(?) penis inserts one, rarely two, conulus into the dorsal vagina. On arrival in the soft bursa, the sperm assumes its typical “bursa conulus” aspect: with the capitulum intact but the sperm body becoming granular, hollow, and open at the tip (Sterrer 1974). In Gnathostomula, and probably all Scleroperalia, the processes following copulation seem still more complex (Riedl 1971b, Müller & Ax 1971). A mature testis follicle is injected, by the penis stylet, under the partner’s skin (or into the vagina where present) in the middorsal body region. The injected follicle turns into the first prebursa and subsequently into the bursa by differentiating typical layers and a mouthpiece. If there



2.3 Reproduction and development 

 181

Fig. 2.46: Conophoralian giant sperm cells (conuli). (A) Austrognathia riedli forma marisrubri from the Red Sea, scale not available. (B) Austrognathia sp. from Florida. (C) Austrognathia sp. from Réunion Island. (D) Austrognatharia sterreri from Belize. (F) Austrognatharia boadeni from Roscoff, France. Abbreviations: c.bo, conulus body; c.ha, conulus hat; c.ma, conulus matrix. (LM micrographs of live specimens.)

is already a bursa from a previous copulation, the prebursa joins and refills the bursa (Riedl 1971b). Sperm are tightly packed in the bursa, from where they presumably reach the mature egg via the nozzle-like bursa mouthpiece.

2.3.2 Oviposition and cleavage Riedl (1969, figures 5 and 6) provided the only observations of oviposition and cleavage, in Gnathostomula jenneri. The animal adheres with its dorsal side to the substrate, pressing the mature egg caudally, between gut and bursa, until the body wall behind the bursa is penetrated. Oviposition is suspected to result in seasonal body fragmentation at least in Gnathostomaria lutheri: all of the 259 specimens collected at the type locality in October 1966 were ante-

rior fragments lacking mature egg, bursa and male organs (Sterrer 1971a), whereas all of the 8 specimens found at the same locality on July 2, 1995, were intact and sexually mature (Sterrer et al. 2001). The emerging zygote in Gnathostomula jenneri (Riedl 1969) becomes spherical, ~55 µm in diameter and sticks slightly to the substrate. Cleavage begins a few hours after oviposition, with the first and second divisions meridional (from pole to pole), nearly equal, and holoblastic. From the third division onward, eggs show a spiral plan, with alternation of dexiotropic and leiotropic planes of cleavage. The first equatorial division is unequal, with clockwise displacement of the first micromere quartet. After 20 hours, further cleavages result in a mass of micromeres covering the animal hemisphere. “Tip” cells of the micromere cross appear on the third day, and on the fourth day, epibolic gastrulation is completed.

182 

 2 Phylum Gnathostomulida

Fig. 2.47: Hatching of a Haplognathia ruberrima. (A) Egg with mature embryo moving inside. (B) Juvenile hatching from egg. (LM micrographs of live specimens, from Sterrer 1974.)

2.3.3 Development (Fig. 2.47) Development is direct. The hatching juvenile of Gnathostomula jenneri is 100 µm long, completely monociliated, and has about the volume of the egg (Riedl 1969). The anterior sensorium and the jaws are missing, but there is a rudiment of the pharynx, including the central (comb) part of the basal plate. Sterrer (1998, figure 14) added observations on Haplognathia ruberrima: in a colorless, ovoid egg capsule, 170 × 120 µm, a juvenile was slowly rotating by means of its ciliary beat until it pierced the capsule with the tip of the rostrum. The hatchling, 330 µm long and 60 µm wide, had a doughnut-shaped pharynx with a fully formed basal plate complete with 9–10 rows of thorns, whereas of the jaws only the anterior-most contour could be discerned. Pharyngeal hard structures are produced in situ and do not change in size or shape during the animal’s life span. Male organs, especially the penis, always appear before female organs, with the stylet being produced from the distal end. In Gnathostomula paradoxa, which grows to 1000 µm length, the genital organs are formed at 400–600 µm length, with the male stylet first, followed by bursa, and finally, the male and female gonads (Müller & Ax 1971). This round completed, the entire genital apparatus may be reduced and replaced by a new set. G. paradoxa mainly reproduces in spring (Müller & Ax 1971). In Austrognathia, the unpaired testis

develops from paired lateral anlagen that then join dorsally of the gut (Sterrer 1971).

2.4 Biology and physiology (Figs. 2.48 and 2.49) Locomotion is a slow gliding by means of the ciliary beat, with occasional body contraction or spiraling when meeting an obstacle and adhesion to the substrate, usually with the tail. The cilia of the rear end seem oriented and are beating, in reverse, seemingly keeping the animal anchored while it is exploring new territory (Sterrer 1969: figures 129, 133, and 134). Gregariousness, observed in Gnathostomula jenneri (Riedl 1969), may facilitate the meeting of potential mating partners. In some Haplognathia species, the jaws and basal plate, particularly of mature specimens, show a “grainy degeneration” (Sterrer 1969), possibly a deposition of excretory products (Fig. 2.48). Respiration rates were measured in two species in Bermuda (Haplognathia cf. ruberrima, with 413  mm3 O2/ hour/g wet weight, and Gnathostomula sp., with 282 mm3 O2/hour/g wet weight), at 20°C (Schiemer 1973). These values are at the lower end of the range for meiobenthic animals with a body size of 1–10 µg wet weight and

2.5 Ecology 

­consistent with life in a strongly reducing environment. Powell et  al. (1979) propose that members of the “thiobios” have a sulfide detoxification system in the body wall, which traps and disarms much of the sulfide that enters it. Rieger et al. (1986) suggest that modified mitochondria, common in thiobios, might be correlated with the occurrence of anoxic biochemical pathways using part of a reversed Krebs cycle as the main source of energy production. A viable egg of Haplognathia ruberrima found in a black, sulfidic sand layer (Sterrer 1974, 1998) suggests that even early development takes place in anoxybiotic conditions. Two species of Austrognatharia have been observed producing a mucous cyst (Fig. 2.49), possibly in response to a deteriorating environment (Sterrer 1971a, 1998).

2.5 Ecology Gnathostomulida are free-living and almost exclusively marine; only one species, Gnathostomaria lutheri, was found in brackish water: in a coastal pond in southern France, with 1.9% to 2.3% salinity (Ax 1956, 1964), and a supratidal pool at the intracoastal waterway in North Carolina, USA (Sterrer 1998). The majority of species have been recorded in shallow water, from the supratidal to 30 m; only two (G. uncinata and Triplignathia(?) bathycola) have been reported from 400 m off North Carolina, USA (Sterrer 1998), to date the deepest record for the phylum. The

 183

typical substratum is a poorly sorted, fine to medium, even coarse sand of organic (as in the tropics) or inorganic origin (as along temperate shores), invariably containing a significant admixture of marine (but not terrigenic) organic detritus, as found in sheltered bays and inlets, coastal ponds, between coral reefs and near mangroves and sea grasses. Between 1974 and 2004, more than 100 sand samples collected between coral reefs and mangroves around CarrieBow Cay (Belize) yielded 25 gnathostomulid species, the largest number from anywhere in the world (Sterrer 2004, figure 1). Gnathostomaria lutheri lives in littoral sand under piled-up leaves of Zostera (Ax 1964, figure 5), and Gnathostomula axi was first described from a decaying layer of dead Thalassia leaves in a shallow mangrove channel (Kirsteuer 1964). Species of Gnathostomula, in particular, can be rather eurytopic; G. paradoxa strays to mud (with Zostera, Karling 1962) as well as to clean sand (Ax 1956) and shell gravel (Ax 1965). Seven gnathostomulid species were found in small pockets of sediment trapped by the rhizomes of Phyllospadix sp., a seagrass growing on high-energy rocky coasts on the west coast of North America (Farris & O’Leary 1985, Sterrer & Sørensen 2006). Gnathostomulids have also been extracted from stromatolitoid microbial nodules (Westphalen 1993) and are numerous alongside permanent tubes of the polychaetes Arenicola marina and Nereis virens (Reise 1981). A meiofaunal assemblage dominated by Gnathostomulida was found beneath a mat of sulfur bacteria close to a brine seep at 72 m in the Gulf of Mexico (Powell

Fig. 2.48: Stages of jaw and basal plate grainy degeneration in Haplognathia simplex. (A) Specimen with regular jaws and basal plate. (B) Partially degenerated jaws and basal plate. (C) Strongly degenerated jaws and basal plate. Abbreviations: bp, basal plate; ja, jaws. (LM micrographs of live specimens, after Sterrer 1969.)

184 

 2 Phylum Gnathostomulida low oxygen concentrations, grazing on chemoautotrophic sulfur-oxidizing bacteria. This suggests an ancient origin for the phylum, having originated in (or at least survived) oceans depleted in oxygen (Pascal et al. 2014).

2.6 Phylogeny

Fig. 2.49: Austrognatharia boadeni inside mucous cyst. (LM micrograph of live specimen.)

et al. 1983). Reise (1981) proposes that the high degree of spatial overlap between most species indicates a narrow ecological niche for the entire taxon, possibly defined by the presence of sulfur bacteria. Pascal et al. (2014) found Haplognathia ruberrima on white mats of filamentous sulfur bacteria (Beggiatoa spp.); based on stable isotope analysis, they conclude that, lacking symbiotic bacteria, H. ruberrima grazes selectively on chemoautotrophic sulfur-oxidizing bacteria. It was the discovery of a diverse gnathostomulid fauna in subsurface marine sands that prompted Fenchel & Riedl (1970) to propose the existence of “the sulfide system: a new biotic community underneath the oxidized layer of marine sand bottoms”. This globally vast biome is characterized by high sulfide concentrations and very low to zero oxygen levels, conditions that had been considered inimical to metazoan life. This black sediment layer is separated from the overlying sediment by a redox potential discontinuity (RPD), where oxidizing processes become displaced by reducing processes. Indeed, studies of the vertical distribution in the sediment show that Gnathostomulida and other members of this “thiobios” are invariably most diverse (and often reach high specimen numbers) at or below the RPD layer (Powell et  al. 1983). We may conclude that Gnathostomulida typically live in marine interstitial environments with high sulfide but

The first attempt to understand gnathostomulid phylogeny was by Sterrer (1972) at a time when the major lineages and most of the gnathostomulid biodiversity at generic level had only recently been discovered. Sterrer (1972) proposed the division of Gnathostomulida into the major clades, Filospermoidea and Bursovaginoidea, of which the latter represents the greatest biodiversity and apparently reaches the highest degree of morphological complexity. Filospermoidea consists of the genera Haplognathia, Cosmognathia, and Pterognathia, with the two latter being putative sister groups. The remaining gnathostomulid taxa known by then were assigned to the other order, Bursovaginoidea. Based on differences in the reproductive system and gametes, Sterrer (1972) proposed a subdivision of Bursovaginoidea into the suborders Scleroperalia and Conophoralia. The latter is identical with the family Austrognathiidae, which, among other traits, are characterized by producing the giant male gametes known as conuli. Austrognathiidae (and hence Conophoralia) were considered sister group to the scleroperalian family Gnathostomulidae, which, in other words, left the second suborder, Scleroperalia, paraphyletic. The remaining scleroperalian taxa were assigned to the four families Agnathiellidae, Mesognathariidae, Gnathostomariidae, and Onychognathiidae. The phylogenetic system proposed by Sterrer (1972) was first tested in a formal cladistic analysis by Sørensen (2002) (Fig. 2.50). The analyses were based on morphological information from 25 terminals, representing the genera that were known by that time. The analyses confirmed most of the hypotheses proposed by Sterrer (1972), leaving only a question mark at the exact interrelationships between the scleroperalian families. Most recently, gnathostomulid phylogeny was analyzed using a combined approach of morphological information and molecular sequence data (Sørensen et  al. 2006) (Fig. 2.51). The analyses recognized the monophyly of the two orders Filospermoidea and Bursovaginoidea, but within the latter clade, the position of Conophoralia was changed: from being sister group to Gnathostomulidae to a position most basally in Bursovaginoidea, hence, for the first time supporting scleroperalian monophyly. The analyses furthermore supported monophyly for the family Mesognathariidae, whereas it was unable to resolve

2.6 Phylogeny 

 185

Fig. 2.50: Strict consensus tree of four most parsimonious trees, based on morphological analysis of Sørensen (2003).

the relationships between the taxa of Onychognathiidae and Gnathostomulidae unambiguously. Terminals representing Agnathiellidae and most of the monotypic families were not included in the analysis because molecular data were unavailable for these species. At the present state, we can consider gnathostomulid monophyly, as well as the division into Filospermoidea and Bursovaginoidea, as well supported by both molecular and morphological data. The filospermoid species are all characterized by having a relatively high body index

(body length/maximum body width ratio), high rostrum index (rostrum length/maximum rostrum width ratio), simple filiform spermatozoa, hypodermic insemination, presence of jaws with a somewhat expanded, anchoring symphysis, and a dentarium with acicular teeth. Furthermore, they are all lacking compound ciliary sensoria, an injectory penis, and a bursal system. Several of these traits can most certainly be considered symplesiomorphic, e.g., the filiform sperm and absence of sensoria and bursa, whereas it is difficult to establish the polarity of

186 

 2 Phylum Gnathostomulida

Fig. 2.51: Shortest tree from analyses of Sørensen et al. (2006) based on data from four molecular loci (18S rRNA, 28S rRNA, histone H3, and cytochrome c oxidase subunit I) and a morphological matrix. The tree only includes taxa from which molecular sequence data were available; hence, not all genera are represented.

some of the other characters. For instance, it is not possible to clarify with certainty if the extremely elongated body and rostrum represent filospermoid autapomorphies or whether they are parts of the gnathostomulid ground pattern. A potential filospermoid autapomorphy could be the hypodermic insemination that apparently happens by sperm actively drilling through the integument of the mate. Even though similar insemination strategies have developed in other Metazoa, it is known from neither the bursovaginoid taxa nor the Rotifera. Also, the expanded symphysis that unites the two jaw halves could represent a potential filospermoid autapomorphy. As the anchoring function of the symphyses has been replaced by the suspensorial caudae in many bursovaginoid species, their symphyses are rarely well developed. In the main jaws of Limnognathia maerski, the symphysis is relatively narrow

as well, and in species of Rotifera, the fulcrum, which is considered the counterpart to the gnathostomulid symphysis (Sørensen 2000), is usually laterally compressed. Bursovaginoid monophyly is supported by several autapomorphies, including the presence of ciliary sensoria, a bursal system, pharynx with tri-lobed musculature, and paired pharyngeal glands. It is furthermore likely that the five-partite basal plate evolved at the bursovaginoid stem, but because no structure homologous to the gnathostomulid basal plate is present in species of Rotifera or Micrognathozoa, this character cannot be optimized unambiguously. Also, the ground pattern of the bursovaginoid jaws is difficult to establish. Sterrer (1972) originally proposed that the open lamellar jaw evolved at the base of Bursovaginoidea, and this viewpoint was subsequently (although not unambiguously) supported by the

2.7 Systematics 

analysis of Sørensen (2002). However, the establishment of Austrognathiidae as the most basal bursovaginoid family (see Sørensen et al. 2006) opens the possibility of the fused lamellar jaw type as being part of the bursovaginoid ground pattern. The division of Bursovaginoidea into the suborders Conophoralia and Scleroperalia is strongly supported by molecular sequence data (Sørensen et  al. 2006). Again, difficulties with unambiguous character polarization ­ make it problematic to point out clear-cut autapomorphies for the two clades. It is, however, reasonable to assume that the simple filiform sperm type was present in the bursovaginoid ground pattern and evolved into giant conuli at the conophoralian branch and into modified dwarf sperm at the branch leading to Scleroperalia. Likewise, we find it likely that the bursa evolved as a soft structure at the base of Bursovaginoidea and subsequently was hardened and became sclerotized at the base of Scleroperalia. This evolution could, eventually, be correlated with the formation of a sclerotized penis stylet. Within Scleroperalia, the relationships remain puzzling. Morphological data support monophyly of the families Agnathiellidae, Mesognathariidae, Onychognathiidae, and Gnathostomulidae (Sørensen 2002), and even though analyses based on combined morphological and molecular evidence do not find support for the two latter (Sørensen et  al. 2006), we regard this as an artifact due to insufficient taxon sampling among the relevant taxa. The relationships between the bursovaginoid families (of which four remain monotypic) also remain unresolved, and we expect that improved molecular taxon sampling and examination of pharyngeal hard parts in species of more poorly known genera will be required to solve these last questions. The ancestral form of Gnathostomulida emerges as a slender acoelomate worm with an equally slender, undelimited rostrum, devoid of compound sensory cirri but equipped with 1. a completely monociliated epidermis bearing singlecell glands and ciliary receptors; 2. an entirely intraepithelial nervous system consisting of a frontal and a buccal ganglion, longitudinal nerves, and a commissure each in the regions of the male organ and tail; 3. body-wall musculature situated between the epidermal basal lamina and the gut epithelium, with weak circular fibers outside of paired, stronger longitudinal bundles; 4. paired three-cell excretory organs; 5. ventral, subterminal mouth; intestine without or with a temporary anus (tissue connection);

 187

6. pharynx as a simple muscular network anchored to the body wall, set with cuticular, rod-enforced plaques in the form of a ventral, four-ridged basal plate and a pair of solid, ventrolateral jaws. The dorsal surface of the basal plate and the anterior jaw may be set with acicular teeth. Hermaphrodites; ovary antero-dorsal, unpaired, with eggs maturing posteriorly. Without a bursa or vagina. Testes paired, situated dorsolaterally, tapering into vasa deferentia, which end subterminally in a simple, glandular male pore. Sperm motile, with one flagellum with 9 + 2 microtubular arrangement. Copulation by adhesion of the male pore (tissue connection) to the partner, which the sperm actively penetrates prior to seeking the mature egg. Oviposition by rupture of the body wall.

2.7 Systematics 2.7.1 Phylum Gnathostomulida (Ax, 1956) Riedl, 1969 Small free-living, worm-shaped acoelomate Bilateria, with a muscular pharynx usually provided with paired jaws and an unpaired cuticular basal plate. Without an anus. Epidermis monociliated. Parenchyma poorly developed. Hermaphrodites. Distribution: Marine.

2.7.1.1 Order Filospermoidea Sterrer, 1972 Gnathostomulida with filiform sperm and without a bursa and vagina. Male opening without an injectory penis. Without paired sensory cirri on the rostrum, but with occipitalia. Body usually very elongated (body index at least 25); rostrum undelimited from trunk, pointed, and slender (index usually more than 3) and not delimited. Pharynx musculature rather loose; jaws more or less compact, with wing-shaped apophyses and a solid symphysis that is usually wider than long. 2.7.1.1.1 Family Haplognathiidae, Sterrer 1972 Filospermoidea with a short pharyngeal bulb (index usually less than 8.0). Basal plate longer, or equally long as wide, or only slightly wider than long (index more than 0.5), often with teeth or ridges on its dorsal surface. Jaws compact, without a crista and with no or few teeth arranged in a ventrolateral arc. One described genus (Haplognathia).

188 

 2 Phylum Gnathostomulida

Genus Haplognathia Sterrer, 1970a (Figs. 2.2 A, 2.3 A–D, G–I, 2.20 A, 2.21 A–D, and 2.22A–I) Haplognathiidae with simple, not horizontally bipartite jaws without or with very few teeth (usually no more than 5). Testes paired. Type species: H. ruberrima (Sterrer, 1969). Ten described species: H. simplex (Sterrer, 1966a); H. filum (Sterrer, 1966a); H. ruberrima (Sterrer, 1966a); H. rubromaculata (Sterrer, 1969); H. rosea (Sterrer, 1969); H. gubbarnorum (Sterrer, 1969); H. lunulifera (Sterrer, 1969); H. lyra Sterrer, 1970a; H. belizensis Sterrer, 1998; H. asymmetrica Sterrer, 1991b. 2.7.1.1.2 Family Pterognathiidae Sterrer, 1972 Filospermoidea with a long pharyngeal bulb (index usually more than 8.0). Basal plate much wider than long (index less than 0.5), often with teeth at the frontal edge. Jaws often slightly lamellar, with a crista, and with several to many teeth arranged in a ventrolateral arc or a ventrorostrodorsal basket. Two described genera (Pterognathia and Cosmognathia). Genus Pterognathia Sterrer, 1966a (Figs. 2.3 J and 2.23 E–I) Pterognathiidae with more or less lamellar, horizontally bipartite jaws with a crista, with many teeth (usually more than 5). Testis usually unpaired and situated dorsally. Type species: P. swedmarki Sterrer, 1966a. Fourteen described species: P. swedmarki Sterrer, 1966; P. meixneri Sterrer, 1969; P. atrox Sterrer, 1969; P. sorex Sterrer, 1969; P. ctenifera Sterrer, 1970a; P. crocodilus Sterrer, 1991a; P. vilii Sterrer, 1991a; P. hawaiiensis Sterrer, 1991b; P. ugera Sterrer, 1991c; P. alcicornis Sterrer, 1998; P. pygmaea Sterrer, 1998; P. sica Sterrer, 2001; P. portobello Sterrer, 2006; P. tuatara Sterrer, 2006. About two undescribed species (Sterrer, unpublished data). Genus Cosmognathia Sterrer, 1991a (Figs. 2.3 E, F, 2.20 C, D, 2.21 E, F and 2.23 A, D) Pterognathiidae with few to many, usually very delicate teeth, arranged in two groups but not equally divided between dorsal and ventral part of jaw nor forming an arc or basket. Rostral edge of basal plate without or with very delicate teeth or ridges; caudal edge usually convex and enclosing a spherical knob. Type species: C. arcus Sterrer, 1991a. Four described species: C. arcus Sterrer, 1991a; C. bastillae Sterrer, 1991a; C. manubrium Sterrer, 1991b; C. aquila Sterrer, 1998.

2.7.1.2 Suborder Conophoralia Sterrer, 1972 Bursovaginoidea with a soft bursa and permanent or temporary vagina. Penis, muscular, without a cuticular stylet. Testis usually unpaired, situated dorsally. Sperm large (from 6 µm to 75 µm long), conical (“conuli”); without flagellum or filaments. Sensorium consisting of a maximum 4 pairs of compound cirri and 2 pairs of apicalia. 2.7.1.2.1 Family Austrognathiidae Sterrer, 1971 Conophoralia with 2 pairs of apicalia and 4 pairs of long (40–55 µm) compound cirri, of which the ventralia originate close to the rostralia. Ciliary pits present. With a pair of pre-pharyngeal glands. Epidermal cells in stripes of 2–3, epidermal inclusions usually in groups. Testis unpaired, situated dorsally. Usually with a vagina. Basal plate winged, with a toothed central part and pronounced lateral, but small rostral wings. Jaws lamellar and closed, with up to 3 horizontal rows of teeth. Without a jugum. Without ability to swim backward. Three described genera (Austrognathia, Austrognatharia, Triplignathia). Although uniquely identified by the possession of conuli (which also provide the most species-distinguishing character set), Austrognathiidae have been allotted to genera somewhat arbitrarily, given the continuous range of variation in jaw and basal plate structure (Sterrer 1991d). Genus Austrognathia Sterrer, 1965 (Figs. 2.2 C, 2.5 A, B, 2.6 H, 2.32 C, D, 2.34 C–E, and 2.35 A, C, D, G, H) Slender to fairly plump Austrognathiidae with two rows of jaw teeth. Basal plate with a median lobe; basal plate teeth more or less equal in diameter. Type species: A. riedli Sterrer, 1965. Eleven described species: A. riedli Sterrer, 1965; A. marisrubri (Riedl, 1966); A. hymanae Kirsteuer, 1970; A. christianae Farris, 1977; A. microconulifera Farris, 1977; A. singatokae Sterrer, 1991a; A. nannulifera Sterrer, 1991a; A. novaezelandiae Sterrer, 1991a; A. macroconifera Sterrer, 1991c; A. clavigera Sterrer, 1997; A. australiensis Sterrer, 2001; About 8 undescribed species (Sterrer, unpublished data). Genus Austrognatharia Sterrer, 1971a (Figs. 2.32 E, F, 2.34 A, B, and 2.35 B, E, F) Fairly slender to plump Austrognathiidae with only the ventral row of jaw teeth well developed; dorsal row with 1 tooth to 2 teeth. Basal plate without a median lobe; basal plate teeth rather unequal in diameter. Type species: A. boadeni Sterrer, 1971b.

2.7 Systematics 

Twelve described species: A. boadeni Sterrer, 1971b, A. sterreri (Kirsteuer, 1969a), A. kirsteueri Sterrer, 1970a; A. strunki Farris, 1973; A. atraclava Ehlers & Ehlers, 1973; A. homunculus Sterrer, 1991a; A. pecten Sterrer, 1991a; A. mooreensis Sterrer, 1991c; A. medusifera Sterrer, 1998; A. stirialis Sterrer, 1998; A. australis Sterrer, 2006; A. barbadensis Sterrer, 2011. About 6 undescribed species (Sterrer, unpublished data). Genus Triplignathia Sterrer, 1991 Plump Austrognathiidae with three rows of jaw teeth. Basal plate with a median lobe; basal plate teeth robust and more or less equal in diameter. Type species: T. adriatica Sterrer, 1991. Two described species: T. adriatica Sterrer, 1991; T.(?) bathycola Sterrer, 1998.

2.7.1.3 Order Bursovaginoidea Sterrer, 1972 Gnathostomulida with a bursa and often a vagina; sperm not filiform. Male opening with an injectory penis. With paired sensory organs on the rostrum, and with occipitalia. Body elongated to fairly plump (index smaller than 25); rostrum rather blunt, often plump (index usually less than 3), and mostly delimited by a sulcus. Pharynx musculature concentrated; jaws open-lamellar to fused-lamellar, without wing-shaped apophyses; solid symphysis, if present, longer than wide. Exceptionally, basal plate and jaws are lacking.

2.7.1.4 Suborder Scleroperalia Sterrer, 1972 Bursovaginoidea with a cuticular bursa and sometimes a vagina. Usually with a male stylet consisting of concentrically arranged cuticular rods. Testes paired, situated ventrolaterally. Sperm small (usually ~3 µm, maximum 13 µm), round, polygonal, or droplet-shaped, often with a bunch of short filaments. Sensorium consisting of 3–5 pairs of compound cirri and usually 1 pair of apicalia. Ciliary pits present or reduced. 2.7.1.4.1 Family Gnathostomariidae Sterrer, 1972 Scleroperalia with 1 pair of apicalia and 4 pairs of short (maximum 35 µm) compound cirri, of which the ventralia originate closer to the rostralia than to the dorsalia. Ciliary pits present. With an unpaired pre-pharyngeal gland. Epidermal cells not in stripes, epidermal inclusions scattered. Male stylet not of the rod type and hardly cuticularized or lacking. Sperm fairly large, round, and with a bunch of fairly long filaments. Basal plate shield-like; jaws more or

 189

less lamellar but not closed, without a cauda; teeth connected basally by a cuticular membrane and arranged in a ventrorostra1 arc. Without a jugum. Without ability to swim backward. One described genus (Gnathostomaria Ax). Genus Gnathostomaria Ax, 1956 (Figs. 2.24 A, B and 2.27) Gnathostomariidae with a fairly long rostrum (index ~1.7) and elongated mouth opening. Jaws with crista-like lamellae. Basal plate wider than long, with teeth at its rostra1 edge. With a muscular seminal vesicle. Type (and only) species: G. lutheri Ax, 1956. 2.7.1.4.2 Family Rastrognathiidae Kristensen & Nørrevang, 1977 Scleroperalia with 1 pair of apicalia and 4 pairs of compound cirri. Ciliary pits present. Paired pre-pharyngeal glands. Epidermal cells not in stripes. Male stylet consisting of 6 rods. Sperm small and round, totally covered with microvilli.Basal plate and jugum lacking. Jaws lamellate with bulbous cauda. Teeth long, arranged in a single arc. One described genus (Rastrognathia). Genus Rastrognathia Kristensen & Nørrevang, 1977 (Figs. 2.6 F, 2.16, 2.25 D, and 2.30) Type (and only) species: R. macrostoma Kristensen & Nørrevang, 1977. 2.7.1.4.3 Family Agnathiellidae Sterrer, 1972 Scleroperalia without paired apicalia, but with 5 pairs of short (30 µm) compound cirri, of which the ventralia originate closer to the rostralia than to the dorsalia. Ciliary pits present. With an unpaired pre-pharyngeal gland. Epidermal cells not in stripes, epidermal inclusions scattered. Male stylet of the rod type. Sperm small, round or polygonal, usually with a bunch of short filaments. Basal plate usually lacking. Jaws lacking or very delicate, lamellar but not closed, without a cauda, but with a solid symphysis; with teeth connected basally by a cuticular membrane and arranged in a ventrorostral bow. Without a jugurn. With ability to swim backward. Two described genera (Agnathiella and Paragnathiella). Genus Agnathiella Sterrer, 1971a (Figs. 2.6 C and 2.12 A) Agnathiellidae without a basal plate and without jaws. Usually with a clover-shaped rostrum; often with a dorsal protrusion in the posterior body region. Type species: A. beckeri Sterrer, 1971a. Two described species: A. beckeri Sterrer, 1971a; A. nominata Sterrer, 2001.

190 

 2 Phylum Gnathostomulida

Genus Paragnathiella Sterrer, 1997 Agnathiellidae with delicate jaws but without a basal plate. Type (and only) species: P. trifoliceps Sterrer, 1997.

Genus Paucidentula Sterrer, 1998 Paucidentulidae with rostrum narrower than the body. Sperm droplet-shaped. Type (and only) species: P. anonyma Sterrer, 1998.

2.7.1.4.4 Family Mesognathariidae Sterrer, 1972 Scleroperalia without or with 1 pair of apicalia and 4 pairs of short (30 µm) compound cirri, of which the ventralia originate closer to the rostralia than to the dorsalia. Ciliary pits present. With an unpaired pre-pharyngeal gland. Epidermal cells not in stripes, epidermal inclusions scattered. Male stylet of the rod type. Sperm small, round or polygonal, usually with a bunch of short filaments. Basal plate shield-like and often very delicate; jaws lamellar but not closed, without a cauda; teeth connected basally by a cuticular membrane and arranged in one or two ventrorostral arcs. Without a jugum. Usually with ability to swim backward. Three described genera (Mesognatharia, Labidognathia, and Tenuignathia).

2.7.1.4.6 Family Clausognathiidae Sterrer, 1992 Scleroperalia with elongated, pointed rostrum (index greater than 3) that lacks paired sensory cirri. Epidermal cells not in stripes. Male stylet of the rod type. Sperm small, round or polygonal. Basal plate may be absent. Jaws lamellar but not closed, without a cauda; teeth uniform, arranged in a ventrorostral arc. Without ability to swim backward. One described genus (Clausognathia Sterrer).

Genus Mesognatharia Sterrer, 1966b (Figs. 2.4 E, 2.6 B, 2.25 B, and 2.26 B, E) Mesognathariidae with 1 pair of apicalia. Lamellar jaws with ventral apophyses, teeth arranged in 1 or 2 ventral arcs. Basal plate shield-like, slightly wider than long. Type species: M. remanei Sterrer. Three described species: M. remanei Sterrer, 1966b; M. bahamensis Kirsteuer, 1969a; M. eastwardiae Sterrer, 1998. Genus Labidognathia Riedl, 1970 (Fig. 2.26 C, F) Mesognathariidae without paired apicalia. Lamellar jaws with long shoulder lamella, teeth arranged in two ventral arcs. Basal plate shield-like, much longer than wide. Type (and only) species: L. longicollis Riedl, 1970a. Genus Tenuignathia Sterrer, 1976 (Figs. 2.25 A and 2.26 A–D) Mesognathariidae with 1 pair of apicalia. Jaws with long ventral apophyses, teeth arranged in one ventrorostral arc. Basal plate lacking. Type species: T. rikerae Sterrer, 1976. Two described species: T. rikerae Sterrer, 1976; T. vitiensis Sterrer, 1991a. 2.7.1.4.5 Family Paucidentulidae Sterrer, 1998 Scleroperalia with one pair of apicalia and 3–4 pairs of compound cirri. Ciliary pits absent. Epidermal cells in stripes. Jaws without a cauda, lamellar but open, with few teeth in a rostroventral arc. Without a jugum. Without ability to swim backward. One described genus (Paucidentula Sterrer).

Genus Clausognathia Sterrer, 1992 (Fig. 2.4 C) Clausognathiidae without a basal plate. Poisterior region of gut without a lumen, consisting of highly vaculolized cells. Male organ with a vesicular seminalis and a vesicular granulorum. Type (and only) species: C. suicauda Sterrer, 1992. 2.7.1.4.7 Family Problognathiidae Sterrer & Farris, 1975 Scleroperalia without distinct paired apicalia and with 4 pairs of short (30 µm), evenly spaced compound sensory cirri. Ciliary pits absent. With paired buccal and pharyngeal glands. Epidermal cells not in stripes; epidermal inclusions scarce. Bursa and prebursa round, interlocking. Male stylet of the rod type. Sperm small and round, possibly with a bunch of short filaments. Basal plate broad; jaws rather lamellar and closed, without a cauda. Teeth arranged in rows. Without a jugum. Without ability to swim backward. One described genus (Problognathia Sterrer & Farris). Genus Problognathia Sterrer & Farris, 1975 (Fig. 2.31) Problognathiidae with a very short rostrum index (ca. 0.6) and elongated mouth opening. Without a tail. Jaws with 2 short rows of teeth; a terminal toothis developed. Basal plate broad (index ca. 0.3); the concave anterior edge set with teeth. Pharynx protrusile. Type (and only) species: P. minima Sterrer & Farris, 1975. 2.7.1.4.8 Family Onychognathiidae Sterrer, 1972 Scleroperalia with one pair of apicalia and 4 pairs of long (50–60 µm) compound cirri of which the ventralia originate equally from or closer to the dorsalia than to the rostralia. Ciliary pits lacking. With an unpaired prepharyngeal gland. Epidermal cells in stripes of 2–3, epidermal inclusions often in groups. Male stylet of the rod type. Sperm small, droplet-shaped. Basal plate broad; jaws rather lamellar but not closed, with a cauda and long

2.7 Systematics 

teeth arranged in a ventrorostrodorsal basket. Without a jugum. Without ability to swim backward. Five described genera (Onychognathia, Valvognathia, Nanognathia, Vampyrognathia, and Goannagnathia). Genus Onychognathia Riedl, 1971a (Figs. 2.4 B, 2.6 E, 2.24 C, D, and 2.28 A–C) Onychognathiidae with a very short rostrum (index ~0.6) and elongated mouth opening. Jaws with long teeth, no terminal tooth developed. Epidermal inclusions in groups, confined to the ventral body surface. Basal plate very delicate, simple, and without a distinct central pad. Cauda short, pear-shaped. Type species: O. filifera Riedl, 1971a. Three described species: O. filifera Riedl, 1971a; O. bractearotunda Ehlers & Ehlers, 1973; O. rhombocephala Sterrer, 1998. Genus Nanognathia Sterrer, 1972 Plump Onychognathiidae with fairly short rostrum (index ~0.7) and elongated mouth opening. Epidermal inclusions in groups confined to the ventral body surface. Jaws with fairly long teeth; a terminal tooth is developed. Cauda elongated. Basal plate delicate, simple and without a distinct central part. Type (and only) species: N. exigua Sterrer, 1972. Genus Valvognathia Kristensen & Nørrevang, 1978 (Fig. 2.29) Plump, with short rostrum and very small, spindle-shaped mouth opening. Provided with long cilia (oralia). Very delicate, broad, 5-lobed basal plate with distinct caudal extension of the median lobe. Type (and only) species: V. pogonostoma Kristensen & Nørrevang, 1978. Genus Vampyrognathia Sterrer, 1998 (Figs. 2.4 D, 2.6 D, 2.26 C, and 2.28 D) Slender Onychognathiidae (index 9–13) with a long rostrum (index 1.5–2) and long mouth opening. Basal plate very delicate, buckle-shaped. Jaws with very long teeth in a ventral and a rostral group; no terminal tooth developed. Type species: V. horribilis Sterrer, 1998. Three described species: V. horribilis Sterrer, 1998; V. minor Sterrer, 1998; V. varanus Sterrer, 2001. Genus Goannagnathia Sterrer, 2001 Slender Onychognathiidae with very slender rostrum (index less than 2) and long tail. Basal plate very delicate, buckle-shaped. Jaws with few teeth on a strong double rim. Type (and only) species: G. susannae Sterrer, 2001

 191

2.7.1.4.9 Family Gnathostomulidae Sterrer, 1972 Scleroperalia with one pair of apicalia and 4 pairs of long (ma. 65 µm) compound cirri, of which the ventralia originate closer to the dorsalia than to the rostralia. Ciliary pits present or lacking. Epidermal cells in stripes of 2–3, epidermal inclusions in furrows between the stripes. Male stylet of the rod type. Sperm small, usually dropletshaped or round with a bunch of short wings. Jaws lamellar and closed, usually with 3 horizontal rows of teeth. Cauda delicate, flanked by a pair of elongated appendages. With a jugum in the upper lip. Without ability to swim backward. Five described genera (Gnathostomula, Semaeognathia, Corculognathia, Ratugnathia, and Chirognathia). Genus Gnathostomula Ax, 1956 (Figs. 2.2 B, 2.4 A, 2.6 G, 2.32 A, B, and 2.33 B, E, G) Fairly plump Gnathostomulidae with fairly short rostrum (index ~0.8) and short mouth opening. Dorsalia normal. Ciliary pits lacking. With or without a permanent vagina. Sperm usually droplet-shaped or round with a bunch of short filaments. Usually with a well delimited tail region. Type species: G. paradoxa Ax, 1956. Twenty species described: G. paradoxa Ax, 1956, G. maldivarum Gerlach, 1958, G. murmanica Mamkaev, 1961, G. axi Kirsteuer, 1964, G. peregrina Kirsteuer, 1969a, G. jenneri Riedl, 1971b, G. microstyla Riedl, 1971b, G. nigrostoma Riedl, 1971b, G. brunidens Riedl, 1971b, G. mediocristata Riedl, 1971b, G. armata Riedl, 1971b, G. karlingi Riedl, 1971b, G. arabica Riedl, 1971b, G. mediterranea Sterrer, 1970; G. salotae Sterrer, 1991a; G. maorica Sterrer, 1991a; G. algreti Sterrer, 1991c; G. uncinata Sterrer, 1998; Gnathostomula, the most eurytopic and geographically widespread of all genera, is also taxonomically the most elusive, combining a paucity of often minute species-specific characters with high intraspecific variability (Sterrer 2006). The 20 species currently described probably represent only an approximation of the existing diversity. Genus Semaeognathia Riedl, 1970b (Figs. 2.4 F and 2.33 C, H) Fairly slender Gnathostornulidae with long rostrum (index ~2.8) and short mouth opening. Dorsalia reduced; ciliary pits present. Sperm droplet-shaped. Tail region hardly delimited. Type (and only) species: Semaeognathia sterreri Riedl, 1970b. Genus Corculognathia Ehlers & Ehlers, 1973 Slender Gnathostomulidae with well-delimited rostrum and tail region. With a jugum. Basal plate heart-shaped to

192 

 2 Phylum Gnathostomulida

trapezoid, delicate, without lateral wings or tooth-bearing central part; with caudal buckle. Jaw with 2 rows of teeth. Type (and only) species: C. apennata Ehlers & Ehlers, 1973. Genus Ratugnathia Sterrer, 1991a Fairly plump (body index ~9) Gnathostomulidae with medium rostrum (index ~1); dorsalia present, ciliary pits lacking. With a massive, unpoaired pre-pharyngela gland. Jaws with three rows of very delicate teeth; basal plate with complex central structure. Type (and only) species: R. makuluvae Sterrer, 1991a. Genus Chirognathia Sterrer & Sørensen, 2006 (Fig. 2.33 A, D, F) Gnathostomulidae with a short rostrum (index 0.99) and long mouth opening. Basal plate very delicate, transverse shield-shaped; rostral wings low-rounded, lateral wings not expressed. Jaws with 3 rows of teeth, of which the median appears as a bundle and with a paired cauda. Without a jugum. Type (and only) species: C. dracula Sterrer & Sørensen, 2006.

2.8 Biogeography To date, some 50 sites worldwide have been specifically sampled for Gnathostomulida, the majority by the first author; half of these sites remain unpublished. Although both sides of the North Atlantic are reasonably well explored, the coasts of South America, Africa (except the Cape), and most of Asia remain terra incognita, as do the northern and southern oceans, and sandy habitats at depths below 30 m (such as on continental shelves, seamounts, and ridges). An analysis of global patterns is further hampered by scarce specimens and the fact that many species (especially of the genus Gnathostomula) are not well characterized; there may well be a significant number of cryptic species. As an example, the jaws of Haplognathia ruberrima in SEM differ significantly between Bermuda and Belize specimens (see Sørensen & Sterrer 2002, figure 4), and interspecific hybridization may occur (Sterrer 1998). While the complex interplay among short-range dispersal (e.g., drifting along and across shores), in situ dispersal (e.g., continental movement, Sterrer 1973b), and speciation, which has led to the current global distribution, remains elusive (Artois et al. 2011), a few patterns are emerging: 7. At least 30% (9 of 28 known species) of Filospermoidea (but no more than 1.3%, i.e., 6 of 46 Scleroperalia) are globally distributed, with Haplognathia ruberrima recorded from 23 (of 54) localities (see Tab. 2.1).

8. About 60% of all known species have only been recorded at or in the immediate vicinity of their type locality. This is best exemplified by at least 6 well-characterized monotypic genera, i.e., Problognathia minima (Bermuda), Paucidentula anonyma (Belize), Ratugnathia makuluvae (Fiji), Goannagnathia susannae (Lizard Island), Valvognathia pogonostoma, and Rastrognathia macrostoma (Helsingør). 9. Many other species have a regional distribution, e.g., throughout the (sub)tropical NW Atlantic (Sterrer 1998), Mediterranean, or Red Sea (Sterrer, unpublished data).

2.9 Paleontology Similarities between gnathostomulid mouth parts and certain micro-conodonts have been pointed out by Ochietti & Cailleux (1969) and debated, although inconclusively (Durden et al. 1969).

Acknowledgments To make this chapter as useful in the field as much as in the laboratory, we have attempted to present the diversity of species and structures as they appear in different techniques, i.e., LM (phase contrast, bright field, and Nomarski), TEM and SEM, and drawings and reconstructions. This was only possible because a number of colleagues provided originals. We thank Pierre-Yves Pascal, Monika Müller, Johannes Achatz, and Reinhardt M. Kristensen for original figures and Gunde Rieger for providing access to the archives of Reinhard M. Rieger. We also attempted to represent Volker Lammert’s work, much of which is available only as locally printed theses, by means of originals he had shared with the first author.

Literature Ahlrichs, W. H. (1995a): Seison annulatus und Seison nebaliae. Ultrastruktur und Phylogenie. Ver. Deut. Zool. Gesell. 88: 155. Ahlrichs, W. H. (1995b): Ultrastruktur und Phylogenie von Seison nebaliae (Grube 1859) und Seison annulatus (Claus 1876), 310 pp. Cuvillier Verlag, Göttingen. Alvestad-Graebner, I. & Adam, H. (1983): Gnathostomulida. In: Adiyodi, K. G. & Adiyodi, R. G. (eds.), Spermatogenesis and sperm function, pp. 171–180. John Wiley & Sons, New York. Artois, T., Fontaneto, D., Hummon, W. D., McInnes, S. J., Todaro, M. A., Sørensen, M. V. & Zullini, A. (2011): Ubiquity of microscopic animals? Evidence from the morphological approach in species identification. In: D. Fontaneto (ed.), Biogeography of microscopic organisms: is everything small everywhere?,

Literature 

pp. 244–283. The Systematics Association, Cambridge University Press, Cambridge. Ax, P. (1956): Die Gnathostomulida, eine rätselhafte Wurmgruppe aus dem Meeressand. Abh. Akad. Wiss. Lit Mainz Math.-Nat. Klasse 8: 1–32. Ax, P. (1964): Die Kieferapparatur von Gnathostomaria lutheri Ax (Gnathostomulida). Zool. Anz. 173: 174–181. Ax, P. (1965): Zur Morphologie und Systematik der Gnathostomulida. Untersuchungen an Gnathostomula paradoxa Ax. Z. Zool. Syst. Evol. 3: 259–276. Ax, P. (1985): The position of the Gnathostomulida and Platyhelminthes in the phylogenetic system of the Bilateria. In: Morris, S,C., George, J. D., Gibson, R. & Platt, H. M. (eds.), The origins and relationships of lower invertebrates, pp. 168–180. The Systematics Association, Cambridge University Press, Cambridge. Ax, P. (1989): Basic phylogenetic systematization of the Metazoa. In: Fernholm, B., Bremer, K. & Jörnvall, H. (eds.), The hierarchy of life. Molecules and morphology in phylogenetic analysis, pp. 229–245. Proceedings from the Nobel Symposium 70. Excerpta Medica/Elsevier, Amsterdam. Ax, P. (1995): Das System der Metazoa I. Ein Lehrbuch der ­phylogenetischen Systematik. Gustav Fischer Verlag, Stuttgart. Durden, C. J., Rodgers, J., Yochelson, E. L. & Riedl, R. J. (1969): Gnathostomulida: is there a fossil record? Science 164: 855–856. Dunn, C. W., Hejnol, A., Matus, D. Q., Pang, K., Browne, W. E., Smith, S. A., Seaver, E., Rouse, G. W., Obst, M., Edgecombe, G. D., Sørensen, M. V., Haddock, S. H. D., Schmidt-Rhaesa, A., Okusu, A., Kristensen, R. M., Wheeler, W. C., Martindale, M. Q. & Giribet, G. (2008): Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452: 745–749. Eernisse, D. J., Albert, J. S. & Anderson, F. E. (1992): Annelida and Arthropoda are not sister taxa: a phylogenetic analysis of spiralian metazoan morphology. Syst. Biol. 41: 305–330. Ehlers, B. & Ehlers, U. (1973): Interstitielle Fauna von Galapagos. II. Gnathostomulida. Mikrofauna Meeresbodens 22: 173–199. Falleni, A. (1993): Ultrastructure of growing oocytes and accessory cells in Austrognathia (Gnathostomulida, Bursovaginoidea). Tissue Cell 25: 777–790. Farris, R. A. (1973): On Austrognathia strunki nov. spec. from the Florida Keys (Gnathostomulida). Int. Rev. Ges. Hydrobiol. 58: 577–586. Farris, R. A. (1977): Three new species of Gnathostomulida from the West Atlantic. Int. Rev. Ges. Hydrobiol. 62: 765–796. Farris, R. A. & O’Leary, D. J. (1985): Application of videomicroscopy to the study of interstitial fauna. Int. Rev. Ges. Hydrobiol. 70: 891–895. Fenchel, T. M. & Riedl, R. J. (1970): The sulfide system: a new biotic community underneath the oxidized layer of marine sand bottoms. Mar. Biol. 7: 255–268. Gerlach, S. A. (1958): Ein neuer Vertreter der Gnathostomulida (Turbellaria?) aus dem Meeressand der Malediven. Kieler Meeresforsch. 14: 175–176. Giribet, G., Distel, D., Polz, M., Sterrer, W. & Wheeler, W. C. (2000): Triploblastic relationships with emphasis on the position of Gnathostomulida, Cycliophora, Platyhelminthes and Chaetognatha; a combined approach of 18S rDNA sequences and morphology. Syst. Biol. 49: 539–562.

 193

Giribet, G., Sørensen, M. V., Funch, P., Kristensen, R. M. & Sterrer, W. (2004): Investigations into the phylogenetic position of Micrognathozoa using four molecular loci. Cladistics 20: 1–13. Glenner, H., Hansen, A. J., Sørensen, M. V., Ronquist, F. Huelsenbeck, J. P. & Willerslev, E. (2004): Bayesian inference of metazoan phylogeny: a combined molecular and morphological approach. Curr. Biol. 14: 1644–169 [erratum in Curr. Biol. 2005, Vol. 15, Issue 4]. Graebner, I. (1968a): Erste Befunde über die Feinstruktur der Exkretionszellen der Gnathostomulidae (Gnathostomula paradoxa, (Ax 1956) und Austrognathia riedli, (Sterrer 1965)). Mikroskopie 23: 277–292. Graebner, I. (1968b): Ergebnisse einer elektronenmikroskopischen Untersuchung von Gnathostomuliden. Ver. Deut. Zool. Gesell. Innsbruck 1968: 580–599. Graebner, I. (1969a): Vergleichende elektronenmikroskopische Untersuchung der Spermienmorphologie und Spermiogenese einiger Gnathostomula-Arten: Gnathostomula paradoxa (Ax 1956), Gnathostomula axi ( Kirsteuer 1964), Gnathostomula jenneri (Riedl 1969). Mikroskopie 24: 131–160. Graebner, I. (1969b): Ergebnisse einer licht- und elektronenmikroskopischen Untersuchung der Spermiogenese einiger Gnathostomula-Arten. Zool. Anz. Suppl. 33: 371–382. Graebner, I. & Adam, H. (1970): Electron microscopical study of spermatogenesis and sperm morphology in gnathostomulids. Accademia Nazionale dei Lincei (Roma). Quaderno 137: 375–382, +14 plates. Halanych, K. (2004): A new view of animal phylogeny. Annu. Rev. Ecol. Evol. Syst. 35: 229–256. Haszprunar, G. (1996): Platyhelminthes and Platyhelminthomorpha – paraphyletic taxa. J. Zool. Syst. Evol. Res. 34: 41–48. Herlyn, H. & Ehlers, U. (1997): Ultrastructure and function of the pharynx of Gnathostomula paradoxa (Gnathostomulida). Zoomorphology 117: 135–145. Jenner, R. A. (2004): Towards a phylogeny of the Metazoa: evaluating alternative phylogenetic positions of Platyhelminthes, Nemertea and Gnathostomulida, with a critical reappraisal of cladistic characters. Contrib. Zool. 73: 3–163. Karling, T. G. (1962): Gnathostomula paradoxa Ax, 1956 (Gnathostomulida) from the vicinity of Bergen, Norway. Sarsia 7: 23–24. Kirsteuer, E. (1964): Eine neue Gnathostomulide aus der Mangroveregion des Archipels Los Roques, Venezuela. Zool. Anz. 172: 436–440. Kirsteuer, E. (1969a): On some species of Gnathostomulida from Bimini, Bahamas. Am. Mus. Nov. 2356: 1–21. Kirsteuer, E. (1969b): Gnathostomulida – a new component of the benthic invertebrate fauna of lagoons. In: Memorias Simposio Internacional Lagunas Costeras, UNAM-UNESCO, pp. 537–544. Kirsteuer, E. (1970): On some species of meiobenthic worms of the Class Gnathostomulida from Barbados, West Indies. Am. Mus. Nov. 2432: 1–10. Knauss, E. B. (1979): Indication of an anal pore in Gnathostomulida. Zool. Scr. 8: 181–186. Knauss, E. B. & Rieger, R. M. (1979): Fine structure of the male reproductive system in two species of Haplognathia Sterrer (Gnathostomulida, Filospermoidea). Zool. Scr. 8: 181–186. Kristensen, R. M. & Funch, P. (2000): Micrognathozoa: a new class with complicated jaws like those of Rotifera and Gnathostomulida. J. Morphol. 246: 1–49.

194 

 2 Phylum Gnathostomulida

Kristensen, R. M. & Nørrevang, A. (1977): On the fine structure of Rastrognathia macrostoma gen. et sp. n. placed in Rastrognathiidae fam. n. (Gnathostomulida). Zool. Scr. 6: 27–41. Kristensen, R. M. & Nørrevang, A. (1978): On the fine structure of Valvognathia pogonostoma gen. et sp. n. (Gnathostomulida, Onychognathiidae) with special reference to the jaw apparatus. Zool. Scr. 7: 179–186. Lammert, V. (1984): The fine structure of spiral ciliary receptors in Gnathostomulida. Zoomorphology 104: 360–364. Lammert, V. (1985): The fine structure of protonephridia in Gnathostomulida and their comparison with Bilateria. Zoomorphology 105: 308–316. Lammert, V. (1986a): Ciliäre Receptoren der Gnathostomuliden. Ver. Deut. Zool. Gesell. München 1986: 283–284 (abstract). Lammert, V. (1986b): Vergleichende Ultrastruktur-Untersuchungen an Gnathostomuliden und die phylogenetische Bewertung ihrer Merkmale. Ph.D. thesis, Univ. of Göttingen, 217 pp. Lammert, V. (1989): Fine structure of the epidermis in Gnathostomulida. Zoomorphology 109: 131–144. Lammert, V. (1991): Gnathostomulida. In F. W. Harrison and E. E. Ruppert, eds. Microscopic anatomy of invertebrates, vol. 4, pp. 19–39. Wiley-Liss, New York. Lanfranchi, A. & Falleni, A. (1998): Ultrastructural observations on the male gametes in Austrognathia sp. (Gnathostomulida, Bursovaginoidea). J. Morphol. 237: 165–176. Leisch, N., Dirks, U., Gruber-Vodicka, H. R., Schmid, M., Sterrer, W. & Ott, J. A. (2011): Microanatomy of the trophosome region of Paracatenula cf. polyhymnia (Catenulida, Platyhelminthes) and its intracellular symbionts. Zoomorphology 130: 261–271. Littlewood, D. T. J., Telford, M. J., Clough, K. A. & Rohde, K. (1998): Gnathostomulida – an enigmatic metazoan phylum from both morphological and molecular perspectives. Mol. Phylogenet. Evol. 9: 72–79. Mainitz, M. (1977): The fine structure of the stylet apparatus in Gnathostomulida Scleroperalia and its relationship to turbellarian stylets. Acta Zool. Fenn. 154: 163–174. Mainitz, M. (1979): The fine structure of gnathostomulid reproductive organs. I. New characters in the male copulatory organ of Scleroperalia. Zoomorphologie 92: 241–272. Mainitz, M. (1983): Gnathostomulida. In: Adiyodi, K. G. & Adiyodi, R. G. (eds.) Oogenesis, oviposition and oosorption, vol. 1, pp. 169–180. John Wiley & Sons, New York. Mainitz, M. (1988): Gnathostomulida. In: Adiyodi, K. G. & Adiyodi, R. G. (eds.) Accessory sex glands, vol. 3, pp. 61–71. John Wiley & Sons, New York. Mamkaev, Y. V. (1961): A new representative of the Gnathostomulids: Gnathostomula murmanica sp. n. Dokl. Akad. Nauk SSSR 141: 1501–1504 [English translation: Dokl. Biol. Sci. Sect. Transl. 141: 1115–1117]. Meglitsch, P. A. & Schram, F. R. (1991): Invertebrate zoology. Oxford University Press, Oxford. Meixner, J. (1938): IVb. Turbellaria (Strudelwürmer) II. System. In: Bronn, H. G. (ed.), Die Tierwelt der Nord- und Ostsee, pp. 147–207 [unpublished page proof only]. Müller, U. & Ax, P. (1971): Gnathostomulida von der Nordseeinsel Sylt mit Beobachtungen zur Lebensweise und Entwicklung von Gnathostomula paradoxa Ax. Mikrofauna Meeresbodens 9: 1–41.

Müller, M. & Sterrer, W. (2004): Musculature and nervous system of Gnathostomula peregrina (Gnathostomulida) shown by phalloidin labeling, immunohistochemistry and cLSM and their phylogenetic significance. Zoomorphology 123: 169–177. Nielsen, C. (2001): Animal evolution. Interrelationships of the living phyla, 2nd edition. Oxford: Oxford University Press. Ochietti, S. & Cailleux, A. (1969): Comparaison des Conodontes et des mâchoires de Gnathostomulides. C. R. Acad. Sci. Paris, 268: 2664–2666. Pascal, P.-Y., Bellemare, C., Sterrer, W., Boschker, H. T. S., GonzalezRizzo, S. & Gros, O. (2014): Diet of Haplognathia ruberrima (Gnathostomulida) in a Caribbean marine mangrove. Mar. Ecol. (in press). Peterson, K. J. & Eernisse, D. J. (2001): Animal phylogeny and the ancestry of bilaterians: inferences from morphology and 18S rDNA gene sequences. Evol. Dev. 3: 170–205. Powell, E. N., Crenshaw, M. A. & Rieger, R. M. (1979): Adaptations to sulfide in the meiofauna of the sulfide system. I. 35S-sulfide accumulation and the presence of a sulfide detoxification system. J. Exp. Mar. Biol. Ecol. 37: 57–76. Powell, E. N., Bright, T. J., Woods, A. & Gittings, S. (1983): Meiofauna and the thiobios in the East Flower Garden Brine Seep. Mar. Biol. 73: 269–283. Reise, K. (1981): Gnathostomulida abundant alongside polychaete burrows. Mar. Ecol. Progr. Ser. 6: 329–333. Reisinger, E. (1961): Morphologie der Coelenteraten, acoelomaten und pseudocoelomaten Würmer. Fortschr. Zool. 13: 1–82. Riedl, R. (1966): Faunitische Studien am Roten Meer im Winter 1961/62, III Die Aufsammlungen in Suez und Al-Ghardaqa, nebst einigen Bemerkungen über Gnathostomulida. Zool. Jahr. Syst. 93: 139–157. Riedl, R. J. (1969): Gnathostomulida from America – this is the first record of the new phylum from North America. Science 163: 445–442. Riedl, R. (1970a): On Labidognathia longicollis, nov. gen., nov. spec. from the West Atlantic Coast (Gnathostomulida). Int. Rev. Ges. Hydrobiol. 55: 227–244. Riedl, R. (1970b): Semaeognathia, a new genus of Gnathostomulida from the North American Coast. Int. Rev. Ges. Hydrobiol. 55: 359–370. Riedl, R. (1971a): On Onychognathia, a new genus of Gnathostomulida from the tropical and subtropical West Atlantic. Int. Rev. Ges. Hydrobiol. 55: 227–244. Riedl, R. J. (1971b): On the genus Gnathostomula (Gnathostomulida). Int. Rev. Ges. Hydrobiol. 56: 385–496. Riedl, R. & Rieger, R. (1972): New characters observed on isolated jaws and basal plates of the family Gnathostomulidae (Gnathostomulida). Z. Morphol. Tiere 72: 131–172. Rieger, R. M. (1976): Monociliated epidermal cells in Gastrotricha: significance for concepts of early metazoan evolution. Z. Zool. Syst. Evol. 14: 198–226. Rieger, R. M. & Mainitz, M. (1977): Comparative fine structure of the body wall in Gnathostomulida and their phylogenetic position between Platyhelminthes and Aschelminthes. Z. Zool. Syst. Evol. 15: 9–35. Rieger, R. M. (1985): The phylogenetic status of the acoelomate organization within the Bilateria: a histological perspective. In: Morris, S. C., George, J. D., Gibson, R. & Platt, H. M. (eds.), The

Literature 

origins and relationships of lower invertebrates, pp. 101–122. Clarendon Press, Oxford. Rieger, R. M., Rieger, G. E., Tyler, S. & Powell, E. N. (1986): Fine structure of mitochondria in meiobenthic organisms of the marine thiobios (abstract). Am. Zool. 26: 196. Rieger, R. M. & Tyler, S. (1995): Sister-group relationship of Gnathostomulida and Rotifera-Acanthocephala. Invertebr. Biol. 114: 186–188. Schiemer, F. (1973): Respiration rates of two species of gnathostomulids. Oecologia (Berl.) 13: 4030–4406. Schram, F. R. & Ellis, W. N. (1994): Metazoan relationships: a rebuttal. Cladistics 10: 331–337. Sørensen, M. V. (2000): An SEM study of the jaws of Haplognathia rosea and Rastrognathia macrostoma (Gnathostomulida), with a preliminary comparison with the rotiferan trophi. Acta Zool. 81: 9–16. Sørensen, M. V. (2002): Phylogeny and jaw evolution in Gnathostomulida, with a cladistic analysis of the genera. Zool. Scr. 31: 461–480. Sørensen, M. V. (2003): Further structures in the jaw apparatus of Limnognathia maerski (Micrognathozoa), with notes on the phylogeny of the Gnathifera. J. Morphol. 255: 131–145. Sørensen, M. V., Funch, P., Willerslev, E., Hansen, A. J. & Olesen, J. (2000): On the phylogeny of the Metazoa in the light of Cycliophora and Micrognathozoa. Zool. Anz. 239: 297–318. Sørensen, M. V. & Sterrer, W. (2002): New characters in the gnathostomulid mouth parts revealed by scanning electron microscopy. J. Morphol. 253: 310–334. Sørensen, M. V., Sterrer, W. & Giribet, G. (2006): Gnathostomulid phylogeny inferred from a combined approach of four molecular loci and morphology. Cladistics 22: 32–58. Sørensen, M. V., Tyler, S., Hooge, M. D. & Funch, P. (2003): Organization of pharyngeal hard parts and musculature in Gnathostomula armata (Gnathostomulida: Gnathostomulidae). Can. J. Zool. 81: 1463–1470. Sterrer, W. (1965): Gnathostomula axi Kirsteuer und Austrognathia (ein weiteres Gnathostomuliden-Genus) aus der Nordadria. Z. Morphol. Ökol. Tiere 55: 783–795. Sterrer, W. (1966a): Gnathostomula paradoxa Ax und Vertreter von Pterognathia (ein neues Gnathostomuliden-Genus) von der schwedischen Küste. Ark. Zool. Ser. 2, 18: 405–413. Sterrer, W. (1966b): Neue Gnathostomulida. Ver. Inst. Meeresforsch. Bremerhaven, Sonderband 2: 201–208. Sterrer, W. (1969): Beiträge zur Kenntnis der Gnathostomulida I. Anatomie und Morphologie des Genus Pterognathia Sterrer. Ark. Zool. Ser. 2, 22: 1–125. Sterrer, W. (1970a): On some species of Austrognatharia, Pterognathia and Haplognathia nov. gen. from the North Carolina coast (Gnathostomulida). Int. Rev. Ges. Hydrobiol. 55: 371–385. Sterrer, W. (1970b): Turbellaria, Gnathostomulida. In: Riedl, R. (ed.) Fauna und Flora der Adria, 2nd edition, pp. 203–205. P. Parey, Hamburg and Berlin. Sterrer, W. (1971a): On the Biology of Gnathostomulida. Vie Milieu Suppl. 22: 493–507. Sterrer, W. (1971b): Gnathostomulida: problems and procedures. Smithsonian Contrib. Zool. 76: 9–15. Sterrer, W. (1971c): Agnathiella beckeri nov. gen. nov. spec. from southern Florida: the first gnathostomulid without jaws. Int. Rev. Ges. Hydrobiol. 56: 215–225.

 195

Sterrer, W. (1972): Systematics and evolution within the Gnathostomulida. Syst. Zool. 21: 151–173. Sterrer, W. (1973a): On Nanognathia, a new gnathostomulid genus from the East Coast of the United States. Int. Rev. Ges. Hydrobiol. 58: 105–115. Sterrer, W. (1973b): Plate tectonics as a mechanism for dispersal and speciation in interstitial sand fauna. Neth. J. Sea Res. 7: 200–222. Sterrer, W. (1974): Gnathostomulida. In: Giese, A. C. & Pearse, J. (eds.) Reproduction of marine invertebrates, Vol. 1. Acoelomate and pseudocoelomate metazoans, pp. 345–357. Academic Press, New York. Sterrer, W. (1976): Tenuignathia rikerae nov. gen., nov. spec., a new gnathostomulid from the West Atlantic. Int. Rev. Ges. Hydrobiol. 61: 249–259. Sterrer, W. (1986): Phylum Gnathostomulida. In: Sterrer, W. (ed.) Marine fauna and flora of Bermuda, pp. 211–213. Wiley, New York. Sterrer, W. (1991a): Gnathostomulida from Fiji, Tonga and New Zealand. Zool. Scr. 20: 107–128. Sterrer, W. (1991b): Gnathostomulida from Hawaii. Zool. Scr. 20: 129–136. Sterrer, W. (1991c): Gnathostomulida from Tahiti. Zool. Scr. 20: 137–146. Sterrer, W. (1991d): Triplignathia adriatica, new genus and species and typology of mouth parts in Austrognathiidae (Gnathostomulida). Proc. Biol. Soc. Wash. 104: 640–646. Sterrer, W. (1992): Clausognathia, a new family of Gnathostomulida from Belize. Proc. Biol. Soc. Wash. 105: 136–142. Sterrer, W. (1997): Gnathostomulida from the Canary Islands. Proc. Biol. Soc. Wash. 110: 186–197. Sterrer, W. (1998): Gnathostomulida from the (sub)tropical northwestern Atlantic. Stud. Nat. Hist. Carib. Region 74: 1–178. Sterrer, W. (2001): Gnathostomulida from Australia and Papua New Guinea. Cah. Biol. Mar. 42: 363–395. Sterrer, W. (2004): Gnathostomulida from the Twin Cays, Belize, mangrove community. Atoll Res. Bull. 520: 1–3. Sterrer, W. (2006): Gnathostomulida from the Otago Peninsula, southern New Zealand. Zootaxa 1172: 1–19. Sterrer, W. (2011): Two species (one new) of Gnathostomulida (Bursovaginoidea: Conophoralia) from Barbados. Proc. Biol. Soc. Wash. 124: 141–146. Sterrer, W. & Farris, R. (1975): Problognathia minima n. g., n. sp., representative of a new family of Gnathostomulida, Problognathidae n. fam. from Bermuda. Trans. Am. Microsc. Soc. 94: 375–367. Sterrer, W., Mainitz, M. & Rieger, R. M. (1985): Gnathostomulida: enigmatic as ever. In: Morris, S. C., George, J. D., Gibson, R. & Platt, H. M. (eds.), The origins and relationships of lower invertebrates, pp. 183–199. Clarendon Press, Oxford. Sterrer, W. & Müller, M. (2004): Musculature and nervous system of Gnathostomula peregrina (Gnathostomulida) shown by phalloidin-labeling, immunohistochemistry and cLSM and their phyletic significance. Zoomorphology 123: 169–177. Sterrer, W., Salvenmoser, W. & Rieger, R. M. (2001): Sperm ultrastructure of Gnathostomaria lutheri Ax (Gnathostomulida: Scleroperalia) I. An hypothesis about the origin of micropodia in scleroperalian sperm. PSZN – Mar. Ecol. 22: 3–11.

196 

 2 Phylum Gnathostomulida

Sterrer, W. & Sørensen, M. V. (2006): Chirognathia dracula gen. et spec. nov. (Gnathostomulida) from the west coast of N. America. Mar. Biol. Res. 2006: 296–302. Tyler, S. (1976): Comparative ultrastructure of adhesive systems in the Turbellaria. Zoomorphologie 84: 1–76. Tyler, S. & Hooge, M. D. (2001): Musculature of Gnathostomula armata Riedl 1971 and its ecological significance. PSZN – Mar. Ecol. 22: 71–83.

Westphalen, D. (1993): Stromatolitoid microbial nodules from Bermuda – a special micro habitat for meiofauna. Mar. Biol. 117: 145–157. 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. Zrzavý, J. (2003): Gastrotricha and metazoan phylogeny. Zool. Scr. 32: 61–81.

Martin V. Sørensen and Reinhardt M. Kristensen

3 Micrognathozoa 3.1 Introduction

Micrognathozoa is a taxon that currently consists of one single species, Limnognathia maerski Kristensen & Funch, 2000. Limnognathia maerski is a microscopic, acoelomate, vermiform animal that ranges in size from about 100 µm to 150 µm (Figs. 3.1–3.5). The body can be divided into three main regions: a head, an accordion-like thorax, and an abdomen. The integument on the dorsal and lateral sides consists of plates that are made up by 1–4 epidermal cells. A body cuticle is generally lacking, but dorsally, the epidermal cells are supported internally by an intracellular protein lamina and externally by a thin glycocalyx. The epidermal cells on the ventral side do not form plates, but are covered by a thick glycocalyx that resembles the thin cuticle found among species of annelids. Besides scattered ciliary sensory bristles, the dorsal and lateral parts of the integument have no ciliation, whereas the ventral side has ciliary bands anterior and lateral to the mouth. Furthermore, two rows of ciliophores extend throughout the ventral side of the thorax and abdomen to the caudal adhesive ciliary pad. A ventral mouth leads through a pharynx with a highly complex jaw apparatus to a short esophagus and a midgut that extends through the thorax and anterior part of the abdomen. An anal opening is either absent or at least present temporarily only. Excretion is done through pairs of protonephridia. Only females have been observed, and males are, if present at all, probably highly reduced and present within a very narrow time span only (Kristensen & Funch 2000, Kristensen 2002). Limnognathia maerski is periphytic and psammobiontic; it lives among submerged mosses and in the sediment of freshwater bodies. It has been recorded from polar and boreal regions, but is best known from its type locality at Disko Island in West Greenland. The species was first discovered during a biological field course at the Danish Arctic Station on Disko Island in 1994. During the course, a group of students and researchers camped on the northeast coast of the island, at a place called Isunngua, to examine the meiofauna of the homothermic springs in the area and at the sandy beach nearby. The campsite was situated next to a cold, heterothermic spring that was used to supply drinking water and water for sample processing. During a routine check of the meiofauna inhabiting this spring water, the new taxon Micrognathozoa was discovered.

The discovery prompted 6 years of detailed research on the new entity until a comprehensive description could be published by Kristensen & Funch (2000). Further detailed observations and interpretations of the pharyngeal apparatus were subsequently published by De Smet (2002) and Sørensen (2003). The phylogenetic position of Micrognathozoa was addressed by Ahlrichs (1995a, b) and Sørensen et  al. (2000) using a morphological approach and subsequently by Giribet et  al. (2004) using molecular sequence data. A contribution to the group’s geographic distribution was given by De Smet (2002) who recorded the species from the Subantarctic island Ile de la Possession in the Crozet Islands archipelago. Additional contributions that summarize our current knowledge about Micrognathozoa have been provided by Kristensen (2002) and Funch & Kristensen (2002). Most recently, the musculature of L. maerski was addressed using confocal laser scanning microscopy (Bekkouche et al. 2014).

3.2 Morphology 3.2.1 General and external morphology Limnognathia maerski is strictly meiofaunal with mature specimens measuring from 101 to 152 µm in total length. Juveniles have been recorded measuring between 85 and 107 µm. The three body regions, head, thorax, and abdomen, are easily distinguished in dorsoventral (Figs. 3.1, 3.2, 3.4 A, C, and 3.5 B) as well as in lateral view (Figs. 3.3, 3.4 D, and 3.5 A). The head is rounded or slightly oval and is broadest medially around the ventral mouth opening. Posteriorly, the head joins the thorax that is distinctly narrower than the head. The thorax has conspicuous broad, transverse wrinkles, which give it an accordion-like appearance (Figs. 3.2, 3.3, 3.4 D, and 3.5 A). The trunk turns broader again as the thorax joins the broadly ovoid abdominal body region. The live animal can occasionally extend its body during crawling to appear elongate and vermiform (Fig. 3.4 D), but it will usually, both during swimming and crawling, maintain a relaxed condition where the broad head and abdomen, and the more narrow thorax make the three body regions easily distinguishable (Fig. 3.4 C).

198 

 3 Micrognathozoa

Fig. 3.1: Line art illustration of Limnognathia maerski in dorsal view. Abbreviations: ac, apicalia; ap, apical plate; at, apical ciliary tuft; cd, caudalia; cp, caudal plate; do, dorsalia; fr, frontalia; ja, jaws; la, lateralia; lp, lateral plate; mg, midgut; oo, oocyte; sg, salivary glands.

3.2 Morphology 

Fig. 3.2: Line art illustration of Limnognathia maerski in ventral view. Abbreviations: ac, apicalia; ad, adhesive ciliary pad; cd, caudalia; cp, caudal plate; ey, eye; fr, frontalia; hc, head ciliophore; ja, jaws; la, lateralia; mo, mouth; oo, oocyte; op, oral plate; pc, preoral ciliation; pr, protonephridium; rb, refractive body; tc, trunk ciliophore.

 199

200 

 3 Micrognathozoa

When observed alive, the cellular, integumental plates are distinct on the dorsal side of the animal (Fig. 3.4 A). The presence of the stiff sensory bristles is furthermore evident in the head region and along the thorax and abdomen (Figs. 3.1 and 3.2). On the ventral side, the two ventral rows of ciliophores form a broad ciliary sole that extends between the oral and the caudal plates (Figs. 3.2 and 3.5 B). Also, the head ciliophores and the preoral ciliary bands of the head are easily observed. The most prominent internal structures are the pharyngeal hard parts (Fig. 3.4 B) and, eventually, the ovary that may contain a large egg (Fig. 3.4 C, D). Live animals can most often be observed either swimming, using the preoral ciliary bands and ventral ciliophores, or crawling on the ventral ciliary sole. Swimming specimens often move slowly, following an α-helical pattern. During crawling, the animal may occasionally stop and attach strongly to the substrate using its caudal adhesive ciliary pad.

3.2.2 Integument The integument of L. maerski consists of unciliated dorsal and lateral epidermal cells that join and form plates (Fig. 3.1) and a ciliated ventral epidermis (Figs. 3.2 and 3.5 B). Most of the dorsal plates are formed by 2 to 4 epidermal

cells with interdigitizing cell borders. The cells are further connected through gap junctions and the unique zipper junctions, which consist of 3 to 7 osmophilic bridges that may open like zippers when treated with sodium hypochlorite (Fig. 3.6 B, C). Each plate is 2–5 µm thick, whereas the integument is less than 2 µm in the joints between the plates. A dorsal cuticle is absent, and extracellularly, the epidermis is covered by a 120-nm-thin glycocalyx only. However, the epidermal cells are supported internally by an intracellular lamina (Fig. 3.6 B). The intracellular lamina is considered homologous with the corresponding lamina found in species of the syndermate taxa Seisonacea, Acanthocephala, and Eurotatoria (Ahlrichs 1995a, b, Kristensen and Funch 2000). The head is covered with 13 plates in total. All head plates consist of 3 to 4 cells, except the large apical plate, located middorsally in the forehead (Figs. 3.1, 3.3, and 3.4 A), which consists of one single epidermal cell only. The thoracic plates are generally thicker and most rigid on the dorsal side, whereas the lateral ones appear more flexible. The anterior-most thoracic plate on the dorsal side is made up by two epidermal cells only, whereas the remaining plates consist of three to four cells. Smaller rod-like or rhomboid plates are inserted between the dorsal and lateral thoracic plates, which add to the flexibility of the thorax. The abdominal integument is made up by dorsal and lateral plates that are more uniform in shape and thickness. One triangular

Fig. 3.3: Line art illustration of Limnognathia maerski in lateral view. Abbreviations: ac, apicalia; ad, adhesive ciliary pad; ap, apical plate; at, apical ciliary tuft; cd, caudalia; do, dorsalia; fr, frontalia; hc, head ciliophore; ja, jaws; la, lateralia; oo, oocyte; rb, refractive body.

3.2 Morphology 

 201

Fig. 3.4: Light micrographs of Limnognathia maerski. (A) Dorsal view showing the dorsal integumental plates. (B) Isolated jaws, dorsal view. (C) More ventral view. (D) Lateral view. Abbreviations: ac, apicalia; ad, adhesive ciliary pad; ap, apical plate; as, accessory sclerite; cd, caudalia; cp, caudal plate; do, dorsalia; fi, fibularium; la, lateralia; lp, lateral plate; mj, main jaws; oo, oocyte; ph, pharynx; pl, pharyngeal lamella; pc, preoral ciliation; rb, refractive body.

plate is located middorsally on the posterior-most part of the abdomen and covers the area where the anus otherwise would have opened. The posterior-most plates are two paired ones, located at the caudal part of the animal and extending down on the ventral side. These plates are referred to as the caudal plates (Figs. 3.1, 3.2, 3.4 A, and 3.5 B, D). The ventral integument generally consists of multiciliated epidermal cells forming a continuous epidermis

without formation of plates. Exceptions are the two caudal plates and the large cuticular oral plate (Fig. 3.5 B). The ciliated ventral epidermis can be divided into four different regions: the preoral ciliary bands, the head ciliophores, the ventral ciliated sole, and the adhesive ciliated pad (Figs. 3.2, 3.4 D, and 3.5 A, B). The preoral ciliation is used for swimming and foraging and includes 6 rows of cilia and one U-shaped ciliary band (Fig. 3.2), all with relatively

202 

 3 Micrognathozoa

Fig. 3.5: SEM of Limnognathia maerski. (A) Lateral view. (B) Ventral view. (C) Oral plate. (D) Caudal plate. Abbreviations: ad, adhesive ciliary pad; cp, caudal plate; do, dorsalia; hc, head ciliophore; la, lateralia; op, oral plate; pc, preoral ciliation; po, paired openings of unknown function; tc, trunk ciliophore.

3.2 Morphology 

 203

Fig. 3.6: TEM micrographs of Limnognathia maerski showing details in the dorsal and lateral integument. (A) Overview of horizontal section of specimen. (B) Detail of cell border in dorsal plate. (C) Detail of framed area in (A) showing a complete zipper junction. Abbreviations: aj, adherens junction; cm, cell membrane; gj, gap junction; gl, glycocalyx; ic, intercellular space; il, intracellular lamina; lp, lateral plates; lu, midgut lumen; mg, midgut; oo, oocyte (note its close contact with the midgut); pa, pharyngeal apparatus; zj, zipper junction.

short, cross-striated ciliary roots. The ciliary bands cannot readily be homologized with bands of the rotifer corona. Four pairs of ciliophores are located in a ventrolateral position behind the level of the mouth opening and lateral to the oral plate (Figs. 3.2, 3.3, and 3.5 B). Each ciliophore consists of a rectangular epidermal cell with numerous stiff compound cilia. The cilia in each ciliophore beat in unison but are not bounded by a common membrane as seen in certain species of gastrotrichs (Ruppert 1991). The ciliary roots are longer than those in the forehead. A large, non-ciliated cuticular plate, the so-called oral plate, is located between the

mouth opening and the thorax (Figs. 3.2, 3.5 B, C, and 3.7 B). Laterally, it is delimited by the head ciliophores. The ventral side of the thorax and abdomen is densely ciliated. This ciliation is formed by two closely set rows of ciliophores, with 16 to 18 cells in each row (Figs. 3.2 and 3.5 A, B). Each ciliophore is rectangular to oval and arranged transversely to the main axis of the trunk. The extracellular covering is conspicuously thicker than the glycocalyx seen on the dorsal and lateral plates and forms a cuticle with a loose, hexagonal substructure. The cilia of the ciliophores penetrate the cuticle, and their bases are enwrapped by a

204 

 3 Micrognathozoa

Fig. 3.7: TEM micrographs of Limnognathia maerski showing details in the ventral integument. (A) Section through the adhesive ciliary pad. (B) Cross section posterior to mouth opening. (C) Section through trunk ciliophores. Abbreviations: bl, basal lamina; ce, centriole; ci, cilium; cr, ciliary root; cu, cuticle; cw, cuticular ciliary wrapping; ep, epidermal cell; gd, gland duct; js, jaw sclerite; mi, mitochondrion; mu, muscle; nu, nucleus; op, oral plate.

cuticular ring (Fig. 3.7 C). The ventral ciliophores are locomotory and are involved in crawling as well as swimming. The adhesive ciliary pad is located between the ventral ciliophores and the caudal plates (Figs. 3.2, 3.3, 3.4 D, and 3.5B). The pad is covered with cuticle and consists of 10 ciliated cells that are arranged in two paired groups with five cells in each. The cilia here are also stiff, but longer than those on the ciliophores, and they do not beat in unison. As in the trunk ciliophores, the basal parts of the cilia are enwrapped in a cuticular ring. The ciliary roots in the adhesive ciliary pad are even longer than those in the ciliophores (Fig. 3.7 A). A small pore between the two groups of cells in the pad marks the outlet of an adhesive gland. The gland is formed by two cells only and resembles a mucous gland rather than a duo-gland system. Otherwise, no epidermal glands are present in the animal.

3.2.3 Musculature The musculature of L. maerski has not yet been described in detail. Previous attempts to analyze the musculature through phalloidin staining and confocal laser scanning microscopy have not been successful;1 hence, most of the available information is based on light microscopy and transmission electron microscopy (TEM). All recorded muscles appear to consist of fibers, and each muscle is made up by a single cell. Myosyncytia or myoepithelia are not present. The muscle cell consists of several (up to nine) sarcomeres marked with z-discs (Fig. 3.8 B). 1 After this chapter went in press, the musculature of L. maerski was actually successfully described in detail using confocal laser scanning microscopy. The results are published in Bekkouche et al. 2014.

3.2 Morphology 

 205

Fig. 3.8: TEM micrographs of Limnognathia maerski showing details in the musculature. (A) Oblique horizontal section through anterior end of specimen. (B) Other section made near framed area in (A) showing cross-striated longitudinal muscles. Abbreviations: br, brain (neural cell bodies); ep, epidermal cell; lm, longitudinal muscle; lu, midgut lumen; mg, midgut; nu(m), nucleus (of muscle cell); pra, abdominal protonephridium; prt, thoracic protonephridium; sa, sarcomere; sg, salivary gland; zd, z-discs.

A number of longitudinal muscles traverse the whole body of the animal (Fig. 3.8 A). Furthermore, some shorter longitudinal fibers connect the thoracic epidermal cells and are responsible for movements in this flexible body region. Dorsoventral muscles are mostly located laterally in the thorax region and run from the dorsalmost edges of the lateral plates down to their attachment points in a more ventrolateral position. Circular trunk muscles have not been detected. The pharyngeal apparatus is associated with a complex musculature, consisting of strong cross-striated fibers that connect the various elements in the jaws. The pharyngeal muscles always attach through a junction formed by an epidermal cells; hence, direct contact

between the muscle cells and the jaw elements never occurs. All muscles in L. maerski are considered to have a mesodermal origin, but this needs confirmation from embryological studies.

3.2.4 Nervous system The central nervous system consists of a brain and a pair of ventral, longitudinal nerve cords. The brain is formed by a large cerebral ganglion that occupies most of the forehead (Figs. 3.8 A and 3.9 C). It consists of a central neuropile, surrounded by neural cell bodies that form a rim around the neuropile. Approximately 175 rounded cell

206 

 3 Micrognathozoa

Fig. 3.9: TEM micrographs of Limnognathia maerski showing details in the nervous system. (A) Cross section through the anterior part of the thorax. (B) Close up of framed area in (A) showing ventral longitudinal nerves and muscles. (C) Sagittal section through head region in specimen with partly protruded jaws. Abbreviations: br, brain (neural cell bodies); ct, collecting tubule from protonephridia; cu, cuticle; dp, dorsal plate; ep, epidermal cells; gc, gut cell; js, jaw sclerite; lm, longitudinal muscle; lp, lateral plate; lu, midgut lumen; mo, mouth; nu(b), nucleus (of neural cell bodies); op, oral plate; pc, preoral ciliation; tc, trunk ciliophore; ve, vesicle; vn, ventral longitudinal nerve cord.

bodies make up the perikarya of the outer rim. The brain appears bilobed in dorsal view, and the two lobes are connected through prominent commissures. A pair of ventral, longitudinal nerve cords extends from each brain lobe and run caudally through the entire trunk (Fig. 3.9 A). A ganglion appears to be present in the thorax and in the posteriormost part of the abdomen. A buccal ganglion may be present as well, but this needs to be confirmed. Owing to difficulties locating the basal lamina, it is difficult to determine the nature of the ventral nerve cords, but they appear to be subepidermal (Fig. 3.9 A, B).

3.2.5 Sensory structures The most prominent sensory structures in L. maerski are the ciliary sensoria that are serially scattered over the entire body (Figs. 3.1–3.3). Each sensorium consists of one to three stiff cilia that emerge through pores in or between the dorsal and the lateral plates, forming tactile bristles. The rim of the pore is formed by a collar from the intracellular lamina (Fig. 3.10 A, B). A ring of nine short microvilli may be present and emerge through the collar. For sensoria consisting of multiple cilia, it remains uncertain

3.2 Morphology 

 207

Fig. 3.10: TEM micrographs of Limnognathia maerski showing sections through sensoria. (A) Longitudinal section through lateralia on thorax. (B) Cross section through lateralia on head. Abbreviations: ci, cilium; co, collar; ec, epidermal cell; jp, junction between plates; mi, mitochondrion; mv, microvilli; sc, sensory cell.

if the cilia arise from a single cell or whether they are formed by several monociliated cells. The forehead bears 4 pairs of sensoria: the apicalia, the frontalia, and 2 pairs of lateralia (Figs. 3.1 and 3.3). The apicalia are situated closely together at the tip of the forehead. A pair of longer frontalia is located lateral to the apialia. More laterally, on the forehead, 2 pairs of lateralia emerge through pores in between the head plates. No sensoria are present in the postoral part of the head. The thorax has 2 pairs of lateralia and 2 pairs of dorsalia (Figs. 3.1, 3.3, and 3.4 A). The lateralia are located on the flexible lateral plates, whereas the dorsalia is situated in a somewhat subdorsal position. The pores of the posteriormost thoracic sensoria are double, which make them appear with an 8-shape (Figs. 3.1, 3.3, and 3.4 A). A pair of lateralia is present medially on the abdomen. More posteriorly, a pair of subdorsal dorsalia emerges. A pair of caudalia projects in a caudal direction near the posterior end of the animal, and even further back, mostly on the ventral side, a second pair of caudalia emerges through pores in the caudal plates (Figs. 3.1–3.3). Other integumentary ciliary sensory structures include a ciliary tuft between the head apicalia, a pair of long flagellum-like cilia near the frontalia, and a pair of frontolateral tufts or brooms of short cilia between the frontalia and the lateralia. Cilia in the mouth and pharynx may have a sensorial function as well, but this needs confirmation. In the forehead, a pair of rounded, vesicular, and osmiophilic structures could be interpreted as lipoid eyes granules (Fig. 3.2). Pigment eyes have been detected in neither the brain nor the forehead.

3.2.6 Intestinal system The mouth is ventral and located near the anterior edge of the oral plate. The posterior mouth lip is attached to the oral plate and appears to be stiff. Opposed to the stiff posterior lip, the anterior rim of the mouth is extremely flexible and capable of expanding to a point that allows the pharyn-

Fig. 3.11: TEM micrographs of Limnognathia maerski showing sections through the posterior part of the intestinal system. (A) Oocyte and posterior part of midgut. (B) Detail of framed area in (A) showing the terminal part of the gut and the caudal plate of the integument. Abbreviations: mg, midgut; oo, oocyte; tp, tegumental plate.

geal apparatus to get extruded through the mouth opening. When this happens, the forehead is lifted and then pulled backward in a rapid movement. The anterior part of the

208 

 3 Micrognathozoa

mouth lumen is ciliated, and the cilia appear to have a sensorial function, rather than being used to move food toward the pharynx. The mouth opening leads through the pharynx with ciliated epithelial cells and the complex pharyngeal apparatus, to a short esophagus without cilia or microvilli. Posteriorly the esophagus expands into a midgut that is devoid of cilia, but has an epithelium covered with microvilli (Figs. 3.6 A, 3.8 A, 3.9 A, and 3.11 A, B). A pair of glands opens at the border between the esophagus and the midgut. These glands are referred to as salivary glands (Fig. 3.8 A), but their precise function is still unknown. Posteriorly, the midgut tapers until a point where an actual gut lumen is no longer present (Fig. 3.11 A, B). Hence, the midgut ends at a point right below an outer epidermal plate (Fig. 3.11 B). A functional anus is not present, due to the posteriorly closing midgut, and even without this tapering of the midgut, the plate would still cover a potential anal opening. Minute muscles attached to the tegumental plate suggest that it may be movable, hence allowing the formation of a temporary anus, but this or any other indication of defecation has never been observed. The posteriormost cells in the midgut differ from the other midgut cells, as they lack the microvilli bordering. This could suggest that these cells have an ectodermal origin. However, this is contradicted by the presence of numerous vesicles and mitochondria, which

is more similar with the endodermal midgut epithelium rather than the outer ectodermal epithelial cells that are poor in these organelles. The pharynx consists of a secretory epithelium, glandular cells, a complex pharyngeal musculature, and the cuticular hard parts that make up the jaw apparatus (Fig. 3.12 A–D). Some of the epithelial cells are ciliated whereas others carry microvilli with a central osmiophilic core (Fig. 3.12 A). During ontogeny, the epithelial cells are involved in the secretion of the thin cuticular brush border that lines the pharyngeal wall. The brush border is in particular well developed in the posterior part of the pharynx, near the two fibularia (Fig. 3.12 A). A large concentration of cilia is also found in the region (Fig. 3.12 B). The epithelial cells furthermore secrete the sclerites of the pharyngeal hard parts. However, there is no sign of secretory activity in epithelial cells of adult specimens, which supports that the pharyngeal hard parts are already fully developed in the newly hatched juvenile. Numerous glandular buccal cells are scattered in the anterior part of the pharyngeal epithelium, and a cluster of ca. 20 glandular cells is located dorsally. The high number of mitochondria and numerous secretory vesicles suggest that these cells have a secretory function as well. The pharyngeal hard parts make up a jaw apparatus with a complexity that has not been recorded in any other

Fig. 3.12: TEM micrographs of Limnognathia maerski showing sections through the pharyngeal apparatus. (A) Horizontal section through fibularium. (B) Oblique section through posterior parts of fibularia. (C) Section through caudal rods of basal plates. (D) Detail showing muscle attachment point on jaw sclerite. Abbreviations: ba, basal plates; ca, cauda; ci, cilia; cu, cuticle; ep, epidermal cells; fi, fibularium; js, jaw sclerites; ju, junction; mi, microvilli; mu, muscle; ne, nerve; op, oral plate; plu, pharynx lumen; scl, scleropilar rods; zd, z-discs.

3.2 Morphology 

 209

Fig. 3.13: SEM micrographs of jaws in Limnognathia maerski. (A) Jaw apparatus, dorsal view. (B) Jaw apparatus, ventral view. (C) Jaw apparatus, lateral view, with pseudophalangia, pharyngeal lamellae, and basal plates twisted down into a ventral position. (D) Jaw apparatus, ventral view. Compare with (B) and note that the pseudophalangia, pharyngeal lamellae, and basal plates have been flipped backward, allowing a ventral view of the main jaws. Abbreviations: as, accessory sclerite; bp, basal plates; ca, cauda; dj, dorsal jaws; fi, fibularium; mjd, main jaws, dorsal part; mjv, main jaws, ventral part; pl, pharyngeal lamellae; pp, pseudophalangia.

microinvertebrate (Figs. 3.13 and 3.14). The jaw apparatus is composed of numerous individual elements, called sclerites, that are interconnected by flexible ligaments and minute muscles. The basal elements that make up each individual sclerite are the minute, tube-like scleropili. The scleropili are about 100  nm in diameter and consist of lucent, osmiophobic material surrounding an electron-dense core. Viewed with TEM in cross section, they appear like white circles with a central black dot (Fig. 3.12 C), whereas when visualized with scanning electron microscopy (SEM), they appear like rounded, knob-like structures on the surface of the sclerites. As mentioned above, the sclerites are secreted not only by the epithelial cells in the pharyngeal wall but also by epithelial cells that are located in small compartments inside some of the sclerites (Fig. 3.12 A, B).

The sclerotized parts of the jaw apparatus can be divided into four main elements: the main jaws, the fibularium, the ventral jaw elements, and the dorsal elements (Fig. 3.13 A–D). The main jaws form a set of pinchers that articulate caudally with an unpaired symphysis. Each half of the main jaw is horizontally bipartite, which divides it into a ventral and a dorsal part. The ventral part is composed of scleropili that are only loosely joined, making this part of the jaws rather flexible (Fig. 3.13 D). The s­ cleropilar composition is evident on all surfaces of the sclerites, where the knob-like terminals of the tubes give it a characteristic ornamentation. Posteriorly, the ventral main jaws attach to the dorsal main jaws, whereas the anterior ends connect with the fibularia. The dorsal main jaws are more solid than their ventral counterparts and their scleropilar composition is not evident on the surfaces. They form an

210 

 3 Micrognathozoa

Fig. 3.14: Three-dimensional reconstruction of the jaws in Limnognathia maerski. (A) Dorsorostral view. (B) Caudoventral view. (C) Ventrorostral view. (D) Laterocaudal view.

elongate shaft with teeth that basally are directed toward the main axis of the jaw apparatus (Fig. 3.13 A, D). Toward the apical part of the shafts, the teeth bend and turn more elongate, which make their direction more parallel to the jaws’ main axis. A small knob, situated medially on the dorsolateral edge of the main jaw shaft, serves as a socket for one additional set of teeth. These teeth are even more elongate and extend in an anterior direction over the tips of the dorsal main jaws. Together with the tooth rows on the inner surfaces of the dorsal main jaws, these elongate dorsal teeth form a basket-like structure that is highly characteristic for the main jaws (Figs. 3.13 A and 3.14 A, D). As stated, both halves of the main jaw pinchers articulate caudally with an unpaired, laterally compressed symphysis. A paired cauda furthermore attaches to the dorsal edge of the symphysis. The cauda is composed of two elongate, slightly bent rods that extend caudally (Figs. 3.13 A and 3.14 A–D). Together with the main jaws, the fibularium constitutes the most prominent part of the pharyngeal hard parts. The fibularium forms two paired sclerites with

internal fibulae that delimit compartments for epithelial cells (Figs. 3.12 A and 3.13 A). Each sclerite is shaped like a slice cut from the surface of a ball, giving it a spacious internal compartment. They are located lateral to the main jaws but fold in ventrally, forming most of the ventral surface of the jaw apparatus as well (Figs. 3.13 D and 3.14 B, C). The ventral and lateral surfaces of the fibularia appear delicate and slightly porous, and parts of it may eventually break open during preparation of the jaws. In the apical parts, a big rostrolateral fenestra is present (Fig. 3.13 C, D). This window appears to be a true opening, and not a preparation artifact. On the internal surface of each fibularium, four obliquely transverse fibulae define five chambers that each contains a large epithelial cell (Figs. 3.12 A, B and 3.13 A). Two tooth rows are present at the most apical part of each fibularium, close to the point where the ventral main jaws attach. The dorsal jaw elements include one pair of sclerites only. They are referred to as the dorsal jaws and are enw­ rapped in the apical folding of the fibularia (Fig. 3.13 A).

3.2 Morphology 

Each dorsal jaw sclerite is rod-shaped and composed of a proximal shaft and three distal tooth rows. The surfaces of the sclerites are densely knobbed, revealing the scleropilar composition of the elements. The ventral jaw elements include the pseudophalangia with their accessory sclerites, the pharyngeal lamellae, and the basal plates (Figs. 3.13 A–D and 3.14 C). The pseudophalangia are a pair of elongate rods with distal, digitiform teeth. Proximally, they articulate with a pair of accessory sclerites, which are small elements, composed of long, twisted scleropili. The pseudophalangia are located in a rostroventral position in the jaw apparatus, whereas the accessory sclerites are more lateroventral (Fig. 3.14 A–C). It is not perfectly clear how the articulating unit attaches to the jaw apparatus, but there appears to a ligamentous connection between the accessory sclerites and the fibularia. The pharyngeal lamellae form a pair of sclerites consisting of thin lamellae that join each other proximally and articulate with a short cylindrical socket (Figs. 3.13 B and 3.14 A–C). Also, the pharyngeal lamellae are located in a rostroventral position and tend to enwrap the distal parts of the pseudophalangia. The third set of major ventral element is composed of two paired basal plates and an unpaired central platelet. Each basal plate consists of a crescentic plate with teeth on its inner margin (Figs. 3.13 B and D and 3.14 C). An elongated, flattened rod extends caudally from the posterior parts of each of the crescentic plates and articulates proximally to the ventral surface of the main jaws through a flexible, ligamentous connection (Fig. 3.13 D). A ligament furthermore connects the two crescentic plates and a small circular platelet with scleropilar surface ornamentation that attaches between the two plates. Only little information is available about the functioning of the jaw apparatus. During crawling, the animal appears to use the pseudophalangia to grab food items. The food is grabbed in very fast snapping movements, and it has not been possible to determine whether the closely associated pharyngeal lamellae exit the mouth opening together with the pseudophalangia. If they leave the mouth as well, one could imagine that they acted like a sieve, disabling too large items to enter the mouth. However, it is more likely that they stay in the pharynx and help supporting the pharynx or mouth lumen during feeding. The fibularia obviously also have a supporting function. The epithelial cells located in the fibularia are most certainly involved in the secretion of material during the formation of the jaws. However, this process appears to happen during the embryological development, and

 211

since the jaws are fully developed by hatching of the juvenile animal, it is uncertain whether the cells play any role beyond this point. The scleropilar composition of the micrognathozoan jaws is identical with the substructure found in jaw sclerites of rotifers and gnathostomulids, which strongly suggests a homology between the pharyngeal apparatuses in these three groups (Ahlrichs 1995b, Rieger & Tyler 1995). However, some incongruence still exists about the interpretation and homologization of the specific micrognathozoan sclerites and their counterparts in rotifer species (compare Kristensen & Funch 2000, De Smet 2002, Sørensen 2003). At the present state, there is a consensus about the interpretation of the micrognathozoan main jaws, which are considered homologous with the rotifer incus (rami+fulcrum) and the gnathostomulid articularium. Kristensen & Funch (2000) and Sørensen (2003) furthermore interpret the pseudophalangia and their accessory sclerites as homologous with the rotifer mallei (unci+manubria), opposed to De Smet (2002) who suggests a homology between the fibularia and the mallei. Sørensen (2003) considers the fibularia autapomorphic for Micrognathozoa.

3.2.7 Body cavities and connective tissue Limnognathia maerski has no coelom, and indications of reduced pseudocoel or a true mesodermal coelom have not been observed (Fig. 3.6 A). Small cavities have, in a few sections, been observed between the epidermis and the gut (Fig. 3.9 A), but these are believed to be artifacts due to shrinking during fixation.

3.2.8 Excretory system Excretion in L. maerski is done through 2 pairs of protonephridia. One pair is located in the anterior part of the thoracic region and one pair anteriorly in the abdomen (Figs. 3.2 and 3.8 B). Each protonephridium consists of four ­terminal cells that form two units, with two terminal cells in each unit (Fig. 3.15 C). The terminal cell connects with a monociliated canal cell, and the two canal cells from each protonephridial unit meet in a shared canal cell (Fig. 3.15 B, C). Each canal cell is wrapped around the distal end of the terminal cell, and the two sides of the wrapped canal cell join along an autodesmosome. The four terminal

212 

 3 Micrognathozoa

cells are monociliated, and have a few mitochondria only (Fig. 3.15 A). Together, the terminal cells and canal cell form a filter region consisting of 9 to 10 stiff microvilli from each terminal cell and about 18 irregular microvilli from the canal cell (Fig. 3.15 B). According to previous descriptions (Kristensen & Funch 2000), the canal cells terminate into a nephridiopore cell that opens ventrolaterally on the animal. However, new observations indicate that the canal cells empty into collecting tubules formed by ciliated cells (Figs. 3.9 B and 3.15 C). Up to nine cilia can be observed in a cross section through the collecting tubule. It still remains uncertain where the collecting tubules terminate. Two paired openings lateral to the caudal plate (Fig. 3.5 B, D) could be interpreted as nephridiopores, which would suggest that the collecting tubules traverse the entire abdomen and join these putative nephridiopores. However, it cannot yet be rejected that the collecting tubules connect the protonephridia with a cloacal opening, which would be close to the condition found among rotifers (Clément & Wurdak 1991, Ahlrichs 1993). Further examinations are still required to clarify this.

3.2.9 Reproductive organs Males of L. maerski have never been recorded; hence, only the morphology of the female reproductive organs is currently known. The reproductive system consists of paired ovaries that join near a ventral gonopore behind the ciliary adhesive pad (Fig. 3.2). A germovitellarium, as found in rotifers, is not present in L. maerski and the yolk is believed to form within the developing oocyte. The mature, paired ovary makes up two, somehow diffuse structures, only surrounded by a thin basal lamina. During its development, the oocyte is in close contact with the midgut (Figs. 3.6 A and 3.11 A), which suggests that nutrition for the yolk is absorbed directly from the midgut. The oocyte has a large nucleus with nucleolus, is rich in mitochondria, and has a prominent endoplasmatic reticulum. Only one oocyte can develop at a time, whereas two or three immature oocytes can be observed in an animal.

3.2.10 Gametes

Fig. 3.15: TEM micrographs of Limnognathia maerski showing details in the protonephridial system. (A) Longitudinal section through terminal cell. (B) Cross section through canal cell and distal ends of terminal cells. (C) Cross section through terminal cells and collecting tubules. Abbreviations: cc, canal cell; ci, cilium; ct, collecting tubule; fi, filter region; mi, microvilli; pt1, protonephridial terminal cell 1; pt2, protonephridial terminal cell 2.

The oocyte develops while it is inside the ovarium. Prior to oviposition the egg have reached a remarkable size, up to 48 µm, and are hence filling up most of the abdomen (Fig. 3.4 D). Oviposition is believed to happen through the ventral gonopore. Two kinds of mature eggs

have been observed. One kind, referred to as the “sticky eggs”, has an unsculptured but sticky surface (Fig. 3.16 A), whereas the second kind, the “sculptured eggs”, displays a thicker and strongly sculptured shell (Fig. 3.16 B).



3.3 Reproduction and development 

 213

The sticky eggs were previously considered aborted eggs, but because both kinds of eggs have been observed developing within live animals, it appears to be two distinctive types. Animals with sticky eggs appear to be most frequent in the midsummer, whereas the sculptured eggs are most numerous in the late summer.

3.3 Reproduction and development Because males have never been observed, reproduction is considered to be purely parthenogenetic. However, the presence of two egg types could suggest that yet ­undiscovered males were involved at some point in the reproductive cycle. In the life cycle found among species of monogonont rotifers, amictic eggs are produced during the parthenogenetic cycle, whereas production of mictic resting eggs involves exchange of gametes between the sexes. Hence, in the mictic phase of the rotifer cycle, reduced dwarf males are present in a short period. Because the sticky, thin-shelled eggs show some resemblance with the amictic rotifer eggs and the sculptured eggs resemble rotifer resting eggs, one may speculate whether male gametes were involved in the production of the latter. This would imply either that males, probably very small and strongly reduced, were present for a very short sexually reproductive period or that some of the known females went through a short hermaphroditic stage that enabled them to produce male gametes. A pair of refractive bodies (Figs. 3.2, 3.3, and 3.4 A) that could be interpreted as testicular structures has been observed in very young stages of L. maerski, which would suggest that they hatched as males and then switched sex during development. In that way, they would be protandric hermaphrodites. However, this is highly hypothetical and needs confirmation from further studies. Other observations suggest that the sticky eggs generally are produced by young individuals, whereas older ones produce the sculptured eggs later in the summer. These observations in themselves do not contradict that the reproductive cycle could be purely parthenogenetic. Nothing is known about cleavage, early embryology and the formation of germ layers. Oviposition has never been observed, whereas it has been possible to observe a juvenile during hatching from the egg. The newly hatched juvenile resembles the adult and deviates at minor points only, thus development is considered to be direct. The juvenile is slightly smaller than an adult specimen, down to 85 µm, and only 10 pairs of ciliophores appear to be present on the ventral sole. However, the jaws are already fully developed at hatching, and behavior and locomotion resemble the patterns observed in adults.

Fig. 3.16: SEM micrographs showing the two kinds of eggs produced by Limnognathia maerski. (A) The “sticky egg”, which is assumed to be parthenogenetic. (B) The “sculptured egg”, assumed to be a resting egg.

3.4 Physiology Most aspects of the physiology in L. maerski still need to be explored. Because its habitat may remain frozen for up to 8 months per year, at least the eggs must have a physiology that enables them to survive this long hibernation.

3.5 Phylogeny The putative homology between the pharyngeal hard parts in species of Micrognathozoa, Gnathostomulida, and Syndermata strongly supports a close relationship between these taxa, and together with the aberrant, parasitic acanthocephalans, these taxa are usually united in the clade Gnathifera (Fig. 3.17) (Ahlrichs 1995a, b,

214 

 3 Micrognathozoa

Fig. 3.17: Cladogram showing the phylogeny of the Gnathifera, with synapomorphies and autapomorphies indicated on branches. (1) Gnathifera: Jaws composed of scleropilar rods with a lucent material and an electron dense core; jaws with pincers caudally articulating into unpaired pedicle. (2) Gnathostomulida: Epidermal cells monociliated; hermaphroditism; jaws with unpaired basal plate. (3) Jaws with pseudophalangia/unci and associate sclerites/manubria; jaws with pharyngeal lamellae/epipharynx; cellular epidermis with intracellular lamina. (4) Micrognathozoa: Intracellular lamina forming epidermal plates, restricted to dorsal and lateral sides; head and trunk with sensoria formed by monociliate or multiciliate cells; cilia on trunk and parts of head arranged as ciliophores; jaws with fibularium, paired basal plates, and dorsal jaws. (5). Rotifera: Syncytial epidermis; trunk ciliation lost; head ciliation arranged in bands forming a wheel-organ; body divided into head, trunk, and foot; germovitellarium. (6) Monogononta: Heterogamic reproduction. (Bdelloid rotifer in tree is redrawn from Hyman 1951.)

3.6 Systematics 

Kristensen & Funch 2000, Sørensen et al. 2000). Within Gnathifera, Micrognathozoa constitute the sister group to Syndermata. This sister-group relationship is supported by the presence of an epidermis with an intracellular lamina and perhaps the formation of differentiated mallei in the pharyngeal apparatus (Kristensen & Funch 2000, Sørensen 2003). De Smet (2002) takes it one step further and includes L. maerski in the Eurotatoria (Bdelloidea+Monogononta), as sister taxon to the Monogononta. However, the presence of a cellular epidermis, monociliated protonephridial terminal cells and the lack of a differentiated wheel organ make this position less likely. Even though morphological characters support gnathiferan monophyly, it has not been possible to confirm this relationship with molecular data. In the only molecular study that has explored the phylogenetic position of L. maerski, the species branched out in various positions depending on the data source and analytical parameter settings (Giribet et al. 2004). The analyses were based on micrognathozoan sequence data from complete 18S rRNA, a 316-bp fragment of 28S rRNA, a sequence from the mitochondrial proteincoding gene cytochrome c oxidase subunit I, and the nuclear protein-coding gene Histone 3. The analyses only supported gnathiferan monophyly when the ribosomal loci were analyzed alone and only under one specific parameter set. Under other parameter sets, Micrognathozoa cluster with one of the rotifer main groups (Monogononta or Bdelloida+Acanthocephala), as sister group to Syndermata (but with Gnathostomulida outside Gnathifera), or as sister group to either Cycliophora or Entoprocta. Interestingly, the latter two often show molecular affinities to the syndermatan taxa, but this result also conflicts with morphological interpretations. Hence, at present, molecular data give only weak hints about the phylogenetic position of Micrognathozoa, and monophyletic Gnathifera based on morphological evidence may currently be considered the best supported hypothesis. Another potential relative to Micrognathozoa could be the enigmatic diurodrilid worms. Diurodrilidae has traditionally been considered a polychaete family that originally was assigned to the now abandoned “Archiannelida”. However, recent studies have questioned their polychaete affinities, and especially their conspicuous ciliophores could indicate a closer relationship head ­ with Micrognathozoa (Worsaae & Rouse 2008). However, the phylogenetic position of Diurodrilidae still remains uncertain, and it needs to be further addressed in future studies.

 215

3.6 Systematics Currently, only a single species, L. maerski, is known from Micrognathozoa. Besides recordings from and near its type locality on Disko Island, the species has been recorded in Wales, UK (J. M. Schmid-Araya, personal communication), and from the Subantarctic Crozet Islands (De Smet 2002). The long distance between the collecting localities would alone imply that at least the Southern Hemisphere species was not conspecific with L. maerski, but it has not been possible to point out any morphological diagnostic differences between the specimens from Greenland and those from the Crozet Islands. However, a 2% difference between 18S rRNA sequences from specimens belonging to the two populations could indicate a cryptic speciation.

3.7 Biogeography The type locality of L. maerski is the spring area of Isunngua, at the Northeast Coast of Disko Island, Greenland. It has furthermore, at a single occasion, been recorded from another spring, about 5 km from the Isunngua spring area, near the Lymnaea Lake (Kristensen & Funch 2000). The only other record of the species on the Northern Hemisphere was done in a swampy spring area in Wales, UK. This location has not been reported in the scientific literature, and is therefore only known as personal communication from Dr. Schmid-Araya. On the Southern Hemisphere, L. maerski has been collected at various localities on Ile de la Possession in the Crozet archipelago, about 2400 km north of Antarctica, and 2400 km southeast of South Africa (De Smet 2002). Even though the species always occurred in relatively low numbers, it appeared to be widespread in freshwater bodies on the island. This bipolar distribution may seem odd, and De Smet (2002) suggests that the species could have been introduced at the Crozet Islands by whalers in the 19th or 20th century or together with the introduction salmon around 1970. He states, however, that these explanations also are less likely. The most plausible explanation, which is also supported by De Smet (2002), would probably be that L. maerski is more widespread than the few occasional recordings would reflect. There are several examples of freshwater rotifer species that have been recorded at single occasions, or if more than one record exist, that they have been recorded on localities situated on different continents or hemispheres. Hence, we find it likely that L. maerski still may have a bipolar distribution, but could be widespread, in low numbers, however, throughout the cold and temperate regions on both hemispheres.

216 

 3 Micrognathozoa

3.8 Paleontology Limnognathia maerski has, as most other soft bodied microinvertebrates, never been found in the fossil record. The pharyngeal hard parts would probably have the capability of fossilization, but due to their minute size, it is not likely that they would ever be recorded in a sample.

3.9 Ecology Limnognathia maerski occurs in well-oxygenated water with low conductivity and a pH around 7 (6.4 to 7.4). In springs on Disko Island, it is always found in association with submerged mosses, whereas specimens from Ile de la Possession were collected from various substrates, including mosses, gravel, and coarse sand. At both localities, the species was present in streams with running water, but on Ile de la Possession, it was also found in water bodies with more stagnant water. The species has been found in water with temperatures ranging from 1.5°C to around 14°C, which indicates that the animal is cold stenothermic. When kept at room temperature, the animals die rather fast. On Disko Island, a few young specimens have been recorded readily after the break of the ice in the end of May, but the population is small until it peaks from mid-July to mid-August. On Ile de la Possession, specimens have been present from November to January, which is the period that the localities have been sampled. The spring area at Isunngua on Disko Island is formed by heterothermic springs, which means that the temperature changes with the seasons and that the springs freeze during the Arctic autumn and stay frozen until the end of May. To survive, L. maerski would have to have mechanisms to withstand this long period of freezing. Apparently, the adult specimens do not possess any cryptobiotic capabilities, as found in species of rotifers, tardigrades, and nematodes; thus, it seems most likely that they survive as resting eggs. The sculptured eggs resemble the rotifer resting eggs that are able to withstand long-term freezing and dehydration, and it is likely that the sculptured eggs of L. maerski have similar abilities. If they are able to withstand not only freezing but also d ­ ehydration, the sculptured eggs could also be the dispersal stage and spread with the wind or attached to birds. It should be stressed, however, that it has still not been possible to

hatch live specimens from sculptured eggs that have been either frozen or dehydrated.

Literature Ahlrichs, W. H. (1993): Ultrastructure of the protonephridia of Seison annulatus (Rotifera). Zoomorphology 113: 245–251. Ahlrichs, W. H. (1995a): Seison annulatus und Seison nebaliae. Ultrastruktur und Phylogenie. Verhand. Dtsch. Zool. Ges. 88: 155. Ahlrichs, W. H. (1995b): Ultrastruktur und Phylogenie von Seison nebaliae (Grube 1859) und Seison annulatus (Claus 1876). Cuvillier Verlag, Göttingen, 310 pp. Bekkouche, N., Kristensen, R. M., Hejnol, A., Sørensen, M. V. & Worsaae, K. (2014): Detailed reconstruction of the musculature in Limnognathia maerski (Micrognathozoa) and comparison with other Gnathifera. Front Zool. 11: 71. Clément, P. & Wurdak, E. (1991): Rotifera. In: Harrison, F. W. & Ruppert, E. E. (eds.) Microscopic anatomy of invertebrates, vol. 4: Aschelminthes, pp. 219–297. Wiley-Liss, New York. De Smet, W. H. (2002): A new record of Limnognathia maerski Kristensen & Funch, 2000 (Micrognathozoa) from the subantarctic Crozet Islands, with redescription of the trophi. J. Zool. 258: 381–393. Funch, P. & Kristensen, R. M. (2002): Coda: the Micrognathozoa – a new class or phylum of freshwater meiofauna. In: Rundle, S. D., Robertson, A. L. & Schmid-Araya, J. M. (eds.) Freshwater meiofauna: biology and ecology, pp. 337–348. Backhuys Publishers, Leiden. Giribet, G., Sørensen, M. V., Funch, P., Kristensen, R. M. & Sterrer, W. (2004): Investigations into the phylogenetic position of Micrognathozoa using four molecular loci. Cladistics 20: 1–13. Hyman, L. H. (1951): The invertebrates: Acanthocephala, Aschelminthes, and Entoprocta. McGraw-Hill, New York, 572 pp. Kristensen, R. M. (2002): An introduction to Loricifera, Cycliophora, and Micrognathozoa. Integr. Comp. Biol. 42: 641–651. Kristensen, R. M. & Funch, P. (2000): Micrognathozoa: a new class with complicated jaws like those of Rotifera and Gnathostomulida. J. Morphol. 246: 1–49. Rieger, R. M. & Tyler, S. (1995): Sister-group relationship of Gnathostomulida and Rotifera-Acanthocephala. Invertebr. Biol. 114: 186–188. Ruppert, E. E. (1991): Gastrotricha. In: Harrison, F. W. & Ruppert, E. E. (eds.) Microscopic anatomy of invertebrates, Vol. 4: Aschelminthes, pp. 41–109. Wiley-Liss, New York. Sørensen, M. V. (2003): Further structures in the jaw apparatus of Limnognathia maerski (Micrognathozoa), with notes on the phylogeny of the Gnathifera. J. Morphol. 255: 131–145. Sørensen, M. V., Funch, P., Willerslev, E., Hansen, A. J. & Olesen, J. (2000): On the phylogeny of the Metazoa in the light of Cycliophora and Micrognathozoa. Zool. Anz. 239: 297–318. Worsaae, K. & Rouse, G. W. (2008). Is Diurodrilus an annelid? J. Morphol. 269: 1426–1455.

Diego Fontaneto and Willem H. De Smet

4 Rotifera

4.1 Introduction Rotifera comprise about 2,000 species (Segers 2007) of microscopic animals, usually much less than 1  mm in length, characterized by the presence of a ciliated corona and a muscular pharynx called mastax (Wallace et  al. 2006). Traditionally, 3 groups are recognized in Rotifera: Bdelloidea, Monogononta, and Seisonacea. Each group is peculiar for general morphology and ecology, but mostly for reproductive modes: Seisonacea are exclusively sexual and live as epibionts on the crustacean Nebalia; Monogononta are facultative cyclical parthenogens, free-living in freshwater and marine waters (Wallace et  al. 2006); Bdelloidea are obligate parthenogens living in any wet or moist habitat, capable of surviving desiccation through dormancy (Ricci & Fontaneto 2009). The position of Rotifera in the tree of life is clear: they belong to the spiralian clade Platyzoa, together with Platyhelminthes, Gnathostomulida, Gastrotricha, Micrognathozoa, and Acanthocephala (Dunn et al. 2008, Hejnol et al. 2009, Edgecombe et al. 2011). But the relationships among the groups of Rotifera, i.e., Bdelloidea, Monogononta, and Seisonacea, are not clear at all, as their relationships with Acanthocephala (see Section 4.5, Phylogeny). Rotifera can be found in any habitat where water becomes available, even for short periods, due to their peculiar ability to produce dormant stages that resist absence of water (Gilbert 1974, Ricci 2001). The mechanisms through which resting stages are produced and survive in the absence of water are different between Bdelloidea and Monogononta, whereas no resting stages are known for Seisonacea. Dormancy makes these animals an interesting model system to understand the mechanisms of desiccation resistance, freeze tolerance, starvation, survival in extreme environmental conditions, and ageing (King 1969, Pouchkina-Stantcheva et al. 2007, Hilhorst 2008, Austad 2009, Snell et al. 2012). Moreover, the ability to recover after dormancy involves mechanisms that restore living conditions, including the potentials for repairing DNA and the resistance to ionizing radiation (Gladyshev & Meselson 2008). Another very peculiar feature of Rotifera is their asexual reproduction (Schön et  al. 2009); Monogononta are cyclical parthenogens, whereas Bdelloidea

are strictly parthenogenetic. Bdelloidea have even been framed as “evolutionary scandal” because they evolved and diversified in the absence of sexual recombination (Maynard Smith 1986, Judson & Normark 1996). The aspects of asexuality are very intriguing (Fussmann 2011), and Bdelloidea are known to have rather peculiar mechanisms to counteract the problems of strictly asexual reproduction (Gladyshev et  al. 2008, Wilson & Sherman 2010). Rotifers are used in aquaculture (Lubzens 1987, Lubzens et  al. 1989) where mass production has been developed (Hagiwara et  al. 1997) to provide live food for fish fry, and as model organisms for ecotoxicology (Ramírez-Pérez et  al. 2004, Dahms et  al. 2011). Most of the experimental work performed on rotifers is based on species that have been cultured for aquaculture or for ecotoxicology, so that a great deal of information is available for the Brachionus plicatilis species complex, the most widely cultured of all rotifers. Rotifer research started with the first microscopist, Anthony van Leeuwenhoek (1632–1723), who described a Bdelloidea in one of his letters from 1687 (Wallace et al. 2006). Since its origin, the history of rotifer research has had 2 main aspects. On the one hand, there are lines of research that focus on the organisms and try to understand rotifers in themselves; on the other hand, scientists from several branches of science work using rotifers as a model system to answer questions of broad interests. The history of rotifer research is, thus, a continuous mingling of these 2 complementary approaches. The rotifer meetings, held every 3  years since 1976, are a combination of different focuses and provide a great avenue for interaction among rather different scientists with different approaches, gathered together by the interest for this fascinating group of animals. The proceedings of these meetings are usually published in Hydrobiologia, and we will continuously refer to papers from such meetings throughout the text.

4.2 Morphology While various investigators contributed to the knowledge of morphology and anatomy of rotifers following their description by van Leeuwenhoek, the start for the

218 

 4 Rotifera

great advance in anatomy and histology was given by Zelinka (1886, 1888, 1891, 1892a, b), who introduced histological techniques, e.g., the use of microtome-cut sections. Zelinka studied different organ systems, especially the nervous system of bdelloids, discovered the mastax and caudal ganglion, and revealed the course of the major nerves and branching-off nerves until their endings. Superior pioneering anatomohistological work was done by de Beauchamp, dealing with the retrocerebral apparatus (de Beauchamp 1905, 1906), corona (de Beauchamp 1907a), integument, intestinal system (de Beauchamp 1907a, b, 1909), and mastax (de Beauchamp 1908, 1909) of all rotifer orders. Eutely was first demonstrated for the brain of Epiphanes, Eosphora, Euchlanis, and Notommata by Hirschfelder (1910) and later on for different organs in Epiphanes senta by Martini (1912) and Asplanchna priodonta by Nachtwey (1925). The latter authors and Hlava (1905), Lehmensick (1926), Seehaus (1930), Waniczek (1930), Peters (1931), Stossberg (1932), Remane (1933), Dehl (1934), Brakenhoff (1937), to mention a few (for a review of studies prior to 1929, see Remane 1933), made painstakingly detailed studies of the different organ systems that are still reliable at large. An important synoptic work on rotifer morphology and anatomy was presented by Remane (1929) and followed by his classical and impressive work published between 1929 and 1933 (Remane 1933). Further comprehensive reviews are by, e.g., Hyman (1951), de Beauchamp (1965), and Wallace et  al. (2006). Specific morphoanatomical information on the different groups of rotifers are presented in the works dealing with taxonomy and identification keys (e.g., Bartoš 1951, Bdelloidea; Voigt 1957, Bdelloidea and Monogononta; Donner 1965, Bdelloidea; Kutikova 1970, Monogononta; Koste 1978, Monogononta; Kutikova 2005, Bdelloidea); an update for several families is presented in the more recent identification guides (Segers 1995, Nogrady et al. 1995, De Smet 1996, De Smet & Pourriot 1997, Nogrady & Segers 2002). In the past 25 years, several new techniques were introduced for the study of rotifer organ and cellular organization. Ultrastructural investigations using transmission electron microscopy (TEM) and/or scanning electron microscopy (SEM) were performed on the different organ systems (e.g., Clément & Wurdak 1991, Clément 1993, Melone 1998, Riemann & Ahlrichs 2010). Detailed study of the structure of the trophi became possible by SEM (e.g., Markevich 1989, Kleinow et  al. 1990, De Smet, 1998, Melone et  al. 1998a). Immunohistochemistry, epifluorescence, and confocal laser scanning microscopy (CLSM) was used to investigate the

nervous system (e.g., Kotikova 1995, 1998, Hochberg 2007, 2009, Leasi et  al. 2009), and epifluorescence and CLSM were likewise used to study musculature (e.g., Hochberg & Litvaitis 2000, Santo et  al. 2005, Sørensen 2005a, b, Hochberg & Ablak Gorbuz 2008, Leasi & Ricci 2010).

4.2.1 General and external morphology Descriptions of rotifer species refer to the females (if not otherwise stated), as males are absent in Bdelloidea and are smaller and of much simpler organization in Monogononta, and moreover have been rarely investigated.

4.2.1.1 Morphology of the female The body shape is enormously diverse, but generally bilaterally symmetrical, with a clear differentiation between both ends and between the ventral and the dorsal sides (Fig. 4.1). Usually, there is a head, trunk, and foot more or less clearly marked off by transversal folds. The foot is always located behind the cloacal aperture, which allows its localization even when the trunk grades imperceptibly into the foot. The body often appears segmented due to the presence of permanent transversal folds in the integument. These are considered pseudosegments, as there is no true segmentation in rotifers. A pseudosegment marked off between head and trunk, called neck, is often present. Head and foot are usually retractable into the trunk. The head bears the characteristic rotatory apparatus or corona, used for locomotion and/or food collection by its cilia, and the mouth and several sense organs, including sensory bristles and pits, eyespots, ciliated tentacles, and one or more dorsal antennae. The apical region can be provided with palps (e.g., Gastropus), ciliated auricles (e.g., Notommata, Synchaeta), a retractile (Colurella, Lepadella) or non-retractile shield (e.g., Squatinella, Cotylegaleata), a snout-like process or proboscis (Rhinoglena), apical tentacles (e.g., Collotheca hoodi, Ptygura pectinifera), a cup-shaped sucker-like structure (Cotylegaleata), a projecting rostrum, etc. The rostrum of the monogononts is a dorsal hood-shaped structure, usually offset by a fold, at the anterior end of the body (e.g., many Dicranophoridae); the apertures of the retrocerebral organ lie ventral to the rostrum. The rostrum (proboscis) of the bdelloids (Fig. 4.2) is a complex, unpaired and retractable, blunt-conical organ with ciliated tip provided with styli, and lamella(e). It is usually composed of 2 pseudosegments, often bearing the apertures of the retrocerebral organ and eyespots when present (e.g., most Rotaria, Adineta oculata). When

4.2 Morphology 

 219

Fig. 4.2: The rostrum of Bdelloidea. (A) Rotaria macroceros, (B) Philodina acuticornis, (C) Mniobia incrassata, (D) Philodina brevipes, (E) Adineta grandis, (F) Zelinkiella synaptae, (G) Macrotrachela natans, (H) Habrotrocha angusticollis, (I) Embata hamata, (J) Adineta cf. barbata, and (K) Adineta barbata. Abbreviations: fe, frontal eye; ln, lateral rostral nerve; mg, medial rostral ganglion; mn, medial rostral nerve; rl, rostral lamella; sc, sensory cell. (Modified from Remane 1933.)

Fig. 4.1: Scheme of the rotifer anatomy. (A) Dorsal and (B) lateral. Abbreviations: H, head; T, trunk; F, foot; b, brain; bl, bladder; c, corona; co, cloacal opening; da, dorsal antenna; gg, gastric gland; i, intestine; lt, lateral antenna; m, mastax; p, protonephridium; o, ovarium; pg, pedal gland; ro, retrocerebral organ; s, stomach; sg, salivary gland; v, vitellarium. (Modified from Wallace & Ricci 2002.)

the corona is withdrawn, the rostrum is extended, forming the frontal part of the head; with unfolded corona, it is retracted, becoming dorsal. The bdelloid rostrum is involved in tactile perception and locomotion: during creeping, the rostrum is brought in contact with the substrate and used for adhesive attachment, producing leech-like movements (e.g., Hochberg & Litvaitis 2000). The head encloses the brain, the glandular retrocerebral organ, and often part of the mastax. The trunk may be cylindrical, fusiform, sacciform, ovate, spherical, etc. In cross section, it is circular, triangular, laterally enlarged, or compressed to various degrees, both dorsoventrally or laterally. The ventral surface is often more or less flattened, and the dorsal one is mostly more or less strongly arched. In Trichocercidae, the cylindrical body is spirally twisted to the left. The posterodorsal end of the trunk is often extended into a more or less prominent projection or tail, overhanging the cloacal aperture. The tail may be unlobed, bilobed, trilobed, or extended into a conical (Notommata tripus) or spiniform process (Proalinopsis staurus, Dorystoma caudatum). The trunk

bears the lateral antennae (lacking in bdelloids), usually situated distally on either side, but displaced to the coronal field in Conochilidae; in Testudinella, the trunk carries the dorsal antenna as well. The integument can be smooth, but in species with stiffened integument or lorica, it often exhibits a great variety of ornamentation, such as longitudinal or transversal ridges or furrows, punctuations, spines, pustules, tubercles, polygonal or circular fossettes, etc.; the different adornments can be arranged in a way to make larger polygonal (e.g., Keratella, Platyias, Trichotria), longitudinal (e.g., Notholca), or transversal (e.g., Lophocharis) patterns on the lorica. SEM of the tegument surface reveals numerous pores on small elevated areas (Ricci & Melone 1984, 1998a). The trunk of many species is provided with movable or nonmovable projections or appendages serving different functions. Projections occur at the anterior and posterior margins of the trunk, on the dorsal surface, or along the entire body (except the ventral surface), and less frequently on the head and foot. Spines enhance buoyancy of planktonic species (e.g., Brachionus, Keratella, Kellicottia) and serve as defense mechanism against being swallowed by predators (e.g., Brachionus, Keratella; see Section 4.4, Physiology). In bdelloids (e.g., Pleuretra hystrix), spines help to anchor to the substrate (Fontaneto & Melone 2003). Paddles (Polyarthra), armlike appendages bearing stiff bristles (Hexarthra), and movable setae (Filinia), all with strong muscles inserted at their base, serve for jumping locomotion to escape predators, or for predator deterrence (e.g., Stemberger & Gilbert 1987, Gilbert 1999). Asplanchna enhances

220 

 4 Rotifera

­ rotection against gape-limited predators by an increase p of size, expanding soft portions of its integument into lateral outgrowths, through an increase of the hydrostatic pressure of its pseudocoel. Two large, dorsal conical outgrowths behind the corona in Synchaeta bicornis and S. fennica are supposed to compensate, by an increase of their volume, for the increased internal pressure upon contraction of the body (Remane 1929). In Philodina alata, a similar function may be attributed to the paired muscularized lateral outgrowths, obvious only in contracted animals. The trunk contains the gastrointestinal, excretory, and reproductive organs. A foot is usually present, and in most species, it is located at the end of the trunk, but it may be displaced ventrally (e.g., Gastropus, Ploesoma, Testudinella) or twisted out of the body axis (e.g., Trichocerca). In planktonic monogononts, the foot is mostly absent (e.g., Asplanchna, Ascomorpha, Horaella, Notholca, Filinia, Keratella, Pompholyx, Trochosphaera). The foot may be very short to very long and is usually composed of a few to many telescopically retractable pseudosegments, or it is cylindrical with a ringed or wrinkled appearance (e.g., Brachionus, Ptygura, Testudinella). The foot is generally partly to completely retractable in the trunk. In soft-bodied rotifers, it can be simply withdrawn into the body, or in species with a stiffened, almost inflexible lorica, the foot is retracted through the foot opening. In sessile species (Flosculariidae, Collothecidae), the foot of adult females is mostly very long and modified into a contractile stalk that may terminate in a non-contractile, narrow columnar pedicel. A short discoid foot is shown by Cupelopagis. In most Ploima, a caudal antenna is usually present dorsally on the distal pseudosegment.

The foot contains the pedal glands producing adhesive secretions and terminates in the attachment organs, such as 1–4 toes, an adhesive disc, or a ciliated ring. Toes may be absent (e.g., Asplanchna, Ascomorpha, Conochilus, Horaella, Notholca, Filinia, Keratella, Pompholyx, Trochosphaera) or vestigial (e.g., Proalides, Synchaeta monopus), especially in the planktonic species. Size, shape, and structure of the toes show a huge range of morphological variation and adaptations in monogonont Ploima (Fig. 4.3). Most of the ploimid genera have 1 to 2 symmetrical toes; unequal toes are present in Monommata and Trichocerca. An additional spur can be present between the toes (e.g., Squatinella rostrum, Trichotria pocillum) or laterally from each toe (Cotylegaleata) and 2–8 needle-shaped substyli at the base of the toes are characteristic in Trichocerca. The toes may be very long and surpass the length of the body (Beauchampiella, Monommata, Scaridium). They may be conical, lanceolate, leaf-shaped, needle-shaped, etc., and straight or curved. They are generally not pseudosegmented but are often provided with an offset distal part, the claw (e.g., Cephalodella, Dicranophorus, Lecane); the toe(s) of some Lecane species show up to 3 pseudosegments. Depending on the stiffening of their integument, toes can be flexible (e.g., Lindia, Notommata, Synchaeta) or rigid (e.g., Trichotria, Mytilina, Lecane); in some species, there is a rigid basal part and a soft distal part (e.g., Encetrum frenoti, Notommata pachyura). Attachment of adults of the sessile Flosculariacea and Collothecacea is by an adhesive disc, or the end of the foot is undifferentiated, whereas their juvenile stages have a ciliated ring at the end of the foot; the foot of the adult free-swimming Testudinella bears a ciliated cup.

Fig. 4.3: Variation of the foot and toes in Monogononta. (A) Cephalodella forficula, (B) Notommata voigti, (C) Cephalodella gibba, (D) Lepadella sp., (E) Lecane luna, (F) Dicranophoroides caudatus, (G) Brachionus diversicornis, (H) Monommata sp., (I) Scaridium longicaudum, (J) Trichocerca pusilla, (K) Trichocerca tigris; (L) Dicranophorus cambari, (M) Lecane bulla, (N) Microcodon clavus, (O) Testudinella sp., (P) Cupelopagis vorax, larva, (Q) Collotheca sp., and (R) Collotheca sp. Abbreviations: a, adhesive disc; c, ciliated cup; f, foot; p, non-contractile pedicel; t, toe. (From Donner 1956, Koste 1978, Kutikova 1970, Wulfert 1957.)

4.2 Morphology 

Variation in morphology of the toes is lower in bdelloids (Fig. 4.4). Toes in bdelloids are generally short, conical, and flexible, and their number is 2, 3, or 4: a pair of ventral toes (Didymodactylos), a single dorsal toe, and a pair of ventral ones (e.g., Macrotrachela, Rotaria, Adineta), or a pair of smaller dorsal and a pair of larger ventral toes (e.g., Dissotrocha, Philodina, Philodinavus). Toes may be lacking, and attachment is by an undivided (e.g., Anomopus, Zelinkiella) or cleft adhesive disc (e.g., Mniobia). The penultimate foot pseudosegment carries dorsally or dorsolaterally 2 appendages, the spurs, characteristic of most bdelloids; Henoceros only shows a single dorsal spur. Depending on the structure and function of the foot and its toes, the following types are recognized in rotifers: creeping foot: foot and toes short, few pseudosegments (Notommatidae, Bdelloidea); swimming foot: foot long cylindrical and unsegmented or with several long pseudosegments (e.g., Brachionus, Testudinella, Rotaria neptunia) or foot short with long toes used for steering when swimming (e.g., Euchlanis, Cephalodella, Trichocercidae); jumping foot: pseudosegments and/or toes very long, provided with strong muscles (Beauchampiella, Monommata, Scaridium), used for, e.g., escape reactions; sessile foot: long retractile foot, often with adhesive disc (sessile Flosculariidae, Collothecidae). Most rotifers are colorless and very transparent, but transparency may be reduced in species with a thick granulated lorica. Rotifers often appear colored by the content of the digestive system because of the pigments ingested with the food (brownish, greenish), or presence of symbiotic zoochlorellae (green). In a few species, whole animals or distinct organs are colored because of their own pigments. A red coloring is most common, especially in species inhabiting the Arctic-alpine environment

 221

(e.g., Philodina gregaria), and shades of yellow, brown, violet, and dark blue occur in the diverse taxa. Several rotifers secrete protective sheaths or tubes (see Section 4.2.2, Integument).

4.2.1.2 Coloniality Some 25 species belonging to 8 genera of mostly sessile monogononts (Flosculariidae, Conochilidae) form permanent colonies varying in size from a few individuals to 3,500 animals (see Wallace 1987, Wallace et al. 2006). Colony formation can be allorecruitive, autorecruitive, or geminative. In the allorecruitive colony formation, the free-swimming solitary juveniles settle on the tubes of conspecifics, forming an arborescent colony (e.g., Limnias ceratophylli, Floscularia conifera, F. ringens). When the juveniles remain within the parental colony, the latter will reach a critical size, and subsequently splits up into 2 or more daughter colonies (autorecruitive colony formation in, e.g., Conochilus hippocrepis and C. unicornis). The colonies are spherical, with the individuals radiating from the centre, each animal having its own mucous tube (C. unicornis), or the tubes coalescing into a single spherical mass (C. hippocrepis). In the geminative colony formation, newborns leave the parent colony together as a planktonic juvenile colony (e.g., Lacinularia flosculosa, Sinantherina socialis). The sheath-forming bdelloid Rotaria macroceros often aggregates in large numbers sticking together.

4.2.1.3 Corona The basic organization of the corona was elucidated by light microscopy (LM) (de Beauchamp 1907a, 1908, 1965, Remane 1933) and later on largely confirmed

Fig. 4.4: Variation of the foot, toes (t), adhesive discs (ad) and spurs (s) in Bdelloidea. (A) Rotaria socialis, (B) Philodina gregaria, (C) P. flaviceps, (D) Habrotrocha pavida, (E) Macrotrachela crucicornis, (F) M. asperula, (G) M. muricata, (H) Embata hamata, (I, J) Zelinkiella synaptae, (K) Mniobia circinata, (L) M. obtusicornis, (M) M. russeola, and (N) M. armata. (From Remane 1933.)

222 

 4 Rotifera

by SEM in its essential outlines (e.g., Clément & Wurdak 1991, Melone & Ricci 1995, Melone 1998, Ricci & Melone 1998a). The ground plan of the corona (Fig. 4.5) comprises a ciliated area, the buccal field, surrounding the usually ventrally located mouth opening. The buccal field is evenly ciliated with short cilia. It extends upward around the head and forms a circumapical band, delimiting an unciliated apical field. The cilia of the ciliary ring at the anterior margin of the circumapical band are strong and form the preoral trochus – a ciliary row of usually finer cilia at the posterior margin forms the postoral cingulum. Between the trochus and the cingulum runs a finely ciliated groove. The apical field bears numerous sensory receptors and is often provided with setae and tufts or rows of cilia (styli, cirri, membranelles). When the circumapical band consists only of a single row of cilia, forming a preoral or postoral ciliary ring

with the anterior or posterior margin of the buccal field, it is called paratrochus and paracingulum, ­respectively. Ciliary rings or arcs formed only by the preoral part of the buccal field are called pseudotrochus. The corona is absent in the parasitic genus Balatro. In many species, the marginal cilia of the circumapical band are elongated and often situated on a pair of lateral ear-like projections (Fig. 4.6), the auricles (e.g., Lindia, Notommata, Synchaeta, Tetrasiphon). Auricles are expanded and assist when swimming; they can be withdrawn in the head when creeping (Notommata). In monogononts, the distal part of the corona on which the cingulum inserts can be developed into a ­prominent pad or fold, the under lip (e.g., Hexarthra, Filinia); in bdelloids, both under and upper lips are present. The ultrastructure of the cilia (Fig. 4.7) was described by, e.g., Lansing & Lamy (1961), Clément (1977), Clément & Wurdak (1991). The axoneme of the motile cilia shows the characteristic 9 peripheral doublets and 2 central tubules. The basal body consists of doublets or singlets instead of the classical triplets. Intramembranous particles are present (Brachionus calyciflorus) over the entire surface

Fig. 4.5: Basic organization of the rotary apparatus and other organs of the head. (A) Dorsal, (B) lateral, and (C) ventral. Abbreviations: af, apical field; b, brain; bf, buccal field; bt, buccal tube; c, cingulum; ca, circumapical band; ce, cerebral eye; da, dorsal antenna; m, mouth; mx, mastax; o, ocellus; op, esophagus; or, orifice of retrocerebral organ; rs, retrocerebral sac; sg, subcerebral glands. (Modified from Donner 1956, de Beauchamp 1965, and Clément & Wurdak 1991.)

Fig. 4.6: Ciliated auricles. (A) Notommata copeus, (B) Lindia fulva, (C) Tetrasiphon hydrocora, and (D) Synchaeta sp. (From Remane 1933.)

Fig. 4.7: Diagram of the cilia of the rotatory apparatus, in longitudinal and cross section. (A) cilia from the trochus and (B) cilia from the buccal field. Abbreviations: bd, belt desmosome; ed, epitheliomuscular desmosome; em, electron-dense material; m, muscle; sr, striated ciliary rootlet. (Modified from Clément & Wurdak 1991, with permission.)

4.2 Morphology 

of the cilia of the tactile bristles or are aligned along the cilia of the pseudotrochus and base of the motile cilia of the corona. In some cilia, an electron-dense material is present at the tip (Notommata copeus, Trichocerca rattus, Philodina roseola), or where doublets become singlets, it extends along the central doublets to the tip (Brachionus sericus). The ciliary rootlets are striated and occur in 2 types (Scholtyseck & Danneel 1962, Clément 1977, 1987, Clément & Amsellem 1989). The vertical rootlets penetrate into the cytoplasm, inserting on epitheliomuscular desmosomes, or to a fibrous intracytoplasmic layer, inserted itself on desmosomes between coronal cells and integument (the desmosomal belt). The horizontal rootlets interconnect the cilia at their base and insert upon the desmosomal belt. Horizontal ciliary roots may be absent (Asplanchna). The muscles inserted on the ciliary roots of the cingulum and pseudotrochus (Brachionus calyciflorus, B. plicatilis) may control the ciliary beat. They are innervated and show characteristics of very fast fibers. Various modifications of the ground plan of the corona have been described, which grossly correlate with the mode of locomotion and feeding habits. Corona types in Monogononta (see de Beauchamp 1907a, 1908, 1965, Remane 1933, Melone 1998) (Figs. 4.8 A–K and 4.9) Notommata type. Extensive buccal field around mostly ventral and central mouth opening. Circumapical band

 223

and apical field present; apical field fairly small; circumapical band laterally with paired tufts of stronger cilia; trochus and cingulum weakly developed. In creeping or slow-swimming animals (Notommatidae). Dicranophorus type. Corona almost exclusively composed of extensive buccal field surrounding central mouth opening. Circumapical band absent or reduced to paired tufts of stronger cilia laterally from buccal field. In creeping or slowswimming animals (Dicranophoridae, some Notommatidae). Asplanchna type. Buccal field strongly reduced. Apical field large. Circumapical band an interrupted circle of cilia around the large apical field; trochus and cingulum not differentiated. Planktonic species (Asplanchnidae, Synchaetidae, Gastropodidae, Trichocercidae, several Notommatidae, e.g., Eosphora, Sphyrias). Euchlanis-Brachionus type. Only supraoral part of buccal field well developed; marginal cilia powerful, enlarged to stiff cirri, the rest of the buccal field with weak cilia, or transverse rows and arcs of cilia/cirri above and laterally from mouth opening, respectively, forming pseudotrochus; arcs and/or tufts of cirri, often set on cushions. Apical field small. Circumapical band present, often interrupted; cingulum not differentiated, may form paracingulum with posterior rows of buccal cilia. In planktonic and semi-planktonic species (Euchlanidae, Epiphanidae, Brachionidae). Conochilus type. Trochus and cingulum well developed, usually interrupted ventrally. Mouth displaced to the dorsal side. In free-swimming species (Conochilidae).

Fig. 4.8: Types of corona: (A–K) Monogononta, (L, M) Bdelloidea. (A) Notommata, ventral, (B) Notommata, lateral, (C) Cyrtonia, (D) Euchlanis, (E) Epiphanes, (F) Brachionus, (G) Synchaeta, (H) Asplanchna, (I) Hexarthra, (J) Conochilus, (K) Floscularia, (L) Macrotrachela, ventral, and (M) Macrotrachela, dorsal. Large dots indicate trochus, medium dots cingulum, small dots ciliation of circumapical band. Abbreviations: af, apical field; bf, buccal field; ca, circumapical band; m, mouth. (From de Beauchamp (1907, 1965.)

224 

 4 Rotifera

Fig. 4.9: SEM picture of the shape of the corona in different genera of Monogononta: (A–C) Notommata, (D, G) Dicranophorus, (E) Asplanchna, (F) Euchlanis, (H, I) Cyrtonia, (J) Brachionus, (K, L) Conochilus, (M, N) Hexarthra, and (O) Floscularia. Scale bar = 10 µm (A, D, F, I, K, L, N), 20 µm (B, C, E, G, H, J, M, O). (Photo courtesy of Giulio Melone.)

Hexarthra type. Buccal field small. Circumapical band and apical field large; strongly developed trochus and weaker cingulum, not interrupted ventrally. In sessile (Flosculariidae), semi-planktonic (Testudinellidae), and free-swimming species (Hexarthridae). Collotheca type. Corona extended into 2–8 blunt or elongate lobes, forming funnel with mouth at center of the bottom. Edges of lobes with motionless cilia or setae or lobes apically with bundles of long setae. In sessile and planktonic species (Collothecidae). The corona of adult females of Atrochidae (Acyclus, Atrochus, Cupelopagis) lacks cilia or setae; a ciliated

corona, used for swimming, is only present in their juveniles and males. Corona types in Bdelloidea (see de Beauchamp 1907a, 1908, 1965, Remane 1933, Melone & Ricci 1995, Ricci & Melone 1998a) (Figs. 4.8 L–M and 4.10) Philodina type. Ventral mouth opening surrounded by buccal field. Buccal field anteriorly delimited by 2 individual retractable trochal disks elevated on pedicels, lined by C-shaped wreath of long cilia, the trochi, and posteriorly demarcated by a single wreath of long cilia, the cingulum. Each trochus is made of about 30 rows of

4.2 Morphology 

Fig. 4.10: SEM picture of the shape of the corona in 2 genera of Bdelloidea: (A) Adineta and (B) Dissotrocha. Scale bar = 10 µm. (Photo courtesy of Giulio Melone.)

 225

Cingulum and cingulum pad with whiskers bordering mouth opening. Corona used for swimming and capturing food; both actions possible at the same moment. In Philodinidae and Habrotrochidae. Abrochtha type. Similar to Philodina type, but with an unpaired trochus composed of a unique row of cilia, and encompassing the rudimentary pedicels and V-shaped cingulum; cingulum pad and whiskers bordering mouth opening lacking. Locomotion and capturing food are 2 separate actions. In Abrochtha and Philodinavus. Adineta type. The corona is a homogenous ventral field of undifferentiated cilia: trochus, cingulum, and cingulum whiskers absent; cingulum pad present. Posterior border of ciliated field with paired cuticular denticulate structures, the food rake. Adineta is unable to swim or to create water currents: the corona is used only for gliding and feeding by scraping the substrate; food is collected with the rake. In Adinetidae.

4.2.1.4 Morphology of the male

Fig. 4.11: Male of Brachionus calyciflorus. Abbreviations: b. brain with eyespot; da, dorsal antenna; eg, “excretion” granules; f, foot; p, penis; pg, pedal glands; pn, protonephrial apparatus; prg, prostate glands; ri, rudiments of intestinal system; sb, sensory bristles; to, toes; t, testis; vd, vas deferens. (Modified from de Beauchamp 1965.)

cilia. The cilia of the cingulum insert on a cingulum pad. Between the pedicels, the epidermis forms an upper lip; an epidermal lower lip lines the mouth opening ventrally.

Males (Fig. 4.11) are usually much smaller than the females and of simpler structure (Wesenberg-Lund 1923, Remane 1933). The sexual dimorphism (Fig. 4.12) is almost a continuum ranging from very weak to very pronounced. Species of Ploima with males of similar appearance to the female and displaying a well developed head, trunk, foot, and toes (e.g., Eosphora najas, Pourriotia werneckii, formerly Proales werneckii, Rhinoglena frontalis, Notommata copeus) are usually slightly smaller (70%–90% of the length of the female) and/or slender, but often have a reduced corona. Melone (2001) studied the coronae of both sexes in R. frontalis by SEM and showed that they are organized in the same way, but the corona of the male is smaller by a reduction of the number of cilia present in its different parts. Other differences concerned the relatively longer foot, proboscis, and lateral antennae. In species with extremely reduced males (e.g., Polyarthra, Trichocerca), the length of the males is only about 15%–50% of the female, and the body is merely a conical or sac-shaped structure without foot, bearing a strongly reduced corona consisting of a terminal ciliated field or tufts of cilia. Spines and appendages of the body and lorica found in the female are lacking or reduced, e.g., the male of Hexarthra only has 3 reduced arms, 1 dorsal and 2 lateral ones, instead of the 6 arms in the female. In some species displaying strong sexual dimorphism, males may show characters absent in females (e.g., Remane 1933, Segers &

226 

 4 Rotifera

Fig. 4.12: Sexual dimorphism in Monogononta: (left) female and (right) male. (A) Rhinoglena frontalis, (B) Notommata copeus, (C) Eosphora najas, (D) Cephalodella catellina, (E) Hexarthra mira, (F) Testudinella patina, (G) Wierzejskiella velox, (H) Lecane psammophila, (I) Collotheca ornata, (J) Brachionus urceolaris, (K) Scaridium longicaudum, (L) Colurella uncinata, (M) Trichocerca pediculus, and (N) Trichocerca capucina. (From different authors.)

Rico-Martinez 2000): the males of Lecane bulla and Synchaeta triophthalma have 2 separate toes instead of the single toe in the female; the neck region in the male of Pompholyx sulcata consists of several plates that are absent in the female; in male Brachionidae, the neck is stiffened and the trunk often shows several plates.

In the sessile Flosculariidae and Collothecacea, the length differences between the sexes may be very great, the males only reaching 5%–10% of the length of the females, and resembling the free-swimming juvenile females. They often show a conical projection at the anterior end, bearing apical eyes, which is absent in the

4.2 Morphology 

 227

4.2.2 Integument

the ICL is very thin in the apical region and thicker and more dense at the level of the trunk, foot, and spines, and thin and flexible at the articulation between the trunk and the corona and at the other joints of, e.g., the foot, toes, paddles, and movable spines (e.g., Clément 1987, Clément & Wurdak 1991). The stiffening can also be restricted to separate plates, e.g., a dorsal and ventral trunk plate in Euchlanis, or 3–5 trunk plates in Cephalodella. The plates are separated by a thin flexible ICL, with the integument taking the shape of a more or less deep invagination or sulcus between the plates. Species showing an extremely thick and stiffened ICL of the major body regions, hence becoming almost inflexible are called loricates (e.g., Brachionidae, Mytilinidae, Trichotriidae) and those with a thin and flexible ICL throughout are illoricates or semi-loricates

The integument is syncytial, and besides its function as an outer protective covering, it plays a role as peripheral skeleton, serves for muscle attachment, and has an endocrine and exocrine function. The skeleton function is provided by a dense intracytoplasmic lamina (ICL) (e.g., Clément 1969, Koehler 1965b, 1966, Dickson & Mercer 1967, Schramm 1978a, Clément & Wurdak 1991, Kleinow 1993), composed of 2 filamentous keratin-like proteins (molecular weight, 39 and 47 kDa, respectively), cross-linked by disulfide bonds (Bender & Kleinow 1988). The ICL lies just under the outer cell membrane of the syncytial integument and is regularly perforated by pores (Figs. 4.13 and 4.14). The cell membrane invaginates through these pores forming spherical bulbs within the cytoplasm. Clément (1969, 1980, 1985, 1993) recognizes 4 types of ICL according to the absence or presence of different layers, and the vertical or horizontal orientation of their composing structures. The bdelloid type ICL shows a thickened and uniformly electron-dense internal layer and a very thin external layer (Philodina acuticornis odiosa, P. roseola, Rotaria sp., Habrotrocha rosa). In monogononts, 3 types may be recognized, characterized by a thick external layer, and a thin or lacking electron-dense internal layer. The stacked lamella type shows several fibrous horizontal layers: 7 in Notommata and 3 in Asplanchna and Synchaeta; the internal layer is strongly reduced. The vertical tube type consists of a layer of juxtaposed vertical tubes and a welldeveloped thin internal layer (Brachionus, Mytilina). The uniformly dense type shows a dense homogenous layer that may be thick (Trichocerca, Keratella) or thin (Filinia, Sinantherina); the internal layer is absent. The thickness of the ICL varies greatly within a single rotifer and between different species. Within the animal,

Fig. 4.13: Diagram of the 4 types of intracytoplasmic lamina. (A) Philodina type, (B) Brachionus type, (C) Notommata type, and (D) Trichocerca-Filinia-Keratella type. Abbreviations: el, external lamina; g, glycocalyx; il, internal lamina; m, membrane; sb, secretory bulb; sc, syncytial cytoplasm. (Modified from Clément 1993.)

adult females; a foot with adhesive disc may be present (­Lacinularia). Males of most benthic monogononts show little or medium reduction in structure and length, unlike the ones of planktonic species, which are the most extremely reduced (Wesenberg-Lund 1923, Remane 1933, Riemann & Kieneke 2008). According to Ricci & Melone (1998b), this phenomenon suggests that the species in the 2 habitats are exposed to different selection pressures. Serra & Snell (1998) suggested that male dwarfism in rotifers could be the result of selection on females, to produce a high number of males in a short period with a minimum reproductive cost.

228 

 4 Rotifera

Fig. 4.14: TEM section of the integument of Macrotrachela quadricornifera. The ICL is visible, with several layers. The section of a pore crossing the lamina is also visible. (Photo courtesy of Giulio Melone.)

(e.g., Epiphanidae, Asplanchnidae, Lindiidae). All stages from illoricate to strongly loricate may occur within a single genus or family; therefore, the degree of stiffness of the ICL generally is of little taxonomic significance. Secretions produced by the syncytial integument are expelled to the outside through secretory bulbs and pores traversing the ICL (e.g., Clément 1977, Clément & Wurdak 1991). These secretions form a very thin to thick extracellular “cuticle”, or glycocalyx, to which foreign particles may stick. Among the secreted substances are glycoproteins that play a role in male mate choice, discriminating females based on species, sex, age, and reproductive status (e.g., Snell et  al. 1995). The integument also has an endocrine function, discharging its secretion products in the pseudocoel via secretory granules produced by the rough endoplasmic reticulum and Golgi apparatus (Clément 1977, Clément & Wurdak 1991). The ICL also serves for the attachment of the body muscles and the cutaneovisceral muscles. Associated with the integument are several glands: pedal glands, the retrocerebral organ, ovifer glands to fix and carry eggs (Sinantherina), modulus glands in tube-forming Flosculariacea, etc.

part of the trunk; smaller accessory pedal glands with an own aperture or an aperture in common with the main glands may be present. In bdelloids, from 2 to 15 pairs of mononucleate pedal glands can be found (Fig. 4.15). The ducts of the pedal glands may be widening locally into reservoirs. The secretion is discharged via apertures in the attachment organs at the end of the foot. In species with reduced or vestigial foot, pedal glands are usually lacking. The apertures of the pedal glands lie on or near the tip of the toes in most Ploima; in Trichocerca, they issue at the base of the toes and the secretion flows externally along the toe(s). In Flosculariacea and Collothecacea, the toes open at the adhesive disc or at the end of the foot. In bdelloids, the tips of both toes and spurs have openings for the ducts of the pedal glands; in the adhesive discs, the pedal glands issue by numerous openings (e.g., up to 14 in Habrotrocha proxima). In monogononts, only one type of secretion is produced by the same animal, whereas in bdelloids, the same animal produces 2 types of secretion (Clément 1987, Clément & Wurdak 1991).

4.2.2.1 Pedal glands

The retrocerebral organ (Figs. 4.5 and 4.16) is located dorsal to the brain and mastax and consists of the unpaired median retrocerebral sac and 2 lateral subcerebral glands (de Beauchamp 1905, 1906, 1965). From the anterior part of the sac, an unpaired duct runs toward the corona and bifurcates anteriorly into 2 ducts opening on the apical field; the openings often lie on a single or paired papilla. The subcerebral glands have ducts alongside those of the sac. In monogononts, the content of the sac often appears strongly vacuolated and may contain light refracting bodies

The foot contains the pedal glands and their reservoirs and ducts. They are unicellular or syncytial, and secrete cement to attach the animal on the substrate temporarily (Ploima, Bdelloidea) or permanently (sessile Flosculariacea and Collothecacea). In some species, the secretion of the pedal glands is also used to anchor eggs to the substrate. Monogononts generally have paired multinucleate pedal glands lying in the foot or extending into in the distal

4.2.2.2 Retrocerebral organ

4.2 Morphology 

 229

Fig. 4.15: Variation of the pedal glands in Bdelloidea. (A) Rotaria socialis, (B) Zelinkiella synaptae, (C) Mniobia tetraodon, (D) Embata parasitica, (E) Mniobia armata, (F) M. russeola, (G) Philodina vorax, and (H) Adineta barbata. Abbreviations: a, adhesive disc; s, spur; t, toe. (From Kutikova 2005.)

Fig. 4.16: Types of retrocerebral organs in Monogononta (A–D) and Bdelloidea (E, F). (A) Notommata copeus, (B) Notommata aurita, (C) Itura myersi, (D) Erignatha clastopis, (E) Rotaria socialis, and (F) Embata parasitica. Abbreviations: b, brain; rs, retrocerebral sac; sg, subcerebral glands. (Modified from Brakenhoff 1937 and de Beauchamp 1965.)

and red-colored pigment granules. The s­ ubcerebral glands are granular and often contain a light refracting globule in Encentrum. The size of sac and glands, and their contents, is highly variable: there may be a sac without glands or glands without a sac (Fig. 4.16). Variations of the subcerebral glands may be apparent also in different populations of the same species. In Ploima, the retrocerebral organ is mostly well developed, but lacking in some species; it is reduced or absent in Flosculariacea and Collothecacea. In bdelloids, the retrocerebral organ is especially developed in creeping species (Brakenhoff 1937), and the different glands produce only one type of secretion (Clément & Wurdak 1991). Ultrastructural investigations show that the ­retrocerebral organ is made up of multinucleate mucous glands, and surrounded by a muscular sheath at its base. The muscle has long thick filaments surrounded by thin filaments. The ratio of thick to thin filaments is 1:6, which is characteristic of a slow and tonic contraction. The ducts are lined by longitudinal microtubules lying below the cell membrane and assisting in

the discharge of mucous secretions (Clément 1977, 1980, Clément & Wurdak 1991). The function of the retrocerebral apparatus is not well known. According to Clément & Wurdak (1991), it probably lubricates the cilia of the corona involved in creeping. In bdelloids, it may be responsible also for the adhesive attachment of the rostrum when creeping (Brakenhoff 1937). Clément et al. (1983) also noticed that the female of Notommata copeus envelopes new laid eggs with mucus secretions from the pedal glands and the retrocerebral organ. The retrocerebral organ is also present in male rotifers (e.g., Wesenberg-Lund 1923, Riemann & Kieneke 2008) and apparently does not show sexual dimorphism.

4.2.2.3 Sheaths and tubes Many rotifers, especially bdelloids (see Donner 1950) and sessile and planktonic monogononts (e.g., Remane 1933, Edmondson 1945, Wright 1950, Fontaneto et  al. 2003),

230 

 4 Rotifera

secrete or construct sheaths or tubes formed of mucus (e.g., Conochilus spp., Gastropus stylifer, Lacinularia spp., Trichocerca cylindrica), mucus and cemented detritus or fecal material (e.g., Habrotrocha gracilis, H. pavida, H. pusilla), gelatinous material (e.g., Acyclus, Collotheca, Ptygura, Stephanoceros), pellets made of bacteria and detritus (Floscularia ringens), fecal pellets (F. janus, Ptygura pilula), and rigid, often ringed material (Habrotrocha angusticollis, Limnias melicerta). The tubes may enclose the whole animal or the foot only. Eggs are laid inside. Some benthic-periphytic monogononts secrete close-fitting mucous coats, often containing foreign particles (e.g., Encentrum umbonatum, Notommata copeus, Paradicranophorus hudsoni, Tetrasiphon). Cephalodella forficula starts its life as a more or less planktonic form before constructing long (up to 5 mm), substrate-bound and detritus-covered mucous tubes, often closed at both ends, in which it swims around (Dodson 1984). Several bdelloids have an outer covering of secretion shaped as

stiff platelets (e.g., Mniobia incrassata, M. mirabilis) or thin rods (e.g., Rotaria socialis). The mucous or gelatinous tubes appear to be secreted by the general body surface in Ptygura and bdelloids or glands near the mastax in bdelloids (Edmondson 1945, Donner 1950, Wulfert 1969) or they are formed by the pedal glands in, e.g., Cephalodella forficula, Collotheca campanulata, and Limnias melicerta (Dodson 1984, Wulfert 1969). The pellets composing the tube of F. ringens are formed in a ciliated groove below the lower lip, the modulus, by collecting and concentrating bacteria and detritus, and gluing them together with sticky secretion from the modulus glands (Fontaneto et al. 2003). For details on tube construction in Collotheca campanulata, F. ringens, F. janus, Limnias melicerta and Stephanoceros fimbriatus, see, e.g., Edmondson (1945), Wright (1950), Wulfert (1969), de Beauchamp (1965), Fontaneto et al. (2003), and Wallace et al. (2006). Sheaths and tubes may protect against predation and drying out and enhance buoyancy in planktonic species.

Fig. 4.17: CLSM of musculature. (A) Adineta ricciae, (B) Brachionus manjavacas female, and (C) Brachionus manjavacas male. Scale bar = 20 µm (A) and 50 µm (B, C). (Photo courtesy of Francesca Leasi.)

4.2 Morphology 

4.2.3 Musculature 4.2.3.1 Musculature of the female The muscles of rotifers comprise smooth, cross-striated, or obliquely striated types, which can be monocellular or bicellular, phasic or tonic, strong or weak, and endurant or not (e.g., Amsellem & Clément 1977, Clément & Amsellem 1989), and occurring as thin filaments or arranged in separate small bands. They are organized into somatic muscles, controlling movement and changes of body form, and serving the head, foot, dorsal antenna, etc., the cutaneovisceral muscles connecting the internal organs with the integument, and the visceral muscles surrounding the digestive tract, cloaca, and reproductive apparatus. The arrangement of the somatic component, studied by LM and histological techniques, received much attention in the past (e.g., Zelinka 1886, Martini 1912, Nachtwey 1925, Remane 1933, Seehaus 1930, Peters 1931, Stossberg 1932, Dehl 1934, Brakenhoff 1937). The results of these earlier observations were largely corroborated and refined by epifluorescence and CLSM (Fig. 4.17) of the phalloidin stained F-actin fibers (e.g., Kotikova et  al. 2001, 2004, 2006, Santo et  al. 2005, Sørensen et  al. 2003, Sørensen 2005a, b, Hochberg & Ablak Gurbuz 2008, Hochberg et  al., 2008, Riemann et  al. 2008, Leasi & Ricci 2009, Wilts et al. 2009, Wilts & Ahlrichs 2010, Leasi et al. 2010). ­Comparison of the somatic muscular system among all rotifer taxa investigated to date reveals a common basic

 231

pattern (e.g., Santo et al. 2005, Leasi et al. 2012), consisting of 2 major groups: a system of outer circular bands and a system of inner, paired, and bilaterally symmetrical longitudinal bands (Figs. 4.18 and 4.19). Both circular and longitudinal muscles insert at their endings on the skeletal syncytial integument and sometimes at the level of muscle Z-elements (Clément & Amsellem 1989). Musculoepithelial junctions are by desmosomes, joined to the ICL by dense material; musculomuscular junctions are by desmosomes and hemidesmosomes and gap junctions (e.g., Koehler 1965b, Clément 1969, Clément & Amsellem 1989). Somatic muscles of bdelloids are smooth or obliquely striated, whereas in monogononts, cross-striated muscles preponderate (Clément & Amsellem 1989, Clément 1993). Circular muscles are localized in the head, trunk, and junction between trunk and foot. Strongly developed circular muscles in the head directly behind the corona form the coronal sphincter, able to contract tightly over the head after retraction of the corona, and thereby protecting the latter (Clément & Amsellem 1989). The circular muscles of the trunk comprise several, mostly 3 to 7, separate muscles. Upon contraction, they exert pressure on the turgid pseudocoel, which results in the extension of the body. There is a great deal of variation in the circular muscle pattern of the trunk, supposed to reflect the ecology, mode of locomotion, shape of the body, and presence of integumentary plates (e.g., Remane 1933, Leasi & Ricci 2009). The circular muscle system may consist of closed muscular rings (Asplanchnopus

Fig. 4.18: Diagram of musculature in Brachionus manjavacas. (A) Female, ventral view, (B) female, lateral view, and (C) male, lateral view. Abbreviations: cs, corona sphincter; dm, dorsal longitudinal muscle; dvm, dorsoventral muscle; lm, lateral longitudinal muscle; pc, muscle pars coronalis; sm, splanchnic longitudinal musculature of the penis; vm, ventral longitudinal muscle. (From Leasi et al. 2010.)

232 

 4 Rotifera

Fig. 4.19: Diagram of somatic musculature in Adineta ricciae (A, B) and Macrotrachela quadricornifera (C, D). (A, C) Dorsal view and (B, D) ventral view; Abbreviations: am, musculature of antenna; cm, circular muscles (1–13); dl, dorsal longitudinal muscles; hl, dorsal longitudinal muscles of the head; hr, ring muscle of the head; rl, longitudinal muscles of the rostrum; tm, musculature of the trochi; vl, ventral longitudinal muscles. (From Leasi & Ricci 2009.)

multiceps, Notommata glyphura) or semi-circular, and open configurations just beneath the integument. The interruptions of the semi-circular muscles may be ventral (e.g., Adineta ricciae, Macrotrachela quadricornifera, Encentrum mucronatum, Epiphanes senta, Proales fallaciosa, Floscularia ringens) or both dorsal and ventral (Proales reinhardti), or the muscles may be split into paired dorsolateral and paired ventrolateral bands connecting the different integumentary plates of the lorica (Dicranophorus forcipatus). In some species, the circular muscles are modified into strands that became free from the integument and only connected to it by their endings (e.g., Testudinella, Euchlanis). Complete or ventrally open muscle rings are mostly characteristic of soft-bodied monogononts and bdelloids. In loricates or semi-loricates, with stiffened integumentary plates, the circular muscles are generally less developed. They are reduced to short lateral dorsoventral strands connecting the dorsal and ventral parts of the tegument or lorica plates in dorsoventrally compressed species (e.g., Brachionus spp., Euchlanis dilatata, Testudinella patina). Transversal muscle bands are found in laterally compressed species (e.g., Colurella, Mytilina) and oblique lateroventral strands are present in species with triangular cross section (e.g., Mytilina). Circular muscles along the trunk are absent in species possessing movable appendages like spines (e.g., Filinia novaezealandiae), arms (e.g., Hexarthra mira) or paddles (e.g., Polyarthra major), but it has been suggested that some muscles serving these appendages are derived from the

circular muscles (e.g., Levander 1894, Hochberg & Ablak Gurbuz 2007, 2008). A pedal sphincter is often present at the junction of the trunk and foot, and a circumpedalis muscle may encircle the junction between foot and toes. The inner, longitudinal retractor muscles are present in the head, trunk, and foot (if present) and insert at different points of the integument; some may span the length of the animal inserting anteriorly in the head or neck region and posteriorly at the base of the trunk. They usually occupy a dorsal, lateral, and ventral position and permit bending of the body and withdrawal of head and foot into the trunk. In bdelloids, the longitudinal muscles are responsible also for the contraction into a tun-shaped stage during desiccation. The different insertion points of the respective head and foot retractors allow the muscles to withdraw these extremities independently. The number of longitudinal muscles as well as their length, width, and insertion points is variable between different taxa and even between closely related species. The longitudinal muscle pattern seems to be determined by different modes of locomotion and adaptation to a specific ecological niche. Major anatomical changes in the somatic muscular system do not take place during the larval development and metamorphosis to the adult stage in Acyclus inquietus, a rotifer with free-swimming larvae and sessile adult females (Hochberg et al. 2010). Information on the cutaneovisceral muscle system is restricted (e.g., Remane 1933, Kotikova et  al. 2006). The number of muscles and position appears very varia-

4.2 Morphology 

ble, even within a single genus. They hold the organs in place, and are especially common at the level of mastax and cloaca. Others have a more specific function, e.g., the dilatator muscles of the mouth in raptorial species (Asplanchna, Synchaeta) allow for the swift opening of the mouth to engulf the prey. Very little information is available on the visceral musculature (e.g., Remane 1933, Kotikova et  al. 2006, Riemann et  al. 2008, Wilts et  al. 2009, Wilts & Ahlrichs 2010, Leasi et al. 2010). The corona, mastax, esophagus, salivary glands, stomach, intestine, cloaca, oviduct, retrocerebral organ, pedal glands and their reservoirs, and protonephridia may be provided with muscle cells or syncytial muscle fibers. Reticulate, circular, and longitudinal arrangements occur.

4.2.3.2 Musculature of the male Little is known on the musculature of the male. The figures presented by Wesenberg-Lund (1923) should be interpreted cautiously, but circular and central, dorsal, and ventral longitudinal retractor muscles are definitely present in many males (Remane 1933, de Beauchamp 1965). Waniczek (1930) found no fundamental differences in body musculature between the sexes in Asplanchna. Comparison of the muscle organization by CLSM of the females and males of Brachionus manjavacas (Fig. 4.18 C) and Epiphanes senta, with different ecology and stiffness of the lorica, reveals an identical condition independent of their ecology and morphology, suggesting that evolution and development constrains the organization of the male muscular system (Leasi et al. 2010). In B. manjavacas and E. senta visceral muscles are present in the reproductive apparatus but absent in the gastrointestinal system, which could be expected because the latter is lacking in these species (Leasi et al. 2010). The reproductive apparatus is supplied by several muscles in longitudinal and circular (B. manjavacas) as well as transversal arrangements (E. senta).

 233

techniques (e.g., Nogradi & Alai 1983, Raineri 1984, Keshmirian & Nogradi 1987, 1988, Kotikova 1995, 1998). Immunochemistry and CLSM (e.g., Hochberg 2006, 2007, 2009, Leasi et  al. 2009) demonstrated the presence of, among others, cholinergic, adrenergic, catecholaminergic, dopaminergic, and serotonergic systems. The nervous system of only few species has been studied. The neuronal organization is fairly conservative in rotifers (e.g., Kotikova 1995, 1998, Hochberg 2006) and shows a basically bilateral symmetry (Fig. 4.20). It consists of a large cerebral ganglion, commonly called brain, located behind the corona dorsally to the mastax, and surrounded by either epithelial or muscular cells. The brain is of varying shape and size: rounded, sacshaped, quadrangular, or triangular, etc. The number of cells is constant for each species (e.g., 183 in Epiphanes senta and 249 in Synchaeta tavina). The neuropilar core of Philodina roseola and Trichocerca rattus is central and surrounded by perikarya at its periphery (Clément 1977, Clément & Wurdak 1991). Perikarya and associated neurites, display a strong symmetry in number, size,

4.2.4 Nervous system Information on the nervous system goes back to the LM and histological observations by, e.g., Zelinka (1888, 1891), Hlava (1905), Hirschfelder (1910), Martini (1912), Nachtwey (1925), Seehaus (1930), Peters (1931), Stossberg (1932), Remane (1933), and Dehl (1934). Additional details were revealed by TEM (e.g., Eakin & Westfall 1965, Clément 1977, Clément et al. 1991) and application of histochemical

Fig. 4.20: Diagram of the nervous system of rotifers. (A) Lateral and (B) ventral. Dark grey, nervous system; light grey, intestinal system, bladder, and pedal glands; medium grey, muscles. Abbreviations: as, apical sense organs; b, brain; ca, caudal antenna; cg, caudal ganglion; da, dorsal antenna; eg, epipharyngeal ganglion; gg, geniculate ganglion; la, lateral antenna; mg, mastax ganglion; nl, nerve to lateral antenna; sn, scalar nerve; so, supra-anal sense organ; vn, ventral nerve; vg, vesicular ganglion; vs, visceral nerve. (Modified from Remane 1933.)

234 

 4 Rotifera

connections, and pathways between the cerebral hemispheres (e.g., Hochberg 2007, 2009). Two ventral main nerves, composed of a bundle of axons, proceed from the sides of the brain to the caudal or foot ganglion, branching of lateral nerves at secondary ganglia. The coronal region with its sensory structures and the dorsal antenna are innervated by a series of paired neurites issuing from the brain that may form neuronal rings (Kotikova 1998, Hochberg & Lilley 2010). The brain also sends nerves to the mastax, salivary glands, and the dorsal, lateral, and ventral retractor muscles. Neurites branching off from the ventral main nerves supply the dorsal, lateral, and ventral retractor muscles, internal organs, and lateral antennae. A pair of perikarya in the foot sends neurites to the caudal ganglion; an unpaired neurite connects the caudal antenna with the caudal ganglion. A transverse commissure between the lateral longitudinal neurites in the head or between the main nerve cords in the anterior and posterior regions of the trunk may be present. In males, brain, nerves, and sensory organs appear well developed, but the mastax ganglion is usually absent in accordance with the reduced non-functional mastax (Remane 1933). In Brachionus plicatilis, Keshmirian & Nogrady (1988) demonstrated catecholaminergic innervation of all major organs and the male copulatory apparatus.

4.2.5 Sensory structures Rotifers have a great variety of sensory cells and sensory organs, which can be classified as mechanoreceptors, photoreceptors, and chemoreceptors. However, many of them are sensory complexes having multiple functions; the function of several others is unknown or conjectural (e.g., Remane 1933, de Beauchamp 1965, Clément et  al.

1983, Wurdak et  al. 1983, Clément & Wurdak 1991). The presence and number of sensory structures vary greatly, even within a single genus or family (for a review, see Remane 1933). The apical field and cingulum of the corona of monogononts are frequently provided with sensory bristles (styles or cirri, and membranelles), sensory papillae, chemoreceptive pores, sensory pits, etc., emanating from a single or a small number of nerve endings (Fig. 4.21). The sensory bristles apparently have a tactile function and participate in sensing water movement and contact with conspecifics and/or prey and male mate choice (e.g., Remane 1933, Clément et  al. 1983, 1991, Snell et  al. 1995, Joanidopoulos & Marwan 1998). They are composed of several cilia, the axoneme of which contains the characteristic 9×2+2 tubules. The sensory endings of their nerve fibers are usually surrounded by a supporting epithelial cell, which is protruding into the pseudocoelom. Sensory papillae, with or without apical cilia, are often present in the apical field as well (Fig. 4.22). Each of the 2 palps of Trichocerca rattus, known as palpar organ, shows 6 symmetrically arranged nerve fibers enclosed at their base by 2 supporting epithelial cells. Within the palps are nerves terminating in microvilli and cilia containing the 9 peripheral doublets but lacking the 2 central tubules. Clément et al. (1983) suggest that the palpar organ is responsible for the recognition of the filamentous shape of the algae they feed on. The finger- to sickle-shaped palp in the apical field of Ascomorpha ovalis and A. saltans serves to hold the prey while its contents are being extracted. In several Dicranophorus species, the head bears 2 to 6 anterolateral palps, supposed to be tactile organs. Chemoreceptive pores located in the anterior syncytial integument beneath the cingulum have been described in Brachionus and Notommata (Clément et  al. 1983,

Fig. 4.21: Head of Synchaeta with antennae, sensory styles and brain. Abbreviations: a, auricle; b, brain; c, dorsolateral part of corona; ca. secondary coronal antenna; da, dorsal antenna; ds, dorsal coronal seta; e, cerebral eyespot; fa. frontal coronal antenna; ls, lateral coronal seta; st, dorsolateral sensory cell of trunk. (Modified from Kutikova 1970.)

4.2 Morphology 

Fig. 4.22: Palps and sensory organs on apical field. (A) Ascomorpha ovalis, (B) Trichocerca stylata, (C) Trichocerca capucina, and (D) Pleurotrocha petromyzon. Abbreviations: at, apical tentacle; bf, buccal field; ca, circumapical band; cg, ciliated groove; da, dorsal antenna; dp, dorsal palp; mp, median palp; om, oral membranel. (A, C, D, From Remane 1933; B, from de Beauchamp 1965.)

Clément & Wurdak 1991). Non-ciliary sensory endings in the form of microvilli are present inside the lumen underlying the pore (Brachionus calyciflorus, B. sericus) or the pore continues internally through a muff-shaped underlying epithelial cell, the cavity of which contains cilia and microvilli borne by a sensory nerve ending (Brachionus plicatilis, Notommata copeus). Ciliated pits with chemoreceptor function and resembling chemoreceptive pores consist of an external cavity enclosing epithelial cilia, and an internal cavity lined with specialized cilia and sensory membranes (Clément et al. 1983, Wurdak et al. 1983). There are paired structures located on the apical field in, e.g., Asplanchna. The prominent sensory organs of the body are mostly the dorsal and lateral antennae and to a lesser extent the caudal antenna. The dorsal antenna is found in all taxonomic groups of rotifer and generally situated medially on the head or neck but displaced to the anterior part of the trunk in Testudinella. In monogononts, it is a more or less long cylindrical projection bearing tufts of cilia or consists of tufts of cilia or styles projecting through a pore in the integument; occasionally, it is reduced to an unciliated pit. The dorsal antenna is paired in the embryonic stage, and becomes fused into an unpaired organ in most of the full-grown species; it is rarely absent (e.g., Conochilus). It consists of 2 cerebral nerve fibers with ciliated ends supported by an epithelial cell, which forms a subsurface pocket containing the bases of the sensory cilia (Fig. 4.23 B). The dorsal antenna is most prominent and of greater complexity in bdelloids, where it forms a long tube, telescopically retractable by paired muscles cells contained within it. The dorsal antenna of bdelloids has 3 pairs of nerve endings and several epithelial supporting cells (Clément 1977, Clément & Wurdak 1991).

 235

Fig. 4.23: Lateral (A) and dorsal antenna (B) in Trichocerca rattus. Abbreviations: b, brain; cn, cerebral neuron terminating at multinucleate, symmetrically arranged supporting cell (sc); i, integument; n, simple neuron in direct contact with integument; p, pseudocoelom. (Modified from Clément & Wurdak 1991, with permission.)

Paired lateral antennae (Fig. 4.24) are present in monogononts, but lacking in bdelloids. They are usually situated laterally and symmetrically in the posterior half of the trunk but may be displaced ventrally or dorsally, or more anteriorly up to the apical field in Conochilus; in Hexarthra, they are situated on the ventral arms; they rarely lie on the caudal lorica spines (e.g., Plationus patulus macracanthus). In numerous Trichocercidae, the lateral antennae are in very asymmetrical positions. Partial to complete fusion of the apically displaced antennae occurs in some Conochilus. The lateral antennae are of similar shape as the dorsal antenna, but usually smaller (Fig. 4.23 A). They consist of a single sensory neuron in direct contact with the integument, with which it forms a pocket containing the bases of the sensory cilia (Clément 1977, Clément & Wurdak 1991). A single, rarely paired, small dorsal caudal antenna, consisting of a ciliated pit or a more or less shallow projection tipped with a tuft of cilia, is generally present on the distal foot pseudosegment, between or above the base of the toes in Ploima (Fig. 4.25). In some species (e.g., Notommata, Proales), there is an unciliated papilla between the toes, and the distal foot pseudosegment of Collurellidae and Cotylegaleata bears a dorsal sensory pit, apparently lacking cilia. Photoreceptors, called eyes, eyespots, or ocelli, are commonly present in monogononts and bdelloids, which most of the species retaining the eyes throughout their life. Eyespots are often lacking in adults of sessile orders, but present in their free-swimming juvenile stages (e.g., Hlava 1908, Hochberg et  al. 2010). Eyespots contain a red to black pigment; the intensity of the color largely

236 

 4 Rotifera

Fig. 4.24: Forms of lateral antennae in Monogononta. (A) Basic type, (B) Lecane ligona, (C) Synchaeta tavina, (D) Tetrasiphon hydrocora, (E) Flosculariidae, scheme, and (F) Conochilus unicornis. (From Remane 1933.)

depends on the food, i.e., the carotenoids assimilated (Birky 1964). Light-refracting globules without pigment have been referred to colorless eyes (e.g., Encentrum) and may represent photoreceptive structures that lack a pigment cup containing screening pigment. The eyes are usually situated frontally or dorsally (rarely ventrally, e.g., Hexarthra) on the apical field, rostrum, and lateral sides of the corona or on the brain. According their specific location, they are called apical, frontal, rostral, or lateral eyespots, etc., and cerebral eyes. Cerebral eyes lie on the brain and are mostly single or sometimes paired but often fused (e.g., Notommatidae, Synchaetidae), unlike the other eye types, which are usually paired. Some species show a combination of frontal, lateral, and cerebral eyespots, e.g., Eosphora najas has 5 eyespots. The ultrastructure of the ocelli and cerebral eyes has been studied by, e.g., Clément et al. (1983) and Clément & Wurdak (1991). The frontal eyespots of Rhinoglena frontalis and apical eyespots of Filinia longiseta consist of anterior ciliated cells forming a cup with screen pigment, and enclosing a nerve ending from which microvilli containing the photosensitive pigment emanate. The paired lateral eyes of Asplanchna brightwellii and the paired median ones of Trichocerca rattus show sensory nerve endings bearing ampulla-shaped photoreceptive cilia (Fig. 4.26), separated from the outside medium by a thin cuticle. A light-refracting lipid globule in front of the microvilli of the ocelli is present in many species and referred to as lens in the older literature (Remane 1933). The cerebral eye is integrated into the brain and associated with the epithelial cells surrounding the latter. The ultrastructure of the cerebral eyes (Fig. 4.27) of the 5 species studied to date is unique for each of them (Eakin & Westfall 1965, Clément 1980, Clément

Fig. 4.25: Forms of caudal antennae in Monogononta. (A) Trichotria tetractis, (B) Lepadella ovalis, (C) Trichotria pocillum, (D) Euchlanis, and (E) Brachionus. Abbreviations: c, caudal antenna; s, sense organ. (Modified from Remane 1933.)

Fig. 4.26: Ocelli and cerebral eye of Asplanchna brightwellii. (A) General scheme of head, (B) Detail of ocellus, and (C) Detail of ampulla-shaped cilium with cross sections. Abbreviations: b, brain; c, corona; ce, cerebral eye; cm, circular skeletal muscle; cn, cerebral neuron; ec, epithelial socket cell; lm, longitudinla skeletal muscle; o, ocellus; sn, sensory nerve ending with ampulla-shaped cilia; ps, pseudocoel. (From Clément & Wurdak 1991, with permission.)

4.2 Morphology 

 237

Fig. 4.28: Cerebral eye of the bdelloid Philodina roseola. (A) Location of the eye at periphery of brain and (B) detail of ampulla-shaped cilium in longitudinal and cross section. Abbreviations: b, brain, e, epithelial cell with pigment granules; ne, nerve ending with 2 ampulla-shaped cilia. (From Clément & Wurdak 1991, with permission.)

Fig. 4.27: Cerebral eyes of monogonont rotifers. (A) Brachionus calyciflorus, (B) Brachionus plicatilis, (C) Trichocerca rattus, and (D) Asplanchna brightwellii. Abbreviations: ax, axon; cl, cytoplasmic lamella of relay neuron; pc, epithelial cell with pigment cup or cavities containing pigment granules or platelets (p); sc, sensory neuron lacking axon; rn, relay neuron with axon leading to the neuropile of the brain; rsn, relay neuron becoming sensory neuron with stacked lamellae. (From Clément & Wurdak 1991, with permission.)

et  al. 1983, Clément & Wurdak 1991). The monogononts (Asplanchna brightwellii, Brachionus calyciflorus, B. plicatilis, Trichocerca rattus) show an unpaired median epithelial cell containing the red screening pigment composed of pigment granules (Brachionus, Trichocerca) or platelets (Asplanchna). This epithelial cell is cup-shaped (Trichocerca, Asplanchna) or has 2 lateral cavities (Brachionus), which are responsible for its x-shape observed in vivo. The cavities and cup lodge a sensory neuron lacking axons (Brachionus, Trichocerca); they are connected to a single (B. calyciflorus, T. rattus) or 2 (B. plicatilis) relay neurons sending off axons to the brain. In Brachionus, each of the relay neurons sends a single cytoplasmic lamina into the sensory neuron; in Trichocerca, the relay neuron forms stacked lamellae penetrating into the unpaired sensory neuron. The structure of the cerebral eye of Asplanchna consists of a relay neuron that becomes the sensory neuron, forming stacked lamellae in the pigmented epithelial cup; 2 axons penetrate into the brain. A median cerebral eye is absent in bdelloids, but paired ones, located on either side of the brain, are present in, e.g., Philodina, Dissotrocha, Embata, Habrotrocha collaris, and Abrochtha. The ultrastructure of

Fig. 4.29: Phaosome of the bdelloid Philodina roseola. Abbreviations: e, epithelial cell; fc, stacks of flattened cilia; sb, spherical bulb; sn, sensory nerve. (From Clément & Wurdak 1991, with permission.)

the cerebral eye of the only bdelloid studied (Philodina roseola) shows that it consists of a nerve ending bearing 2 ampulla-shaped cilia and lodged in an epithelial cell containing granules of screening pigment (Fig. 4.28). The presumed components for photoreception that contain the visual pigment of the above 5 species are different as well: tubular formations of the endoplasmic reticulum (B. calyciflorus), a stack of plasma and ER membranes (B. plicatilis), stacks of intracytoplasmic plasma membrane (T. rattus), lamellar rhabdomeres (A. brightwellii), and electron-dense ampulla-shaped cilia (P. roseola). A photoreceptive function was also suggested for the phaosome (Fig. 4.29) of the bdelloid Philodina roseola (e.g., Clément 1980, Clément & Wurdak 1991). The unpaired phaosome is located at the base of the rostrum, near the outlets of the retrocerebral apparatus. It consists

238 

 4 Rotifera

of a sensory nerve, ending in a spherical bulb containing a stack of fan-shaped flat cilia bearing lateral membrane expansions. The bulb is supported by an epithelial cell. Other receptors, consisting of ciliated sensory cells, occur in the mouth region, buccal tube, mastax, pseudocoel, penis, cloaca, etc. (Clément et  al. 1983, Clément & Wurdak 1991).

4.2.6 Intestinal system 4.2.6.1 Intestinal system of the female The anatomy of the intestinal system is thoroughly documented in Remane (1933). For studies of its ultrastructure relying on TEM, see, e.g., Clément et al. (1980a, b, c, 1983) and Clément & Wurdak (1991). The digestive system is similar in most bdelloids and monogononts and consists of a mouth, buccal tube, pharynx with mastax, esophagus, stomach, intestine, cloaca, and cloacal aperture or anus. In most species, the gastrointestinal system runs straight from mouth opening to anus but is U-shaped in species with anteriorly displaced anus (e.g., Conochilidae). The mouth commonly opens apically to ventrally in the centre of the buccal field and frequently at the base of a more or less strongly developed funnel (Fig. 4.30). Several sensory receptors, supposed to participate in food intake, are present in the mouth region (Wurdak et al. 1983). The mouth leads to the buccal tube or opens directly into the pharynx. The buccal tube is formed by a simple epithelium, consisting of mushroom-shaped cells with the caps organized in an imbricate way forming the tube wall, and the stalks representing the cell bodies containing nuclei and most of the cytoplasm (Clément et al. 1980b). Sheaths of circular and longitudinal muscles surround the buccal tube (e.g., Clément et al. 1980c). The buccal tube is short and devoid of cilia or absent in raptorial species (e.g., Asplanchna, Dicranophorus) and more or less long and cylindrical and covered with cilia in filter feeders (e.g., Brachionus, Philodina). The buccal cilia are characterized by an electron-dense tip. In many rotifers, the buccal tube ends in a supple myelin-like structure, the buccal velum, separating the buccal lumen from the pharynx or mastax lumen, thus preventing rejection of the ingested food (Clément et al. 1980a, 1983). The buccal epithelium is provided with several sensory receptors (Wurdak et al. 1983). The pharynx is short and usually ciliated but may be lacking in species with trophi that can be thrust out through the mouth opening (e.g., Dicranophoridae). Pharyngeal cilia have a classical 9 × 2 + 2 configuration

Fig. 4.30: Diagram of oral apparatus: mouth, mastax, and esophagus. Abbreviations: ar, anterior coronal mechanoreceptor; b, brain. be, buccal epithelium; bt, buccal tube; bv, buccal velum; c, cilium; cr, chemoreceptor; m, mastax; mg, mastax ganglion; ml, mastax lumen, M1–M6, muscles and their innervations; o, esophagus; p, pseudotrochus; pe, pharyngeal epithelium; sr, sensory receptor; t, trophi. (Modified from Clément & Wurdak 1991, with permission.)

but are surrounded by a double membrane that is an extension of the double membrane around the pharyngeal lumen (Clément & Wurdak 1991). The pharynx leads to the cavity of a muscular masticatory apparatus, the mastax (Figs. 4.30 and 4.31). This cavity occupies the anterior, dorsal, or anterodorsal part of the mastax and opens posteriorly into to the esophagus. The epithelial wall of the mastax cavity is lined with a thin cuticle. The cuticle is lamellar (“myelinic cuticle”) and formed by the superimposition of cellular unit membranes coalescing in 2 to 30 and more bilayers (Clément 1993). The basal part of the mastax ventrally contains the sclerotized jaws or trophi and their associated muscles, epithelial cells, salivary glands, sensory receptors, and a small nerve ganglion. The trophi comprise a set of hard, cuticularized extracellular elements formed by various epithelial cells

4.2 Morphology 

Fig. 4.31: Diagram of mastax. (A) Dorsal view, trophi forcipate, (B) lateral view, trophi forcipate, and (C) cross section, trophi malleate. Abbreviations: cu, cuticle; e, epithelium; f, fulcrum; m, manubrium; ml, mastax lumen; mo, mouth; mr, mastax sensory receptor; mu, muscles; r, ramus; s, stomach; sg, salivary gland; t, trophi; u, uncus. (Modified from de Beauchamp 1965 and Riemann & Ahlrichs 2008.)

(e.g., Koehler & Hayes 1969a, b, Clément & Wurdak 1991). Each element is build up of tubular sclerite bodies composed of an electron-dense core surrounded by electron-lucent material (e.g., Rieger & Tyler 1995). Some elements (ramus and manubrium) are hollow and contain the nucleus and perinuclear cytoplasm of an epithelial cell(s); others (uncus and fulcrum) are solid, lying next to their epithelial cell nucleus and cytoplasm. The main trophi elements (Fig. 4.31 C) consist of an unpaired median fulcrum, and 3 paired elements: ramus, uncus, and manubrium, connected by ligaments and moved by muscles connecting the different elements with each other and with the wall of the mastax. The development of the different elements and associated muscles is considerably modified in the different families or species, in relation to their mode of life and feeding habits (see below for a detailed description of the different trophi types). The highly complex musculature of the different trophi types is poorly known and relies on histological observations (e.g., Martini 1912, Seehaus 1930, Stossberg 1932, Remane 1933, de Beauchamp 1909, 1965), and TEM and/or CLSM of the monogononts Bryceella stylata, Dicranophorus forcipatus, Notholca acuminata, Pleurotrocha petromyzon, Proales tillyensis Trichocerca rattus, Asplanchna brightwellii, Notommata copeus, Bra-

 239

Fig. 4.32: Diagram of the musculature of the trophi apparatus of Dicranophorus forcipatus: (A–G) dorsal and (H–J) ventral. (A) Musculus fulcro-ramicus, (B) Musculus transversus manubrii, (c) musculus fulcro-manubricus, (d) musculus manubrico-uncus, (e) musculus caudo-ramicus, (f) musculus circumglandis, (g) musculus manubricus perioralis, (h) mastax receptor retractor, (i) musculus hypopharyngeus, and (j) musculus manubricohypopharyngeus. (Modified from Riemann & Ahlrichs 2008.)

chionus calyciflorus, and the bdelloid Philodina roseola (Clément 1987, Clément & Wurdak 1991, Sørensen et  al. 2003, Riemann & Ahlrichs 2008, Wilts et  al. 2010, Wulfken et al. 2010). There usually is a series of paired muscles (e.g., 13 in T. rattus, 6 in B. stylata) joining specifically paired trophi elements or trophi elements and mastax wall (Fig. 4.32). Paired muscles found in several mastax types studied to date are, e.g., abductor muscles interconnecting cauda of manubria with base of rami, muscles interconnecting the distal end of the fulcrum with the head of the manubria, malleus flexors interconnecting the cauda of the manubria with the unci, muscles connecting manubria to both the median and peripheral mastax floor, etc. The number of unpaired muscles is restricted: e.g., a strong hypopharynx muscle connects the enlarged dorsodistal end of the fulcrum with the opposite wall of the mastax lumen in species with virgate trophi; a weak retractor lies on the dorsal edge of the fulcrum and terminates in the mastax sensory receptor in species with forcipate and modified malleate

240 

 4 Rotifera

trophi; the rami adductor muscle connects both rami distally in, e.g., malleate and incudate trophi, and a transversal manubrium adductor connects the heads of both manubria in, e.g., malleate, modified malleate, forcipate, and virgate trophi. Comparison of the musculature in different species, and identification of homologous muscles is difficult. Possible homologous muscles are: the paired muscle connecting fulcrum and caudal part of rami, the paired muscle attached to the caudal end of the fulcrum and the dorsal edge of the head of the manubria, the paired muscle joining the distal end of the manubria and the unci, the paired muscle interconnecting the distal end of the manubria with the anterolateral part of the rami, and the unpaired hypopharynx muscle/ fulcrum sensory receptor retractor (e.g., Riemann & Ahlrichs 2008, Wilts et al. 2010, Wulfken et al. 2010). In bdelloids, mastax muscles are often laminar, and short with few cross-striations, unlike the longer and regularly striated muscles observed in monogononts (Clément & Wurdak 1991, Clément 1993). In T. rattus, all muscles proved unicellular, except one. The attachment of the muscles to the trophi elements and their adjacent epithelial cells is by desmosomes. The flexible ligaments connecting the trophi elements contain electron-dense cuticular material. The muscles are innervated by the mastax ganglion situated at the mastax floor. Several sensory receptors, such as ciliated pits and ciliated cells, occur between the trophi and the mastax floor, and on the roof of the mastax (Clément et al. 1983, 1991, Wurdak et al. 1983). A variable number of mononucleate or multinucleate salivary glands are generally present, closely associated with the mastax ventrolaterally (Fig. 4.33), or incorporated in the mastax wall (Fig. 4.31 C); apparently, they also may be lacking. Their anterior part is often transformed into a large reservoir (e.g., Asplanchna, Flosculariacea, median salivary gland of several bdelloids). The ducts open anterior to the trophi or in the buccal tube. The esophagus is short to long. de Beauchamp (1909), and later confirmed by Clément et al (1980b, 1991), distinguished 2 sections: the cuticular and the ciliary esophagus. The cuticular esophagus is the tube emanating from the mastax toward the stomach. It is an expansion of the dorsal pharyngeal wall of the mastax, composed of a thin epithelial layer surrounded by a thicker muscular layer and lined by a fine cuticle. The ciliated esophagus is the section between the cuticular esophagus and the stomach and made of ciliated cells, the cilia of which end in the stomach lumen. The cilia are long and form a vibratile flame, helping to move down the food and preventing regurgitation. The periesophageal musculature may consist of longitudinal and circular muscles or a reticular muscle layer and appears

Fig. 4.33: Diagram of the intestinal system in rotifers. Abbreviations: bt, buccal tube; bv, buccal velum; c, cloaca; cio. ciliary esophagus; cuo, cuticular esophagus; gg, gastric gland; i, intestine; m, mastax; s, stomach. (Modified from Clément & Wurdak 1991, with permission.)

correlated with different modes of feeding (Remane 1933). Several types of esophagus have been recognized, based on its total length, length and width of cuticular and ciliary sections, presence of a vibratile flame, etc. (Remane 1933). The stomach wall is syncytial in bdelloids and cellular with usually clearly defined cells in monogononts. In bdelloids (Philodina roseola, Habrotrocha rosa), the stomach lumen is lined by a thick, dense fibrillar terminal web (Fig. 4.34), lying directly under the plasma membrane (Mattern & Daniel 1966a, Schramm 1978a, Clément & Wurdak 1991). This web is perforated by large pores containing invaginations of the stomach cell membrane, pinching off digestive vesicles and vacuoles toward the cytoplasm. In monogononts, a thin fibrous terminal web, pierced by pores, may be present (Asplanchna) or lacking (Brachionus, Filinia, Notommata copeus, Rhinoglena, Trichocerca rattus). Digestive vacuoles are formed by endocytosis as well (Wurdak 1987, Clément & Wurdak 1991). The stomach is enveloped by a mesh of circular and longitudinal muscles or by syncytial muscle fibers. The anterior part of the stomach may be provided with an offset proventriculus (e.g., many Encentrum and Proales). Blind sacks of the stomach wall often occur in species showing intracellular digestion and/or symbiotic zoochlorellae (e.g., Ascomorpha, Birgea, Dicranophoroides caudatus, Gastropus, Itura). Usually, there is a pair of syncytial gastric glands opening at the junction of ciliated esophagus-stomach by a pore directly or by a more or less long duct. The shape of the glands is variable and may be ovate, rounded, lobate, kidney-shaped, tubular, etc. The glands are rich in rough

4.2 Morphology 

Fig. 4.34: Diagram of the apical part of the syncytial wall of the stomach in Bdelloidea. Abbreviations: cm, cell membrane; g, glycocalyx; ICL, intracytoplasmic lamina; p, pore; pd, part of cytoplasm with digestive vacuoles; sl, stomach lumen. (From Clément 1993, with permission.)

endoplasmic reticulum and Golgi complexes; secretory granules accumulate in the basal part of the glands next to the stomach (e.g., Clément & Wurdak 1991). The secretions of the gastric glands play a primary role in the extracellular digestion. Other gland-like structures of variable shape and number (Remane 1933), remembering gastric glands by structure and inclusions (de Beauchamp 1909), and mostly situated on the dorsal stomach wall occur in several species (e.g., Enteroplea lacustris, Epiphanes clavulata, Microcodon clavus, Tetrasiphon hydrocora). Gastric glands are rarely absent (e.g., Albertia typhlina), but mostly lacking in species with stomach blind sacks and/ or symbiotic zoochlorellae. The stomach either is distinctly constricted off from the intestine by a muscular sphincter, or merges gradually into it. The syncytial intestine is thin-walled and provided

 241

with motile cilia and microvilli; the latter may be replaced by atypical flattened cilia containing microtubules (Clément 1977, Clément & Wurdak 1991). In bdelloids, the microvilli show submembrane helicoidal filaments, which are lacking in monogononts. Externally, the intestine is provided with a continuation of the musculature of the stomach. The intestine empties in a short, contractile cloaca, opening dorsally at the base of the foot. In species with ventrally displaced foot, it opens terminally or even ventrally (Ploesoma); in Flosculariidae, it opens on a papilla. The lumen of the cloaca is lined by a thin epithelial syncytium surrounded by a reticular muscle layer. The cloacal wall (Asplanchna brightwellii) includes a multicellulary sensory receptor, with cilia and microvilli projecting toward the cloacal opening. The sensory cells are near the cloacal ganglion innervating the cloacal muscles. The syncytial wall of the cloaca merges imperceptibly into the syncytial integument, and is seldom ciliated (e.g., Clément et  al. 1983, Clément & Wurdak 1991, Ahlrichs 1993). Oviducts, bladder, or protonephridial ducts open in the cloaca as well. In some species, there is no cloaca or cloacal aperture, and fecal matter is ejected through the mouth (e.g., Asplanchna) or stored as defecation pellets in the stomach (e.g., Ascomorpha). There are many variations (Fig. 4.35) on the general scheme of the intestinal system as outlined above (see Remane 1933). In Collothecacea, the anterior end is transformed into a large buccal funnel. A semicircle of cilia in this funnel separates an anterior part or vestibulum, from a posterior chamber, the infundibulum. The mouth is at the base of the infundibulum, and leads to a narrow esophagus hanging freely in a very large cavity, the proventriculus. The small trophi are situated at the bottom of the proventriculus. The proventriculus is homologous to the mastax with its lumen enlarged into a food-storage organ.

Fig. 4.35: Variation of the intestinal system in Monogononta (A–E) and Bdelloidea (F, G). (A) Encentrum, (B) Synchaeta, (C) Asplanchna, (D) Collotheca, (E) Ascomorpha ecaudis, (F) Philodina, and (G) Habrotrocha. Abbreviations: b, bladder; cw, ciliary wreath; fp, fecal pellet; gg, gastric gland; i, intestine; if, infundibulum; mx, mastax; o, esophagus; p, proventriculus; pt, pharyngeal tube; s, stomach; v, vestibulum. (From Donner 1956, 1965.)

242 

 4 Rotifera

Digestion is generally extracellular, although intracellular digestion can take place (Asplanchna) after an initial extracellular digestion (Wurdak 1987). In some monogonont genera (e.g., Ascomorpha) and the bdelloid family Habrotrochidae, digestion is intracellular. In Habrotrochidae, the stomach is a syncytial protoplasmic mass without lumen. The food is formed into pellets or food vacuoles in a short chamber behind the mastax; the vacuoles become digested when circulating in the stomach protoplasm.

4.2.6.2 Trophi The trophi (Figs. 4.36–4.39) may be organized in a strictly bilaterally symmetrical (most species) to highly asymmetrical way (Aspelta, Trichocercidae). The fulcrum (absent in bdelloids) serves as an attachment for the 2 rami; together, these 3 elements are referred as incus. Adjoining them on each side are an uncus and manubrium, which hinging together form the malleus. The fulcrum is of variable length and can be plate- or rod-shaped and distally tapering, expanding, or forked, or provided with a basal plate. The rami are tall or flat in cross section, and their shape varies from roughly triangular to strongly elongate and almost parallel-sided. Their inner margin can be smooth or bears elongate sclerite

bodies, the rami scleropili, which may fuse into a ridge and/or a series of tooth-like projections. At the outer part of their base, rami may show lateral projections, the alulae. Rami and fulcrum lie in the same plane, or form a more or less great angle with each other. The rami operate like a forceps with the fulcrum as the base. The unci consist of a single tooth, or few to many teeth, often firmly fused into rigid plates. Each uncinal tooth consists of a head and shaft; the teeth are mostly unequal. A subuncus, composed of minute teeth, rods, or plate-shaped elements, is mostly present, and situated distally beneath the uncus. The manubria are more or less triangular, or crescent- or rod-shaped supports of the unci. They are composed of an expanded head or clava, connected to the uncus, and a more or less strongly elongate shaft terminating in a handle-like distal end, the cauda, which is present in Ploima but absent in Gnesiotrocha and Bdelloidea. The head or clava is composed of 3 chambers that may be strongly reduced or absent. The malleus lies in the same plane as the incus, or the unci and manubria stand in different planes to each other and/or to the incus. A variable number of diverse accessory elements, e.g., epipharynx, hypopharynx, suprarami, etc., is often present in monogononts. Studies by SEM of the embryonic development of the malleate trophi (Fig. 4.37) show that the first observable

Fig. 4.36: The main trophi types. (A) Basic plan, (B–D) malleate (Euchlanis: B, right; C, ventral view); Proales decipiens, (D) right; (E–G) virgate (Notommata copeus: E, ventral; D, right; G, Trichocerca rattus, ventral), (H–I) forcipate (H, Dicranophorus epicharis, ventral; I, Encentrum), (J) cardate (Lindia janickii, ventral), (K) incudate (Asplanchna), (L) uncinate (Collotheca), (M) malleoramate (Filinia, ventral), and (N) ramate (Macrotrachela). Abbreviations: ep, epipharynx; im, intramalleus; f, fulcrum; m, manubrium; r, ramus; u, uncus.

4.2 Morphology 

 243

Fig. 4.37: Development of malleate trophi. (A) Early embryo and (B) adult. Abbreviations: f, fulcrum; m, manubrium; r, ramus, ra, rami apophysis; rr, reinforced ramus ridges; u, uncus. Scale bar=10 µm.

and distinctly sclerotized structures are a double row of elongate sclerite bodies along the longitudinal axis, wherein the future unci, reinforced ramus ridges, rami apophyses, and fulcrum are recognizable (De Smet, personal observation). By addition of sclerite bodies and apposition of amorphous sclerite material, the trophi attain their definitive shape and size in the fully grown embryo. After hatching ramate, malleate, virgate, forcipate, malleoramate, and uncinate trophi apparently do not change in size or shape and remain constant during the life of the rotifer (Fontaneto et al. 2003, Fontaneto & Melone 2005, 2006), although some post-embryonic growth cannot be ruled out in some taxa with incudate trophi, viz. Asplanchna (Fontaneto & Melone 2005).

Nine main types of trophi are recognized (Fig. 4.36), based on the shape and size of the elements, the presence of any accessory parts, and the way they operate. Several transitional types are known, and some other types are that specialized and modified that they cannot be classified into any of the main types. Shape of trophi is an utmost important character in rotifer taxonomy (Wallace et  al. 2006), and even subtle differences in trophi shape may be useful in solving complexes of cryptic species; in monogononts, small details on the trophi shape can distinguish the species of the Epiphanes senta complex (Schröder & Walsh 2007) and between Brachionus manjavacas and B. plicatilis (Fontaneto et  al. 2007). In bdelloids, the newly described Abrochtha from the USA have statistical differences in the number of minor teeth (Birky et al. 2011).

Fig. 4.38: SEM pictures of the ramate trophi in Bdelloidea. (A, B) Dissotrocha aculeata, (C) Anomopus telphusae, (D) Rotaria tardigrada, (E) Otostephanos donneri, (F) Abrotrochtha carnivora. (A, C–F) Dorsal view and (B) ventral view. Scale bar = 5 µm.

244 

 4 Rotifera

Fig. 4.39: SEM pictures of trophi types in Monogononta. (A, B) Floscularia ringens, (C) Hexarthra sp., (D) Cupelopagis vorax, (E) Cyrtonia tuba, (F, G) Brachionus manjavacas, (H) Proales similis, (I) Cephalodella sp., (J) Pleurotrocha atlantica, (K, L) Notommata glyphura, (M) Notommata codonella, (N) Eothinia elongata, (O, P) Asplanchna priodonta, (Q) Asplanchnopus multiceps, (R) Encentrum algente, (S, T) Lindia deridderae, and (U, V) Dicranophorus forcipatus. Scale bar = 10 µm.

4.2 Morphology 

SEM pictures of the trophi can be found in the references mentioned below for the different types and in Figs. 4.38 and 4.39. Ramate. Rami sickle-shaped, flat. Fulcrum absent. Unci broad, usually with numerous teeth, occasionally broad semi-circular striated plates, usually with broad and narrow teeth. Manubria sickle-shaped, as lateral bands. Grinding. In subclass Bdelloidea only (e.g., Melone et al. 1998b, Melone & Fontaneto 2005). Malleoramate. Rami more or less triangular, flat. Fulcrum short. Unci broad, with numerous teeth, occasionally resembling striated plates, proximal teeth usually enlarged. Manubria crescent-shaped with 3 superimposed major chambers, without shaft. Grinding. In order Flosculariacea only (e.g., Nogrady & Segers 2002, De Smet 2005a). Uncinate. Similar to malleoramate type, but all trophi elements except unci strongly reduced. Unci teeth 2 or from 4 to 5, and only the first or first 2 of stout build, elongate and curved, forming supporting rods for mastax. Tearing. Only in Order Collothecacea: Atrochidae, Collothecidae. Malleate. All parts of stout build. Rami more or less triangular, flat, inner margin usually toothed. Fulcrum short. Unci with several (4–12) firmly connected teeth, often fused into plate. Manubria provided with a fairly short shaft. The malleate trophi are adapted for gripping, grinding, and pumping. In Order Ploima: e.g., Epiphanidae, Brachionidae, Euchlanidae, Mytilinidae, Trichotriidae (e.g., Segers et  al. 1994b, De Smet & Gibson 2008). The submalleate type of Lecanidae is characterized by manubria showing a relatively long and incurved shaft (e.g., Segers 1995). Virgate. All parts can be of slender build and/or thin. Rami broad, more or less triangular, recurved dorsally forming hemispherical dome. Fulcrum strongly elongated, distal end usually more or less expanded or strongly bent dorsally. Unci with few teeth, often only the first of stout build or teeth reduced. Manubria mostly with elongate shafts. Often strongly asymmetrical (e.g., Trichocerca). The virgate trophi type is the most variable of all. Used for swallowing food by pumping without crushing or piercing and sucking. The pumping action is produced by a powerful hypopharyngeal muscle. In Order Ploimida, e.g., Gastropodidae, Trichocercidae, Synchaetidae (e.g., Nogrady & Segers 2002). The malleovirgate type showing many uncinal teeth is a transition between the malleate and virgate type. In Proalidae (e.g., De Smet 1996). Incudate. Rami strongly elongate, curved, pincerlike. Fulcrum short. Unci and manubria strongly reduced.

 245

Specialized for seizing. Only in Asplanchnidae (e.g., Gilbert et al. 1979). Cardate. Rami lyrate. Fulcrum medium long. Unci a few distinct teeth, first largest, or a striated plate. Manubria with well developed head and shaft, head with characteristic crescent- or rod-shaped ventral apophysis. Species-specific accessory trophi elements present, often numerous. Adapted for pumping; the pumping action is produced without the hypopharyngeal muscle: the lumen is widened by a rolling motion of the trophi. Only in Lindiidae (e.g., Nogrady & Segers 2002, De Smet 2005b). Forcipate. Rami strongly elongate, straight, or curved, pincer-like, with toothed tips, inner margins often with few to numerous teeth. Fulcrum usually short to medium long. Unci strong, a single or few teeth only. Manubria rod-shaped, long, head reduced; often with intramalleus between uncus and manubrium. Seizing; rami and unci can be thrust out from mouth (Dicranophoridae). In Dicranophoridae and Ituridae (e.g., De Smet & Pourriot 1997). In the hemiforcipate type, rami lack apical and medial teeth. In Asciaporrectidae (De Smet 2006). Fulcrate. Fulcrum long, well developed. Rami short with strong, characteristic alulae. Unci small. Manubria absent. Pumping action performed by hypopharynx muscle attached to fulcrum. Only in Order Seisonacea (e.g., Segers & Melone 1998).

4.2.6.3 Intestinal system of the male The intestinal system of the male shows transitions from fully developed to completely absent. To date, Rhinoglena frontalis is the only species studied in which the males possess a fully developed and functional system similar to the female (Melone 2001). The male trophi are similar to the female but display fewer uncinal teeth and a reduction of the size to about 70%. In several species, e.g., Asplanchnopus multiceps, Eosphora najas, and Lacinularia flosculosa, the male intestinal system is well developed, but trophi are apparently lacking (Hamburger 1907, de Beauchamp 1965). In other males, the digestive tract consists of a strand of stomach cells with distinct lumen and indications of gastric glands (e.g., Mytilina, Synchaeta) or it is a strand without lumen (e.g., Asplanchna priodonta); the cellular strand supports the testis. The digestive tract may be further reduced to a globular mass of cells without lumen (e.g., Asplanchna brightwellii, Encentrum martes), supposed to form an energy source. The intestinal system is completely absent in males of, e.g., Conochilus, Filinia,

246 

 4 Rotifera

Hexathra, Keratella, Polyarthra, Pompholyx, and Trichocerca (e.g., Wesenberg-Lund 1923). The sensory receptors in the mouth region of the female, involved in feeding, are absent (Wurdak et al. 1983).

4.2.7 Body cavity The body cavity of the rotifers is considered a pseudocoel, as it is not lined by an epithelium, but by extracellular matrix. It is usually a spacious cavity, apparently lacking fibrils or microfilaments of collagen (Clément & Wurdak 1991, Clément 1993). The pseudocoel of several taxa, e.g., Asplanchna, Proales, Synchaeta, and Collothecacea, contains free amoeboid cells or amoebocytes (Nachtwey 1925, Remane 1933, Baumann et al. 2000); these cells could not be demonstrated in Brachionus, Notommata, and Trichocerca (Clément 1980). The amoebocytes form a highly dynamic, 3-dimensional polygonal network of filopodia. The cytoskeleton of the filopodia contains F-actin and microtubules that are often organized in bundles. Filopodial motion types include lateral junction displacement, formation and extension of free-ending filopodia, and fusion of filopodial strands, resulting in enlargements, diminutions, and extinctions of the filopodial polygons, and in the formation of new polygons (Baumann et al. 2000). An intense and fast particle transport takes place in the filopodial strands. The amoebocytes are often vacuolated, and contain inclusions, “bacteroids”, pigments, oil droplets, etc. In some species (e.g., Cupelopagis vorax, adult sessile Collotheca, Proales), specialized immobile amoeboid cells, the excretophores, apparently accumulate catabolites as excretion particles. The excretophores often pile up symmetrically near, e.g., the longitudinal and circular muscles, and the anterior part of the protonephridial tubule; their number and size increases with age (Remane 1933). Salt composition and volume of the pseudocoelomic fluid are regulated by the protonephridia (e.g., Braun et al. 1966, Pontin 1966). The pseudocoel functions as a hydrostatic skeleton, and supposedly as respiratory and circulatory system, its pseudocoelomic fluid being replenished by the oxygen carrying water taken in by the digestive tract, and subsequently eliminated through the protonephridia.

4.2.8 Excretory system 4.2.8.1 Excretory system of the female The excretory system of rotifers has been studied by LM (e.g., Remane 1933, Brakenhoff 1937, Pontin 1964) and

TEM (e.g., Braun et  al. 1966, Mattern & Daniel 1966b, Warner 1969, Schramm 1978b, Clément & Wurdak 1991, Ahlrichs 1993, Riemann & Ahlrichs 2010). It consists of 2 similar protonephridia lying ventrolaterally in the pseudocoel. The protonephridial apparatus is formed by 3–4 multinucleate cells and consists of few to several flame bulbs or terminal organs, attached to collecting tubules or capillary canals, which are connected to the main canals; the main canals discharge in an unpaired and contractile urinary bladder, or in a contractile cloaca (Fig. 4.40). The flame bulbs (Fig. 4.41) are conical and laterally flattened, appearing cylindrical or fan-shaped depending on their orientation (e.g., Asplanchna priodonta, A. brightwelllii, Dicranophorus forcipatus, Notommata copeus) or almost cylindrical and round in cross section (e.g., Encentrum mucronatum, Erignatha clastopis, Proales reinhardti). They are hollow, and their lumen drains into the capillary canals of the syncytium. The distal end of each flame bulb is closed by a protoplasmic cap. In monogononts, the lateral wall of the flame bulb usually consists of cytoplasmic columns and electron-dense microvilli (pillars), arranged in concentric rings around a central

Fig. 4.40: Protonephridium and contractile bladder of Asplanchna priodonta. Top 2 flame bulbs shown frontally; other 2 shown from the side. Abbreviations: cb, contractile bladder; ct, collecting tubule; ef, external filament; fb, flame bulb; mc, main canal; ps, protoplasmic strand; sc. syncytial cytoplasm; uv, uterovesicular duct; vf, vibratile flame. (Modifed from Pontin 1964.)

4.2 Morphology 

Fig. 4.41: Flame bulb. (A) Flame bulb partly opened to show internal organization, (B) cross section, and (C) detail of the filtering membrane in (1) bdelloids, (2) the monogonont Trichocerca rattus, (3) other monogononts (Asplanchna, Brachionus, Notommata copeus, Rhinoglena). Abbreviations: bl, basal lamina; cc, cytoplasmic columns; f, filtering membrane; m, microvilli; pc, protoplasmic cap; vb, vibratile flame. (A, B, From Braun et al. 1966; C, from Clément & Wurdak 1991, with permission.)

 247

lumen with vibratile flame. Columns, and occasionally microvilli, are connected by a filtering membrane. In bdelloids and some monogononts (Proales reinhardti), microvilli are absent or not distinct from columns, and the filtering membrane is sustained by both. The cilia of the vibratile flame are connected with each other and inserted on the protoplasmic cap containing their basal bodies. Ciliary rootlets extending into the filter region are usually absent in monogononts, but present in bdelloids (see, e.g., Riemann & Ahlrichs 2010). The basal membrane is often present at the outside. The distal cap of the flame bulbs is connected to the integument by protoplasmic strands, maintaining the position of the protonephridial apparatus. The number of flame bulbs on each side of the body varies from 2 to 100 according to the species and appears positively correlated with the surface area of the species (Pontin 1964); the number is fairly constant for any one species. The lumina of the flame bulbs are connected to the hollow cells of the protonephridial collecting tubule, and the latter joins with the intracytoplasmic lumen of the main canal syncytium. The intracytoplasmic lumen of the collecting tubules may be ciliated. The collecting tubules are mostly straight, whereas the main canals are straight, coiled, looped, or branched and often connected by a transversal canal, the Huxley anastomose, above the brain (e.g., Epiphanes, Lacinularia, Stephanoceros). Several protonephridium types (Fig. 4.42) have been described (for a review, see Remane 1933).

Fig. 4.42: Variation of the protonephridial system in Monogononta. Abbreviations: b, bladder; cc, capillary canal; fb, flame bulb; Ha, Huxley anastomose; mc, main canal. (From Remane 1933.)

248 

 4 Rotifera

A urinary bladder is only found in monogononts. It opens ventrally in the cloaca, or when occasionally absent, the protonephridia empty directly in the contractile cloaca (e.g., Conochilus, Lacinularia, Testudinella). The bladder syncytium is surrounded by a binuclear syncytial muscle, enabling the bladder to contract. In bdelloids, a separate urinary bladder is absent, and the protonephridia discharge in the terminal section of the intestine functioning as a bladder. The protonephridial system of the rotifers has both an excretory and osmoregulatory function (e.g., Braun et al. 1966, Pontin 1966). 4.2.8.2 Excretory system of the male Very little is known on the excretory system of the male (Wesenberg-Lund 1923), and results must be considered with caution (Remane 1933). Protonephridia have been found in most of the males described but appear absent in the strongly reduced ones. In several species flame bulbs, capillary ducts, and main canals are distinct (e.g., Asplanchna, Asplanchnopus, Epiphanes); the Huxley anastomose is present in male Epiphanes. The numbers of flame bulbs of the male and the female are identical (e.g., Epiphanes senta, Asplanchna priodonta, Mytilina) or apparently reduced in the male (Collothecacea). A contractile urinary bladder is usually absent but has been found in, e.g., Asplanchna, Asplanchnopus, Cyrtonia, Stephanoceros, and Cupelopagis. The protonephridia may discharge into the urinary bladder, into the vas deferens by a common duct, to the outside via separate pores near

the genital opening, or into a bladder-like structure lying dorsally from the testis.

4.2.9 Reproductive organs 4.2.9.1 Reproductive organs of the female The female reproductive system (Fig. 4.43) comprises the ovarium, the vitellarium, and a follicular layer surrounding both completely (bdelloids) or partly (monogononts) and continuing as an oviduct (e.g., Remane 1933, de Beauchamp 1965, Bentfeld 1971a, b, Amsellem & Ricci 1982, Clément & Wurdak 1991). In most species, the oviduct opens into the cloaca beyond the urinary bladder, or when the cloaca is reduced or absent, it directly leads to the outside. The female organs of monogononts consist of a single ovarium and vitellarium, whereas both organs are always paired in bdelloids, with the 2 oviducts joining into a single duct. They lie ventrally under the stomach and are mostly displaced to one side in monogononts. The syncytial ovarium is small; it lies close to the vitellarium and contains the oocytes. In full-grown females, the syncytial vitellarium is a very large, yolk-producing gland with large polyploid nuclei, the number of which is usually constant within a species. The most frequently observed number is 8, but 4, and a multiple of 4 up to 32, occurs. Individual variations in number of nuclei in function of diet and doses of vitamin E have been described (e.g., Birky & Field 1966, Amsellem & Ricci 1982). The

Fig. 4.43: Diagram of the female genital apparatus. (A) The bdelloid Philodina roseola and (B) the monogonont Asplanchna brightwellii. Abbreviations: cb, cytoplasmic bridge; de, developing embryo; ff, follicular folds; fl, follicular layer; i, integument; mo, maturing oocyte; n, nucleus with large nucleolus; ne, nucleolar extrusion; o, oocyte in ovarium; od, oviduct; ps, pseudocoelom; u, urogenital pore; v, vitellarium. (Modified from Clément & Wurdak 1991, with permission.)

4.2 Morphology 

shape of the vitellarium is rounded, lobed, band-shaped, horseshoe-shaped, cylindrical, etc., and often characteristic of the species.

4.2.9.2 Reproductive organs of the male The male reproductive system (Fig. 4.44) consists of the unpaired, large, globular to pyriform testis, one to several accessory or prostate glands, the short to long, and mostly ciliated vas deferens, the copulatory organ or penis, and the dorsally lying genital aperture that is usually surrounded by a ring of cilia (e.g., Wesenberg-Lund 1923, Remane 1933, de Beauchamp 1965). If a digestive tract or its rudiments are present, the testis lies ventral to it. The wall of the testis is syncytial. The tip of the testis bears a wreath of cilia and often tufts of sensory cilia and several openings (Brachionus, Asplanchna); its inside is lined by integument (Aloia & Moretti 1974, Clément et al. 1983, Clément & Wurdak 1991). The testis is filled up with spermatids, spermatozoa, and testicular rods. The penis is protrusible (e.g., Brachionus, Euchlanis, Gastropus) or projecting continually (e.g., Anuraeopsis, Keratella, Notholca). In some species, the penis is absent, and the evertable vas deferens acts as penis (e.g., Ascomorpha, Asplanchna, Epiphanes, Rhinoglena), or the tube-shaped posterior extremity of the male is specialized for copulation (e.g., Filinia, Hexarthra, Polyarthra). Two pairs of cutaneovisceral muscles may insert on the vas deferens and act as retractores penis.

Fig. 4.44: Male reproductive system. (A) Scheme of male genital organ in Monogononta, penis redrawn and (B) penis everted. Abbreviations: c, cloaca; cr, ciliary ring at genital opening; pg, prostate glands; s, spermatozoa; t, testis; tr, testicular rods; vd, vas deferens. (Modified after Remane 1933 and Koste 1978.)

 249

4.2.10 Gametes Oogenesis was studied by, e.g., Nachtwey (1925), Lehmensick (1926), Bentfeld (1971a, b), and Clément & Wurdak (1991) for monogononts and by Hsu (1956a, b), Clément & Wurdak (1991), and Pagani et al. (1993) for bdelloids. The number of oocytes is fixed at birth (e.g., Buchner et  al. 1965, Pagani et al. 1993). Maturation of the oocytes is by 2 subsequent equatorial divisions in bdelloids, with the extrusion of 2 polar bodies. Oogenesis of mononogonts is likewise by 2 equatioral divisions and extrusion of 1 polar body for the production of amictic eggs and 2 for the production of mictic ones (Gilbert 1993, Hsu 1956a, b). Important nucleolar extrusions are present in the cytoplasm of the immature oocytes of monogononts, but absent in bdelloids. During maturation, the volume of the oocyte increases considerably by the transfer of cytoplasm from the vitellarium through a cytoplasmic bridge; each oocyte has its own bridge (e.g., Bentfeld 1971b, Amsellem & Ricci 1982, Clément & Wurdak 1991). Subsequently, at the end of the growth period oocytes become detached from the ovary. After deposition of a shell in the oviduct, eggs are laid in oviparous species (most monogononts and bdelloids) or develop in the oviduct in ovoviviparous species. The term “viviparous” is often used for species who give live birth (e.g., the monogononts Albertia, Asplanchna, Lindia, Rhinoglena, Trochosphaera, and the bdelloids Rotaria), unlike the species who lay eggs enclosing a full-grown embryo that leaves the shell immediately after egg deposition. At hatching, juvenile females of free-swimming species usually have the adult shape. In sessile species, the females hatch as free-swimming larvae of typical rotifer appearance (Fig. 4.45), attaining the characteristic shape of the sessile stage after attachment and development into adults (e.g., Hochberg et  al. 2010, Fontaneto et al 2003). Males are sexually mature at hatching and do not grow. Three types of eggs are commonly produced in monogononts. The asexual or amictic egg (also called subitaneous egg) is a thin-shelled diploid egg produced by amictic females. The amictic egg develops into either amictic or mictic females. The sexual or mictic egg, also called male egg, is a small, thin-shelled haploid egg produced by mictic females; if unfertilized, it develop into males. Resting or dormant eggs are thick-shelled, fertilized mictic eggs, hatching into amictic females (Fig. 4.46). Resting eggs are actually resting embryos and not eggs (Boschetti et  al. 2011a; see Section 4.3.3, Development). A fourth type of egg is the pseudosexual egg (e.g., in Keratella hiemalis, Synchaeta), an unfertilized resting egg produced via parthenogenesis in the

250 

 4 Rotifera

absence of males (Ruttner-Kolisko 1946, 1974). Bdelloidea lack males, and their diploid eggs develop into females by parthenogenesis. Eggs are spherical, ellipsoid, ovate, kidney-shaped, etc. They may be free floating, glued by a short stalk to the substrate, carried attached to the body of the female by a gelatinous thread, etc.; in several species, the resting egg is retained within the lorica. Amictic and mictic eggs are usually smooth, unlike the resting eggs, which are mostly ornamented with spines, ridges, pits, gas-filled vacuoles, etc. The ornamentation is often characteristic of the genus and/or species (e.g., Pourriot et al. 1983). The resting egg is mostly provided with a groove at one pole, where the shell opens on hatching of the young female. The shell of the amictic egg in Trichocerca consists of an external shell

underlain by a thin internal envelope, joined to short microvilli at the periphery of the egg cytoplasm (Clément & Wurdak 1991). The shell of the resting egg of Asplanchna is composed of 3 coats: an external shell consisting of 2 coats separated by a space from an inner coat, which in its turn is separated by a space from the embryo (Wurdak et al. 1977). During spermatogenesis, the primordial germ cells produce the spermatozoa, and atypical germ cells, which produce the testicular rods, formerly described as rudimentary, non functional, or atypical spermatozoa (e.g., Whitney 1917, 1918, Tannreuther 1919, Koehler 1965, Koehler & Birky 1966, Aloia & Moretti 1974). The spermatozoa of a few monogononts, belonging to the genera Asplanchna, Brachionus, Cephalodella,

Fig. 4.45: Free-swimming larvae and sessile adults. (A) Sinantherina socialis, (B) Stephanoceros fimbriatus, and (C) Cupelopagis vorax. Abbreviations: a, adult female; l, larva. (From Remane 1933, Koste 1978, and Kutikova 1970.)

Fig. 4.46: SEM pictures of resting eggs of Monogononta: (A) Asplanchna priodonta, (B) Brachionus plicatilis, (C) B. calyciflorus, (D) Epiphanes brachionus, (E) Rhinoglena frontalis, (F) Notommata codonella, (G) Conochilus natans, (H) Filinia longiseta, and (I) Hexarthra mira. Scale bar = 20 µm, except for (G) 10 µm. (A–F, Photo courtesy of Giulio Melone; G–I, Photo courtesy of Hendrik Segers.)



Epiphanes, and Lacinularia, have been described with LM (Hamburger 1907, Whitney 1917, 1918, Tannreuther 1919, Tauson 1927). Ultrastructural studies are limited to Asplanchna brightwellii (Koehler 1965, Koehler & Birky 1966, Aloia & Moretti 1974), A. sieboldii (Koehler 1965), Brachionus plicatilis (Melone & Ferraguti 1994), B. sericus (Clément 1977, Clément & Wurdak 1991), and Epiphanes senta (Melone & Ferraguti 1999). The general organization of the spermatozoon is similar for the species studied, except for some minor differences concerning shape and size (Fig. 4.47). The spermatozoon is an elongated cell with a posterior cell body containing the nucleus and an anterior complex composed of a flagellum and undulating membrane; an acrosome is absent. The single axoneme is intracellular and shows the usual 9 × 2 + 2 tubules, in which the inner dynein arms only are present (Melone & Ferraguti 1994, 1999). The elongate nucleus is lobate and located anteriorly behind the flagellar region (Koehler 1965, Melone & Ferraguti 1994, 1999). The cytoplasm contains randomly distributed elongate mitochondria, different types of vesicles, free ribosomes, dense granules, etc. The shape and length of the spermatozoa studied by EM is as follows: Asplanchna: gourd-shaped, 15–20 µm; Brachionus plicatilis: filliform, 70–80 µm (flagellum 40–45 µm, cell body 30–35 µm); Epihanes senta: sausageshaped, bent, and weakly twisted, 12–15 µm (cell body 8–10 µm).

4.3 Reproduction and development 

 251

The testicular rods are rod-shaped structures developing late in embryonic life, i.e., in essentially mature males, from atypical germ cells showing intense secretory activity (Koehler 1965, Koehler & Birky 1966, Clément 1977, Melone & Ferraguti 1994). The rods contain dense material originating from Golgi vesicles and are organized as densely packed microtubules; a nucleus and flagellum are lacking. They are extruded from the cell within 2 membranes, one derived from the Golgi vesicle and one from the cell membrane. Testicular rods occur in almost all monogonont families and are supposed to assist, mechanically or enzymatically, in the penetration through the female integument during copulation (Remane 1933, Koehler 1965). They are always much smaller than the spermatozoa, e.g., 15 × 2 µm in Asplanchna, 16 × 1.5 µm in Brachionus plicatilis, and 14 × 1 µm in Epiphanes senta (Melone & Ferraguti 1999).

4.3 Reproduction and development 4.3.1 Reproductive biology Rotifers are a textbook example of the different types of reproduction in animals. Seisonacea are characterized by sexual reproduction, which is in common with most animals (see chapter on Seisonacea). Monogononta can

Fig. 4.47: Spermatozoon. (A) Diagram of spermatozoon of Monogononta, (B) spermatozoon of Brachionus plicatilis with representative cross sections, and (C) Spermatozoon of Epiphanes senta. Abbreviations: c, centriole; m, mitochondrion; n, nucleus; u, undulating membrane; v, vesicles containing tubules. (A, Modified from Melone & Ferraguti 1999; B, C, modified from Melone & Ferraguti 1994.)

252 

 4 Rotifera

alternate parthenogenesis, both through the development of diploid eggs producing females and through the development of haploid eggs producing males, to sexual reproduction (Serra & Snell 2009). Bdelloidea are the most famous case of ancient asexuals and the whole group diversified and persisted in the absence of any known form of sexual reproduction (Mark Welch et al. 2009). Clear absence of sex in Bdelloidea cannot be proven unambiguously, as hidden sex may always occur. Yet, a large amount of observation on Bdelloidea never produced any evidence of meiosis, males, hermaphrodites, or vestigial genital structures (Birky 2010). Moreover, all the available indirect cytological and genetic evidence concurs in supporting their actual asexuality (Normark et al. 2003, Mark Welch et al. 2009): high sequence divergence between gene copies, functional divergence of former alleles, and multiple functionally divergent and conserved copies have been found (Mark Welch & Meselson 2000, Pouchkina-Stantcheva et  al. 2007, Eyres et  al. 2012), there is lack of high-copy-number retrotransposons (Arkhipova & Meselson 2000, 2005), an accumulation of deleterious mutations has been found (Barraclough et  al. 2007, Swanstrom et  al. 2011), and the diversification rates in Bdelloidea are clearly different from those in Monogononta (Fontaneto et  al. 2012b). According to our current knowledge, Bdelloidea produce only diploid eggs that hatch into females that can lay eggs that hatch into females, without any form of sexual recombination. Thus, the whole genome of the mother is inherited by all the daughters, producing clonal lines. Because of the evolutionary success of Bdelloidea in the absence of sexual reproduction, Maynard Smith (1986) dubbed them as an “evolutionary scandal”. This makes bdelloids an excellent study system for the origin of sex in animals (Bell 1982) by understanding how these peculiar organisms thrive in the absence of sex. One caveat is nevertheless present due to the possibility of horizontally transferred genes, which are abundant and functional in Bdelloidea (Boschetti et al. 2012) and may represent an alternative mechanism to sexual recombination to maintain genetic variability (Gladyshev et al. 2008). A rather peculiar reproductive behavior is present also in the other group of Rotifera, the Monogononta, characterized by cyclical parthenogenesis. Cyclical parthenogenesis is a combination of asexual, parthenogenetic, and sexual reproduction (Serra & Snell 2009). The conditions thought to favor cyclical parthenogenesis are seasonal and/or unpredictable temporal heterogeneity (Carmona et  al. 1995, Serra et  al. 2003, Ricci 2001), typical of the ephemeral habitats where most Monogo-

nonta thrive. In Monogononta, asexual females, called amictic females, produce asexual daughters by ameiotic (apomictic or amictic) parthenogenesis. In response to certain environmental cues such as population density, photoperiod, or chemical triggers (Pourriot & Clément 1981, Stelzer & Snell 2003, Snell 2011), asexual females produce sexual daughters, called mictic females (Gilbert 1963b). These sexual females produce meiotic eggs, which, if not fertilized, develop into haploid males. If fertilized by a male, eggs develop into cysts, called resting eggs, which undergo diapause. When resting eggs hatch, the cycle starts again and asexual females reproduce by parthenogenesis until the next round of sexual reproduction. Variations to this life cycle have been described, including amphoteric females producing both ameiotic and meiotic eggs (Gilbert 1974, King & Snell 1977), parthenogenetic resting eggs (Gilbert 1995, Gilbert & Schreiber 1998), and sexual females hatched from resting eggs (Schröder et  al. 2007). The mechanisms involved in mating behavior in monogononts has been studied mostly for those species used in aquaculture (Hagiwara et al. 1995b, Rico-Martínez & Snell 1996, 1997), and a set of molecules potentially involved in mate recognition have already been tested and described (Snell 2011) (see Section 4.4, Physiology).

4.3.2 Cleavage Most Rotifera are oviparous, but several ovoviviparous species retain the developing embryos inside the body (Gilbert 1989). Oviparous species may carry the eggs, attach them to the substrate, or simply release them in the water column (Wallace et al. 2006). Rotifera possess a deterministic cleavage that follows a modified spiral pattern (Gilbert 1989). The fate of the cells is established very early in the development (Pray 1965). Moreover, Rotifera are eutelic (Clément & Wurdak 1991); cell divisions occur only during embryogenesis, whereas the newborn already possesses the same number of cells of the adult. Embryo development follows the 3 typical steps of cleavage, gastrulation, and organogenesis (Boschetti et al. 2005). Eggs are poor in yolk; they are laid unsegmented and extrude the polar body before undergoing cleavage (Hsu 1965a, b, Gilbert 1989). During cleavage, subsequent divisions produce smaller and smaller cells of unequal size and amount of cytoplasm, called blastomeres. Cleavage is holoblastic and cell divisions are unequal. For a review of the current knowledge on cleavage, see Boschetti et al. (2005). A typical 16-blastopore stage is present with 4

4.4 Physiology 

rows of 4 cells each, and at this stage, gastrulation starts (Gilbert 1989, Boschetti et al. 2005).

4.3.3 Development Gastrulation in Rotifera occurs by epibolic movements and consequent involution; the resultant gastrula is a stereogastrula without a recognizable internal cavity. Lechner (1966) described 2 stages in the gastrulation process, a first epibolic growth of the blastoderm cell and a second stage with a further epibolic growth coupled with the involution of the blastoderm to form the blastopore. Not all authors agree on this 2-stage process, on the position of the blastopore and the origin of the mouth, and on the origin on the digestive system (Boschetti et al. 2005). Modern analyses should be performed to obtain a clear description of the embryo development in Rotifera. Cell lineage mapping has been performed only on few species (Zelinka 1892b, Pray 1965, Lechner 1966), and organogenesis is not well known, with discrepancies in the scattered available information. The main differences concern the origin of the digestive and the reproductive systems (Boschetti et al. 2005). According to Gilbert (1989), the stomodaeum, the pharynx, the nervous system, the excretory system, and the muscles originate from the ectoderm; the reproductive system, the germarium, and the vitellarium from the mesoderm; the digestive system from the endoderm. It has to be reminded that Rotifera are eutelic (Clément & Wurdak 1991) and possess a constant number of cells (or better, of nuclei) throughout all life because several tissues are syncytial. Cell divisions occur during embryogenesis only, and the fixed number of cells is supposed to be species specific (Boschetti et al. 2005). The newborn possesses the same number of cells of the adult, and the cells will only increase in size during life. In sessile Rotifera, the juveniles that hatch from the eggs are very different from the adults, and undergo radical developmental changes (Fontaneto et  al. 2003, Wallace et  al. 2006). Such changes include formation of the adult corona and elongation of the foot (see Section 4.2.1.2, Coloniality). Development of resting eggs is also poorly understood (Wurdak et al. 1978, Hagiwara et al. 1995a, Boschetti et al. 2011a). The so-called resting eggs are actually diapausing embryos that will resume their development after being activated by external and/or internal stimuli (Gilbert 1974, Pourriot & Snell 1983, García-Roger et al. 2006).

 253

4.4 Physiology Very few studies deal with the general physiology of Rotifera. For example, almost nothing is known on the digestive system; we only know some of the enzymes involved and the absorptive process (Lindemann et al. 2001). More efforts have been implemented to analyze the triggers of the switch between sexual and asexual reproduction in Monogononta (Snell 2011), the physiological aspects of dormancy in Bdelloidea (Tunnacliffe & Wise 2007), hormone-induced phenotypic plasticity (Gilbert 1999), and the ecotoxicological and ecophysiological mechanisms (Dahms et al. 2011). No rotifer hormones have been yet described (Snell 2011), but the rotifers that have been tested respond to a variety of hormones and neurotransmitters from vertebrates and insects (Gallardo et al. 1997, 1999, 2000, Snell & DesRosiers 2008), including growth hormone, human chorionic gonadotropin, triiodothyronine, juvenile hormone, serotonin, GABA, and progesterone. Several putative endocrine disruptors, sufficiently similar to hormones that they interfere with normal endocrine signaling in aquatic animals have effects on rotifers (Snell & Joaquim-Justo 2007). Moreover, rotifers indeed produce growth-promoting substances (Ohmori et al. 2011). Evidence that rotifers use chemical signals as pheromones is available in rotifers, especially to regulate their reproduction (Snell 2011). These include compounds excreted in the medium that trigger the switch from sexual to asexual reproduction (Gilbert 1963a, Stelzer & Snell 2006), and mate recognition pheromones (Snell et al. 1995, Snell & Stelzer 2005, Snell et al. 2009). Also, environmental triggers have been described, such as dietary α-tocopherol, photoperiod, and crowding (Gilbert 1981, Pourriot & Clément 1981, Stelzer & Snell 2003). The mechanism that switches asexual to sexual reproduction in Monogononta is controlled by the accumulation of a signaling molecule produced by the rotifers themselves (Gilbert 2004). Such population-level metabolic synchrony, regulating gene expression in response to fluctuations in population density, is similar to what known as quorum sensing in bacteria (Miller & Bassler 2001). Rotifera, together with other microscopic aquatic animals such as tardigrades and nematodes, are able to withstand lack of water, in a desiccated and/or frozen state, entering dormancy. Bdelloidea may enter dormancy in any stage of their life cycle (Ricci & Fontaneto 2009), whereas Monogononta produce resting eggs that can remain dry and/or frozen for extended periods before hatching (Gilbert 1974, Schröder 2005). In the dry state,

254 

 4 Rotifera

there is little or no evidence of metabolic activity (Clegg 1986, 2001). The desiccating process in Bdelloidea is influenced by the nature of the substrate and the humidity; the process involves morphological changes such as the contraction of the body into a tun shape and the packing of internal structures (Ricci et al. 2003, 2008). The biochemical mechanisms used to survive by microscopic animals during dormancy are not fully understood (Clegg 2001, Oliver et al. 2005, Tunnacliffe & Wise 2007, Lubzens et al. 2010), but the main actors are thought to be specific molecules that protect the intracellular and extracellular environment against the various damages induced by the lack of water (Crowe et al. 1998, Rebecchi et  al. 2007). Sugars such as trehalose and sucrose were, for a time, considered the main molecules involved in the process in plants and animals (Alpert 2006). Yet, trehalose, present in nematodes and tardigrades, is absent in Bdelloidea (Lapinski & Tunnacliffe 2003) and found only in traces in Monogononta (Caprioli et al. 2004). It has been recognized that other molecules are important as well in the physiology of dormancy (Tunnacliffe & Lapinski 2009): these include heat shock proteins, late embryogenesis abundant proteins, chaperones, antioxidants, and others (Goyal et  al. 2005, Denekamp et  al. 2009, Clark et  al. 2012). Intriguingly, even foreign genes, acquired through horizontal gene transfer, seem to be involved in the desiccation response of Bdelloidea (Boschetti et al. 2011b). Whereas Bdelloidea recover from their dormant stage as soon as water becomes available (Ricci 1998a), hatching of resting eggs of Monogononta is triggered by a more complex mechanism, with temperature and/or light conditions, interacting with the length of desiccation (Minkoff et al. 1983, Schröder 2005). Several planktonic rotifers exhibit considerable environmentally controlled variation in spine development and body size. The most striking case is the elongation of spines in loricated rotifers in response to the presence of specific predators (Gilbert 1999, 2011). Such elongation of spines occurs due to chemical compounds, called kairomones, identified in the environment by rotifer mothers, which produce daughters with longer spines (Gilbert 1967). The physiological response to chemical changes in the environment is studied in detail in Rotifera, and ecotoxicological analyses have been performed both in the laboratory and in the field (Snell & Janssen 1995). Rotifers are used in the risk assessment of pharmaceuticals, endocrine disruptors, and heavy metal pollution, through whole-animal bioassays and gene expression

studies (Dahms et al. 2011). Responses regarding growth condition, reproduction, population dynamics, and toxicity are well known, especially for the laboratory model of the genus Brachionus (Yúfera 2001, Sarma et al. 2001, 2005, 2009).

4.5 Phylogeny The close relationship between the 3 traditional groups of Rotifera (Bdelloidea, Monogononta, and Seisonacea) and Acanthocephala is well accepted in traditional morphological studies; Rotifera and Acanthocephala share a peculiar syncytial epidermis with an ICL (Clément 1993, Wallace et al. 1996, Garey et al. 1998) and a similar morphology of the sperm cell (Melone & Ferraguti 1994, Ferraguti & Melone 1999). In addition, phylogenetic analyses using several molecular markers support their close relationship (Garcia-Varela & Nadler 2006), so that they have been grouped in the taxon Syndermata (Ahlrichs 1997). The monophyly of the 3 major groups of Rotifera is not debated (Herlyn et al. 2003, Sørensen & Giribet 2006). Nevertheless, their evolutionary relationships are still unclear, and different analyses usually provide discordant results regarding their position and the position of Acanthocephala (Lasek-Nesselquist 2012). This latter group can be supported either as a sister group of monophyletic Rotifera, or at different positions within Rotifera (Herlyn et al. 2003, Sørensen & Giribet 2006, Fontaneto & Jondelius 2011). The 6 hypotheses regarding the possible phylogenetic relationships within the clade Rotifera+Acanthocephala (Syndermata) are, in historical order, (a)  Bdelloidea, Monogononta, and Seisonacea (traditionally named Rotifera) as a monophyletic clade, sister to Acanthocephala (Fig. 4.48 A): This hypothesis is based on general morphological similarities and supported by morphological cladistic analyses (Melone et  al. 1998b, Sørensen & Giribet 2006), but never supported by molecular phylogenies. (b)  Lemniscea (Fig. 4.48 B): The hypothesis of Bdelloidea and Acanthocephala as sister groups is based on the supposed presence of a proboscis and lemnisci in Bdelloidea (Lorenzen 1985). Ricci (1998b) argued that the morphological basis of the hypothesis was false, due to a misinterpretation of the rostrum and the hypodermic cushions of Bdelloidea as homologous to the proboscis and lemnisci of Acanthocephala. Notwithstanding the unreliability of the morphological basis, this first analysis brought attention to the Rotifera+Acanthocephala relationship. Moreover, the first phylogenetic studies on

4.6 Systematics 

 255

Fig. 4.48: Alternative phylogenetic hypotheses for the clade Rotifera+Acanthocephala: (A) classic theory of a monophyletic origin of the Rotifera, (B) Lemniscea hypothesis, (C) Pararotatoria hypothesis, (D) Acanthocephala as Eurotatoria sister group, (E) Hemirotifera hypothesis, and (F) Seisonacea as Lemniscea sister group. Abbreviations: e, Eurotatoria; h, Hemirotifera; le, Lemniscea; p, Pararotatoria; r, Rotifera. (Modified from Fontaneto & Jondelius 2011.)

16S and 18S rDNA supported such relationship (Garey et al. 1996). The position of Seisonacea and Acanthocephala was not included in the discussion on Lemniscea. (c)  Pararotatoria (Fig. 4.48 C): The hypothesis of Eurotatoria (Bdelloidea+Monogononta) as a sister group of Pararotatoria (Acanthocephala+Seisonacea) is based on morphological characters such as sperm morphology and epidermal ultrastructure (Ahlrichs 1997). Moreover, this relationship was supported by the inclusion of Seisonacea in the 18S molecular data set (Herlyn et al. 2003). Acanthocephala as a sister group to Eurotatoria (d)   (Bdelloidea+Monogononta) (Fig. 4.48 D): This hypothesis is supported by a phylogenetic analysis of the nuclear gene hsp82 (Mark Welch 2000) and from a combined analysis of hsp82+18S (Mark Welch 2005). Seisonacea were not included, as no hsp82 was available for them. (e)  Hemirotifera (Fig. 4.48 E): In this hypothesis, Monogononta are the sister group of Hemirotifera (Acanthoc ephala+Bdelloidea+Seisonacea), based on combined molecular and total evidence analyses of 74 morphological characters and DNA sequence data from 18S, 28S, histone H3, and COI (Sørensen & Giribet 2006). (f)  Monogononta as a sister group of the other 3 groups, and Seisonacea as a sister group of Lemniscea (Fig. 4.48 F), inferred from a phylogenetic analysis of concatenated 18S+28S+COI (Garcia-Varela & Nadler 2006) and supported by EST phylogenomics (Witek et al. 2008). Almost any plausible relationship among Acanthocephala, Bdelloidea, Monogononta, and Seisonacea has been suggested, with no agreement between different analyses (Fontaneto & Jondelius 2011, Lasek-Nesselquist 2012). Two

main patterns emerge from all the analyses performed until now: (i) the traditional hypothesis of Acanthocephala as a sister clade of monophyletic Rotifera has never been supported by molecular phylogenies and (ii) the most recent analyses even split the clade Eurotatoria (Min & Park 2009, Witek et  al. 2009). Nevertheless, Seisonacea, Bdelloidea, and Monogononta share many morphological and ecological features, and there is no morphological support for the splitting of Eurotatoria (Melone et al. 1998b). The only unambiguous pattern is that each of the 3 groups of Rotifera (Bdelloidea, Monogononta, and Seisonacea) is a monophyletic clade. But the relationships between them are still not clear.

4.6 Systematics Classically, 3 groups are recognized within Phylum Rotifera: Seisonacea, Bdelloidea, and Monogononta. The classification we report here is that of Segers (2002), based on morphology alone, given that the phylogenetic relationships have not been solved yet for rotifers using molecular approaches (see Section 4.5, Phylogeny).

4.6.1 Classification Class Pararotatoria Sudzuki, 1964 Order Seisonacea Wesenberg-Lund, 1899 Class Eurotatoria De Ridder, 1957 Subclass Bdelloidea Hudson, 1884 Order Adinetida Melone & Ricci 1995 Order Philodinavida Melone & Ricci 1995 Order Philodinida Melone & Ricci 1995

256 

 4 Rotifera

Subclass Monogononta Plate, 1889 Superorder Pseudotrocha Kutikova, 1970 Order Ploima Hudson & Gosse, 1886 Superorder Gnesiotrocha Kutikova, 1970 Order Flosculariacea Harring, 1913 Order Collothecacea Harring, 1913 Subclass Bdelloidea (Fig. 4.49) is a group of completely asexual animals, reproducing by apomictic thelytoky. Ovaries and vitellaria paired. Body composed of telescopically retractable pseudosegments. Usually illoricate. Corona characteristic, composed of 2 trochal discs, may be reduced to buccal field. Head bears dorsal antenna and ciliated rostrum. Lateral antennae lacking. Number of toes 2, 3, or 4 or adhesive disc. Penultimate pseudosegment with spurs. Stomach syncytial. Trophi ramate. The bdelloids comprise 4 families in 3 orders (Melone et al. 1998b), to accommodate 19 genera and about 460 species (Segers 2007): order Adinetida with family Adinetidae (2 genera), order Philodinida with families Habrotrochidae (3 genera) and Philodinidae (11 genera), and order Philodinavida with family Philodinavidae (3 genera). Species count for bdelloids is done considering all subspecies as valid species; given the problems in

defining species in asexual taxa, we here follow the rule that every morphological variability described at the subspecies level has the validity of species. Order Adinetida is a group of bdelloids with fusiform to vermiform body, mostly compressed dorsoventrally, characterized by a corona of the Adineta type, without trochus and with a ventral ciliated field with rake of teeth at its base or at its lateral sides. Only 1 family, Adinetidae, with 20 species. Order Philodinavida is a group of bdelloid rotifers characterized by trophi close to the mouth opening and extruded when animal is feeding. The corona is poorly developed, reduced to small ciliated field. Only 1 family, Philodinavidae, with 9 species. Order Philodinida is the richest and most diverse group of bdelloids, characterized by a Philodina-type corona, deep and not protrusible trophi. Two families with about 435 species. Subclass Monogononta shows cyclic parthenogenesis, interrupted by sexual reproduction. Males usually strongly reduced. Fertilization internal, resulting in dormant (resting) eggs. Single ovary, vitellarium, and testis. Body pseudosegmented, shape very various, usually with distinct head, trunk, and foot. Loricate or illoricate. Foot and

Fig. 4.49: SEM pictures of Bdelloidea. (A) Adineta tuberculosa, (B) Rotaria macrura, and (C) Dissotrocha aculeata. Ventral view. Scale bar = 20 µm. (Photo courtesy of Giulio Melone.)

4.6 Systematics 

toes very various, usually 2 toes, or adhesive disc; toes may be absent. Lateral antennae always present. Corona and trophi of various types. Dorsal antenna present, ciliated rostrum absent. Stomach cellular. About 1,600 species are recognized in 3 orders: Ploima, Flosculariacea, and Collothecacea, comprising 23 families and 113 genera. Order Ploima (Fig. 4.50) is extremely varied in body plan, corona, toes, and trophi. Never sessile permanently. Trochus never surrounding apical field; corona often with pseudotrochus differentiated by the buccal field. Usually

 257

2 toes, rarely single toe, toes often absent in planktonic species. Trophi forcipate, incudate, malleate, or virgate. About 1,410 species. Order Flosculariacea (Fig. 4.51) is a group of monogonont rotifers, characterized by malleoramate trophi. Body plan variable; free-swimming juveniles differ in shape from adult females. Foot, if present, without toes; in juveniles and free-swimming species terminating in ring of cilia or ciliated cup. Loricate or illoricate. Corona of Hexarthra or Conochilus type. About 140 species.

Fig. 4.50: SEM pictures of Ploima. (A–C), Brachionus manjavacas, (D) Proales theodora; E–G, Dicranophorus forcipatus, (H) Notommata glyphura, (I) Mytilina ventralis, and (J) Trichotria tetractis. Scale bar = 25 µm. (A–I, photo courtesy of Giulio Melone; J, photo courtesy of Eike F. Wiltz.)

258 

 4 Rotifera

Fig. 4.51: SEM pictures of Flosculariacea, (A) Conochilus hippocrepis, (B) Filinia longiseta, (C) Flosculariacea ringens, and (D, E) Hexarthra mira. Scale bar = 50 µm. (A, C–E, photo courtesy of Giulio Melone; B, photo courtesy of Eike F. Wiltz.)

Fig. 4.52: SEM pictures of Collothecacea. (A–C) Cupelopagis vorax and (D) Stephanoceros fimbriatus. Scale bar = 50 µm. (Photo courtesy of Giulio Melone.)

Order Collothecacea (Fig. 4.52) is a group of rotifers characterized by uncinate trophi. Body elongate conical or saccate; foot always without toes, long with small adhesive disc, or modified into ventral adhesive disc, or reduced;

foot of free-swimming juveniles with ciliary ring. Illoricate. Corona of Collotheca type, a modified funnel with or without long setae, and often lacking cilia. Most species are sessile, but several live in the plankton. About 51 species.

4.6 Systematics 

4.6.2 Keys

01b – Esophagus long, trophi deeper, never extruded. Proximal and distal minor teeth of uncus plate always present. Corona otherwise, with trochi absent or present

4.6.2.1 Key to higher taxa 01a – Trophi fulcrate. Corona rudimentary. Body with small oval head, long slender neck, fusiform trunk, and stalk-like foot terminating in adhesive disk. Epizoic on marine crustaceans of the genus Nebalia. Males and females in equal numbers. Paired ovaries

… Class Pararotatoria, Order Seisonacea

01b – Trophi not fulcrate. Corona mostly well developed. Body not as above. Not epizoic on Nebalia. Mostly or only females. Ovaries paired or ……………………………… 02 single ovary 02a – Paired ovaries. Trophi ramate 02b – Single ovary. Trophi not ramate (Class Eurotatoria, Subclass Monogononta) 03a – Trophi malleoramate. Corona elliptical to round, heartshaped, horseshoe-shaped, or 4-lobed, without long setae. Free-swimming or sessile 03b – Trophi uncinate. Corona funnel-like with tentacles, lobes, or knobs bearing long setae; there may be cilia between the tentacles, lobes, etc. Usually sessile, a few planktonic species 03c – Trophi otherwise: cardate, forcipate, incudate, malleate, or virgate

…………………………… 02

…………… Adinetidae

…… Class Eurotatoria, Subclass Bdelloidea

02b – Corona frontal, with trochi, mostly elevated on pedicels. Foot with 2 spurs of variable length. Toes present (2, 3, or 4) or absent. Vortex feeding

…………………………… 03

03a – Stomach without recognizable lumen, with round pellets in its wall

…… Habrotrochidae

03b – Stomach with thick wall and visible lumen, not filled with round pellets

………… Philodinidae

……………………… Order Flosculariacea

…………………………… 03

4.6.2.3 Key to families of Ploima

……………………… Order Collothecacea

…………… Order Ploima

4.6.2.2 Key to families of Bdelloidea 01a – Esophagus short; trophi usually close to the mouth opening, generally extruded when animal is feeding. Proximal minor teeth of uncus plate usually reduced or absent. Corona reduced to small ciliated field, trochi absent, or small

02a – Corona modified to a ventral ciliated field, no trochi. Foot long and extensible, with 3 toes and 2 spurs, or short and plump, with disc-shaped distal pseudosegment, and spurs absent. Common behavior is scraping and browsing the substratum

 259

…… Philodinavidae

01a – Anterior half of body composed of 16 pseudosegments with finely denticulate distal margin. Endoparasitic in earthworm Pheretima

……………… Clariaidae

01b – Anterior half of body not composed of 16 pseudosegments. Not parasitic in Pheretima

………………………… 02

02a – Vitellarium strongly elongate, cylindrical, wound around intestine, with 20–30 nuclei. Connection between stomach and intestine narrow, surrounded by wreath of ca. 6 spherical glands

…… Tetrasiphonidae

260 

 4 Rotifera

02b – Vitellarium spherical, ovate, U-shaped, lobed, etc., not cylindrical and wound around intestine, usually with 8 nuclei. No wreath of glands between stomach and intestine 03a – Mouth opening ventral, surrounded by hollow cupshaped structure. Head covered by non-retractile shield. Trophi specialized virgate with 2 stilettoshaped elements 03b – Not as above 04a – Trophi highly modified, manubria heart-shaped, 2 large hook-shaped accessory elements. Stomach with blind sacks, stomach wall with green zoochlorellae. Gastric glands absent 04b – Not as above 05a – Trophi hemiforcipate, manubria with long rod-shape shaft and small axe-shaped head. Free-living or symbiotic in shells of testate amoebae

………… ……………… 03

…… Cotylegaleatidae …………………………… 04

……………… Birgeidae …………………………… 05

…… Asciaporrectidae

05b – Trophi not hemiforcipate with long manubria, if hemiforcipate with medium long manubria with broad head, and 2 trapezoid epipharyngeal plates. Free-living or symbiotic with organisms other than testate amoebae …………………………… 06 06a – Trophi cardate. Illoricate

……………… Lindiidae

06b – Trophi not cardate. Loricate or illoricate

…………………………… 07

07a – Trophi forcipate or hemiforcipate with medium long manubria. Illoricate

…………………………… 08

07b – Trophi not forcipate. Loricate or illoricate

…………………………… 09

08a – Trophi with hook-shaped alulae, not protrusible. Stomach and intestine mostly greenish, filled with zoochlorellae

………………… Ituridae

08b – Trophi without alulae or with differently shaped alulae, protrusible. Stomach and intestine rarely with zoochlorellae

…… Dicranophoridae

09a – Trophi incudate. Illoricate

……… Asplanchnidae

09b – Trophi otherwise. Loricate or illoricate

………………………… 10

10a – Trophi virgate

………………………… 11

10b – Trophi strictly malleate, submalleate, or malleovirgate

………………………… 16

11a – Foot and toes very long, combined longer than body

…………… Scaridiidae

11b – Foot shorter, toes variable, combined shorter than or as long as body

………………………… 12

12a – Body more or less strongly asymmetrical. Trophi very asymmetrical, asymmetry concerns all trophi elements. Foot terminally, a short single pseudosegment bearing several bristles (substyli) and elongate spine-like toe(s) of unequal length. Trunk usually cylindrical with dorsal crest (usually asymmetrical); ventral fissure absent

……… Trichocercidae

12b – Body symmetrical. Trophi usually symmetrical, sometimes weakly asymmetrical (asymmetry most pronounced in rami and unci). Toes absent or present and usually of equal length, never substyli; occasionally, a single toe. Body sacciform, fusiform, conical, etc., never with dorsal crest, or if indication of dorsal, crest lorica with ventral fissure

………………………… 13

13a – Stomach colored yellowish or brownish, with blind sacs. Body saccate to ovate, weakly loricate 13b – Stomach without blind sacs. Trunk fusiform, conical, vasiform, or cylindrical and illoricate, or ovate to beanshaped and distinctly loricate

……… Gastropodidae

…………………………… 14

4.6 Systematics 

17b – Head shield absent

14a – Corona with stiff setae and sensory palps. Strongly developed V-shaped hypopharynx muscles. Illoricate or loricate. With or without 6 sword-shaped paddles

……… Synchaetidae

14b – Not as above

………………………… 15

15a – Body conical, dorsum strongly arched, head continuous with trunk. Foot long, narrow. Single toe. Mouth opening bordered by long stiff cilia forming pseudotrochus 15b – Body more or less strongly fusiform, head usually distinctly offset. Foot usually short and broad. Toes usually paired, rarely a single toe. Corona of Notommata-type 16a – Trophi submalleate, number of unci teeth reduced, usually composed of 3 stout subequal fused teeth, shaft of manubria elongate. Foot short, a single pseudosegment, inserted ventrally on ventral plate. Toes 2 or fused (partly or completely) to 1. Usually loricate with dorsal and ventral plate separated by lateral furrows (sulci); dorsal and ventral plate not distinguishable in illoricate species 16b – Trophi strictly malleate, or malleovirgate, unci with many teeth, gradually decreasing in length, shaft of manubria not particularly elongated; when number of teeth reduced, foot absent or composed of 2–5 pseudosegments in line with trunk. Loricate or illoricate. Dorsal, ventral and/or lateral sulci present or absent 17a – Head shield present, retractile or non-retractile

………………………… 18

18a – Loricate, lorica of 1 piece with longitudinal dorsal sulcus or dorsal keel (not part of polygonal facets). Foot present

………… Mytilinidae

18b – Loricate or illoricate, without dorsal sulcus or keel (if present part of polygonal facets). Foot absent or present

………………………… 19

19a – Illoricate or very weakly loricate

………………………… 20

…… Microcodonidae 19b – Distinctly loricate 20a – Mouth set at end of shallow or deep, large funnelshaped buccal field. Corona usually with conspicuous series of tufts of long cirri. Foot distinct with 1 or 2 toes or …… Notommatidae rudimentary and toes lacking. Trunk occasionally with several protruding transversal folds; without lateral sulci. Trophi strictly malleate 20b – Mouth superficial, no large funnel-shaped buccal field. Corona usually without tufts of long cirri [except in Bryceella]. Foot distinct with 1 or 2 toes. Trunk without protruding transversal folds; usually without lateral sulci [shallow ones in Bryceella]. Trophi malleate to …………… Lecanidae malleovirgate 21a – Conspicuous lateral sulci present. Foot with 2 stout toes 21b – Lateral sulci absent, shallow, or inconspicuous. Foot absent or present; toes absent or present

………………………… 17

 261

22a – Lorica covering clearly defined head, trunk, and foot

22b – Lorica only covering trunk and occasionally foot. Foot ………… Lepadellidae absent or present

………………………… 21

………… Epiphanidae [illoricate Brachionidae may key out here]

…………… Proalidae

………… Euchlanidae

………………………… 22

……… Trichotriidae

……… Brachionidae

262 

 4 Rotifera

4.6.2.4 Key to families of Flosculariacea

4.6.3 Characterization of families

01a – Body loricate. Free-swimming 01b – Body illoricate. Free-swimming or sessile in tubes of mucus, pellets, or rigid material

Subclass Bdelloidea Hudson, 1884 Order Adinetida Melone & Ricci, 1995 Family Adinetidae Hudson & Gosse, 1886 (Figs. 4.49 A and 4.53 A, B) Body fusiform to vermiform, mostly compressed dorsoventrally. Esophagus long, trophi deep, not protrusible. Stomach tube-shaped with lumen. Corona a ventral ciliated field with rake of teeth at its base (Adineta) or at its lateral sides (Bradyscela), trochi absent; rostrum terminal, not retractile. Eyespots usually absent. Foot narrow, 3 toes, 2 spurs (Adineta), or foot short, plump, distal foot pseudosegment disc-shaped with papillae, spurs absent (Bradyscela). Proximal and distal minor teeth of uncus always present. Oviparous, rarely viviparous. Gliding movement. Feeding by scraping and browsing. Two genera: Adineta Hudson & Gosse, 1886 (18 spp.) and Bradyscela Bryce, 1910 (2 spp.). Limnoterrestrial, in aerophytic mosses, leaf litter, soil, Sphagnum, hygropsammon, and littoral submerged vegetation.

…………… Testudinellidae

…………………………………… 02

02a – Body with 6 arm-like appendages with fanwise arranged setae. Freeswimming ………………… Hexarthridae 02b – Body without arm-like appendages. Free-swimming or sessile …………………………………… 03 03a – Apical field domeshaped. Body with or without 2 long lateral movable setae and 1 caudal seta. Foot absent. Free-swimming ………… Trochosphaeridae 03b – Apical field not dome-shaped. Movable setae always absent. Foot present. Free-swimming or sessile …………………………………… 04 04a – Corona round, horseshoe- to U-shaped. Teeth of left uncus longer than those of right one. Free-swimming, often colonial

………………… Conochilidae

04b – Corona round, heart-shaped, or with 2–8 lobes, not horseshoe- or U-shaped. Teeth of left and right uncus equally long. Adults sessile or swimming in colonies

……………… Flosculariidae

4.6.2.5 Key to families of Collothecacea 01a – Corona funnelshaped without lobes or cilia …………………… Atrochidae 01b – Corona with lobes, tentacles, or points bearing long setae and cilia ……………… Collothecidae

Order Philodinavida Melone & Ricci, 1995 Family Philodinavidae Harring, 1913 (Fig. 4.53 C–E) Esophagus short. Stomach tube-shaped with lumen. Corona poorly developed, reduced to small ciliated field, trochi absent or small, cheeks present, stout non-retractable rostrum, short dorsal antenna (Ricci & Melone 1998a). The trophi are close to the mouth opening, and extruded when animal is feeding. Proximal minor teeth of uncus plate reduced or absent. Foot with 4 toes, and 2 spurs. Oviparous. The aberrant carnivorous Abrochtha carnivora has a long pharynx that can be widened to become a large funnel, used in combination with the mouth cone to envelop the prey (rotifers); its trophi are not protrusible, and the uncus shows numerous minor teeth (Ricci et al. 2001). Three genera: Abrochtha Bryce, 1910 (5 sp.), Henoceros Milne, 1916 (2 spp.), and Philodinavus Harring, 1913 (2 sp.). Benthic-periphytic in stagnant and running waters. Order Philodinida Melone & Ricci, 1995 Family Habrotrochidae Bryce, 1910 (Fig. 4.53 F–I) Body fusiform to vermiform. Esophagus long, trophi deep, not protrusible. Stomach without recognizable lumen, with round pellets in its wall. Corona frontal, with small but well developed trochi, mostly elevated on pedicels. Upper lip not extending beyond ciliary disc, or hoodshaped. Eyespots present or absent. Foot with 2 spurs of variable length. Toes present (2, 3, or 4) or absent. Proximal and distal minor teeth of uncus plate always present.

4.6 Systematics 

 263

Fig. 4.53: Bdelloidea. (A) Adineta vaga, (B) Bradyscela clauda, (C) Abrochtha intermedia, (D) Henoceros falcatus, (E) Philodinavus paradoxus, (F) Otostephanos monteti, (G) Scepanotrocha rubra, (H) Habrochtha serpens, (I) Habrotrocha pusilla textrix, (J) Anomopus telphusae, (K) Ceratotrocha cornigera, (L) Didymodactylos carnosus, (M) Dissotrocha a. aculeata, (N) Embata laticeps, (O) Macrotrachela multispinosa, (P) Mniobia orta, (Q) Philodina citrina, (R) Pleuretra brycei, (S) Rotaria neptunia, (T) Rotaria sordida fimbriata, (U) Rotaria citrina, and (V) Zelinkiella synaptae. (After different authors.)

264 

 4 Rotifera

Oviparous, rarely viviparous. Vortex feeding. Some Habrotrocha species form secretions and more or less bottle-shaped shells or occupy shells of testate amoebae (Centropyxis, Nebela) or the cups (amphigastrae) under the leaflets of hepatic mosses (Frullania, Lejeunia); others reported from empty Sphagnum and plant (Typha) cells (Donner 1950). Three genera: Habrotrocha Bryce, 1910 (128 sp.), Otostephanos Milne, 1916 (13 spp.), and Scepanotrocha Bryce, 1910 (11 spp.). Limnoterrestrial, in aerophytic mosses and lichens, leaf litter, wet soil, Sphagnum, hygropsammon, and littoral submerged vegetation; occasionally epizoic on insect larvae, molluscs, etc. Family Philodinidae Ehrenberg, 1838 (Figs. 4.49 B, C and 4.53 J–V) Shape of body variable, fusiform, vermiform, cylindrical, etc. Integument soft or stiffened and often ornamented with grooves, spines, platelets, warts, etc. Esophagus long, trophi deep, not protrusible. Tube-shaped stomach with lumen. Corona frontal, well developed, with trochi mostly elevated on pedicels. Rostrum retractile. Eyespots present or absent. Foot with 2, 3, or 4 toes, or with adhesive disc and toes absent. Two spurs of variable length. Proximal and distal minor teeth of uncus plate always present. Vortex feeding. Oviparous and ovoviviparous. Eleven genera: Anomopus Piovanelli, 1903 (2 spp.), Ceratotrocha Bryce, 1910 (4 spp.), Didymodactylos Milne, 1916 (1 sp.), Dissotrocha Bryce, 1910 (34 spp.), Embata Bryce, 1910 (5 spp.), Macrotrachela Milne, 1886 (97 spp.), Mniobia Bryce, 1910 (49 spp.), Philodina Ehrenberg, 1830 (50 spp.), Pleuretra Bryce, 1910 (14 spp.), Rotaria Scopoli, 1777 (27 spp.), and Zelinkiella Harring, 1913 (1 sp.). Limnoterrestrial, in aerophytic mosses and lichens, Sphagnum, leaf litter, soil, dung; in benthos, periphyton, hygropsammon and hydropsammon of freshwaters, rarely planktonic; a few species in brackish and marine environments. Several species epizoic on crustaceans: Mniobia branchicola on gill plates of terrestrial isopods (Ligidium); Embata commensalis, E. laticeps, E. parasitica, Rotaria magnacalcarata, R. murrayi, R. socialis on freshwater isopods and amphipods, e.g., Asellus, Gammarus; Anomopus telphusiae and Embata parasitica in gill chamber of freshwater crayfish, e.g., Astacus, Austropotamobius, and crab (Potamon), and Anomopus chasmagnathi in gill chamber of brackish-water crab (Chasmagnathus); Zelinkiella synaptae on body surface and tentacles of sea cucumbers (Synaptidae) and polychaetous annelids (Terebellidae); several freshwater species have been reported epibiotic on insect larvae, molluscs, etc.

Subclass Monogononta Plate, 1889 Order Ploima Hudson & Gosse, 1886 Family Asciaporrectidae De Smet, 2006 (Fig. 4.54 A) Illoricate. Body fusiform with offset head, trunk, and foot. Foot short to fairly long, broad. Two short toes. Trophi hemiforcipate organized for grasping, with long rod-shaped manubria bearing axe-shaped head, long fulcrum, and 2–3 large uncinal and 3–5 subuncinal teeth pivoting on the broad tips of the flat rami. Oviparous. Three species: the free-living benthic-periphytic Asciaporrecta hyalina (formerly Pleurotrocha hyalina), and A. difflugicola and A. arcellicola inhabiting shells of live testate amoebae of the genus Difflugia and Arcella, respectively. Probably parasites, feeding on the protoplasm of their hosts (De Smet 2006). Family Asplanchnidae Eckstein, 1883 (Fig. 4.54 B, C) Body sac-shaped, sometimes with lateral protuberances, head differentiated, foot and toes present or absent, hyaline. Illoricate. Corona Asplanchna type, a single ring of cilia. Eyespot cerebral or absent. With or without intestine and anus. Vitellarium spherical, saccate, or horseshoe-shaped. Gastric glands on esophagus. Trophi incudate. Oviparous and ovoviviparous. Three genera: Asplanchna Gosse, 1850 (9 spp.), Asplanchnopus Guerne, 1888 (4 spp.), and Harringia de Beauchamp, 1912 (2 spp.). Planktonic (Asplanchna), semi-planktonic (Asplanchnopus), and benthic-periphytic (Harringia) species. Family Birgeidae Harring & Myers, 1924 (Fig. 4.54 D) Illoricate. Body plump, vase-shaped, head separated from trunk by neckfold. Foot extremely slender, 3 pseudosegments. Two lanceolate short toes. Corona frontal, circumapical band interrupted dorsally, passing laterally to auricle-like area of strong cilia, buccal field close-set with short cilia. Stomach with 10 blind sacks, stomach wall crowded with zoochlorellae. Gastric glands absent. Large eyespot ventrally on small brain. Trophi unique, highly specialized with large, hooked pseudunci expanded at their proximal ends into broad laminae; at rest hooks protrude slightly through mouth opening. Fulcrum long, rod-shaped, directed toward mouth. Rami elongate, narrow, attenuated toward tiops. Unci with 4–5 linear teeth. Manubria heart-shaped with very short cauda. Monotypic family, with a single species: Birgea enantia Harring & Myers, 1922, reported from freshwater swampy pond (Wisconsin and New Jersey, USA).

4.6 Systematics 

Family Brachionidae Ehrenberg, 1838 (Figs. 4.50 A–C and 4.54 E–L) Loricate, occasionally very weakly loricate. Lorica usually more or less dorsoventrally depressed, angular, rectangular, or hexagonal; anterior margins of lorica often with spines, caudally with or without spines or extensions; head and foot retractable in lorica. Foot present or absent, if present long and wrinkled or with pseudosegments; 2 small toes. Corona Brachionus type, frontal with reduced ciliation, pseudotrochus with 3‒5 humps with short cilia on top and cirri between. Eyespots frontal, cerebral, or lacking. Trophi malleate. Oviparous. Seven genera: Anuraeopsis Lauterborn, 1900 (5 spp.), Brachionus Pallas, 1766 (55 spp.), Kellicottia Ahlstrom, 1938 (2 spp.), Keratella Bory de St. Vincent, 1822 (45 spp.), Notholca Gosse, 1886 (42 spp.), Plationus Segers, Murugan & Dumont, 1993 (3 spp.), and Platyias Harring, 1913 (3 spp.).

 265

Planktonic and semi-planktonic in littoral, temporarily attached to substrate; Brachionus rubens and B. sessilis epizoic on cladocerans, occasionally on Corixa sp. Freshwater, a few species in brackish and marine environments. Family Clariaidae Kutikova, Markevich & Spirodonov, 1990 (Fig. 4.54 M) Illoricate. Body elongate fusiform, anterior half with 16 pseudosegments, showing longitudinal striations and finely denticulate distal margin. Toes very long, weakly inflated. Trophi aberrant with thin elongate forceps-like rami, inner margin of rami lined with numerous, and loosely bound short teeth, rami with set of 4 small apical teeth. Single uncus tooth, short manubria, and long fulcrum. Monotypic family, with a single species: Claria segmentata Kutikova, Markevich & Spirodonov, 1990, parasitic in earthworm Pheretima modigliani Rosa, 1889 (Megascolecidae).

Fig. 4.54: Monogononta. (A) Asciaporrecta arcellicola, (B) Asplanchna priodonta, (C) Harringia eupoda, (D) Birgea enantia, (E) Brachionus mirabilis, (F) Platyias quadricornis, (G) Plationus patulus, (H) Keratella serrulata, (I) Anuraeopsis fissa, (J) Kellicottia longispina, (K) Notholca kozhovi, (L) Notholca olchonensis, (M) Claria segmentata, and (N) Cotylegaleata perplexa. (After different authors.)

266 

 4 Rotifera

Family Cotylegaleatidae De Smet, 2007 (Fig. 4.54 N) Loricate. Head relatively small, covered by non-retractile head shield, offset from rectangular, and slightly dorsoventrally compressed trunk. Foot long, 4 pseudosegments, distal by one longest, bearing 2 toes, and 2 ventrolateral spurs. Corona reduced, ventral; mouth opening surrounded by large, shallow cup-shaped structure with stiffened wall. Trophi modified virgate, with small unci composed of 5–6 teeth, short manubria, long fulcrum, and 2 relatively long stilletto-shaped epipharyngeal elements. Monotypic family, with a single species: Cotylegaleata perplexa De Smet, 2007, found in benthos of freshwater lake at 8–9 m depth; ectoparasite? (De Smet 2007). Family Dicranophoridae Harring, 1913 (Figs. 4.50 E–G and 4.55 A–F) Illoricate or semi-loricate. Body elongate cylindrical, fusiform, usually with offset head, trunk, and foot. Head usually with dorsofrontal rostrum. Trunk mostly with dorsal fold or tail, covering foot more or less. Foot and toes (2) usually short. Corona of Dicranophorus type, ventral or oblique, often 2 lateral ciliary tufts of longer cilia; corona absent in parasitic genus Balatro. Eyespots frontal, 2, rarely 4, or absent. Trophi forcipate, protrusible. Oviparous, ovoviviparous. The family comprises 19 genera: Albertia Dujardin, 1838 (7 spp.), Aspelta Harring & Myers, 1928 (21 spp.), Balatro Claparède, 1867 (4 spp.), Dicranophoroides De Smet, 1997 (4 spp.), Dicranophorus Nitzsch, 1827 (52 spp.), Donneria De Smet, 2003 (1 sp.), Encentrum, Ehrenberg, 1838 (subgenera Encentrum, Isoencentrum, and Pseudencentrum De Smet, 1997, 105 spp.), Dorria Myers, 1933, Erignatha Harring & Myers, 1928 (5 spp.), Glaciera Jersabek, 1999 (1 sp.), Inflatana Kutikova, 1985 (1 sp.), Kostea De Smet, 1997 (1 sp.), Myersinella Wiszniewski, 1936 (4 spp.), Paradicranophorus Wiszniewski, 1929 (5 spp.), Parencentrum Wiszniewski, 1936 (2 spp), Pedipartia Myers, 1937 (1 sp.), Streptognatha Harring & Myers, 1928 (1 sp.), Wierzejskiella Wiszniewski, 1934 (2 spp.), Wigrella Wiszniewski, 1932 (2 spp.). Benthic-periphytic, and psammic in freshwater, brackish, and marine environments; the most diverse rotifer family in saline waters. Several species live epizoically on freshwater crustaceans. Encentrum grande and E. kulmatyckii live on the thoracal and abdominal appendages, and the gills of Asellus aquaticus and Gammarus pulex; Dicranophorus siedleckii is also found on the gills and pleopods of G. pulex. Dicranophorus hauerianus and D. cambari inhabit the branchial cavities of several species of freshwater crayfish. Nearly all

Albertia and Balatro species are obligatory parasites of terrestrial and aquatic oligochaetes. Albertia species live in the intestine of Naididae, Lumbricidae, and Lumbriculidae; Balatro is parasitic in the intestine or on the exterior of Lumbriculidae and Enchytraeidae. Encentrum kozminskii lives ectoparasitic on the skin and gills of carp (Cyprinus carpio). Family Epiphanidae Harring, 1913 (Fig. 4.55 G-J) Illoricate, occasionally weakly stiffened. Body fusiform, cylindrical, or conical to vase-shaped. Foot short to long, mostly indistinctly offset from trunk. Toes short and small, 2 or 1. Mouth set in funnel-shaped ventral buccal field with short cilia. Corona of Euchlanis-Brachionus type, a circumapical band of cilia; apical field with tufts of cilia, buccal field with short cilia; apical field can be elongated to form broad proboscis. Eye(s) pigmented or colorless; a single cerebral eye, or 2 eyes on proboscis. Trophi malleate. Oviparous or ovoviviparous. Five genera: Cyrtonia Rousselet, 1894 (1 sp.), Epiphanes Ehrenberg, 1832 (6 spp.), Mikrocodides Bergendal, 1892 (3 spp.), Proalides de Beauchamp, 1907 (2 spp.), and Rhinoglena Ehrenberg, 1853 (4 spp.). Planktonic and semi-planktonic, periphytic. Freshwater and brackish waters. Epiphanes daphnicola (formerly Proales daphnicola) is epibiotic on Daphnia spp., rarely on other cladocerans, copepods, amphipods, and oligochaetes. Family Euchlanidae Ehrenberg, 1838 (Fig. 4.55 K, N) Loricate. Body usually oval; trunk usually depressed, composed of 2‒3 plates connected by thin membranes forming longitudinal sulci; head offset from trunk by constriction, foot well-differentiated with 2 stout, more or less long toes. Single cerebral eyespot. Corona Euchlanis type, with interrupted ciliary band, apical field with several tufts of cilia. Trophi malleate. Oviparous. The family comprises 5 genera: Beauchampiella Remane, 1929 (1 sp.), Dipleuchlanis de Beauchamp, 1910 (3 spp.), Diplois Gosse, 1886 (1 sp.), Euchlanis Ehrenberg, 1832 (14 spp.), Tripleuchlanis Myers, 1930 (2 spp.). Benthic-periphytic, littoral of freshwaters, rarely brackish and marine. Family Gastropodidae Harring, 1913 (Fig. 4.55 L, M) Loricate. Body ovate to sac-shaped, more or less compressed laterally and/or dorsoventrally. Lorica smooth, transparent, with or without longitudinal folds. Foot absent or present, more or less placed ventrally, retractable, usually annulated. Occasionally surrounded by mucus

4.6 Systematics 

 267

Fig. 4.55: Monogononta. (A) Albertia vermiculus, (B) Balatro calvus, (C) Encentrum plicatum, (D) Dicranophorus hercules, (E) Dicranophorus cambari, (F) Dicranophoroides caudatus, (G) Epiphanes brachionus, (H) Proalides tentaculatus, (I) Rhinoglena fertoeensis, (J) Mikrocodides robustus, (K) Euchlanis meneta, (L) Ascomorpha saltans, (M) Gastropus stylifer, (N) Beauchampiella eudactylota, (O) Lecane sylvae, (P) Lecane satyrus, (Q) Colurella halophila, (R) Squatinella longispinata, and (S) Lepadella ovalis. (After different authors.)

268 

 4 Rotifera

sheath. Stomach lobate or with blind sacks containing darkcolored defecation pellets; stomach wall often with zoochlorellae. Corona with well developed cingulum, apical field with sensory organs, e.g., styli, palps, tufts of bristles. Cerebral eyespot present. Trophi virgate. Oviparous. The family comprises 2 genera: Ascomorpha Perty, 1850 (9 spp.) and Gastropus Imhof, 1888 (3 spp.). Planktonic and semi-planktonic in littoral vegetation of freshwaters, rarely in brackish habitats. Family Ituridae Sudzuki, 1964 (Fig. 4.56 A) Illoricate. Body fusiform. Head more or less widening laterally; usually neck distinct. Foot short, 2 pseudosegments. Stomach with blind sacks; stomach and intestine with green zoochlorellae. Gastric glands absent. Corona reduced to a buccal field with 2 lateral, non-retractable ciliated auricles. Two frontal and single cerebral eyespot. Trophi similar to forcipate type of genus Dicranophorus. Oviparous. A single genus Itura Harring & Myers, 1928, with 6 species. Benthic-periphytic in freshwater habitats, occasionally in brackish environment. Family Lecanidae Remane, 1933 (Fig. 4.55 O, P) Loricate, rarely illoricate. Lorica cylindrical when extended; outline truncate-oval or shield-shaped; flattened dorsoventrally when contracted; with dorsal and ventral plates separated by lateral sulcus. Head opening of lorica slit-like, broad and shallow, often with the lateral corners forming angles or spines. Anterior margin of dorsal and ventral plate straight or with more or less deep sinus. Foot subdistally, very short, a single pseudosegment. Two separate or partly fused toes, or a single toe; tips of toes simply acute, or with completely or incompletely separated claws or incompletely separated pseudoclaws. Corona Euchlanis type. Single cerebral eyespot. Trophi submalleate. Oviparous. A single genus Lecane Nitzsch, 1827, with about 170 spp. Benthic-periphytic in littoral of freshwaters, several species in brackish environment, rarely marine. Lecane branchicola lives in the gill chambers of the freshwater crab Potamon fluviatile. Family Lepadellidae Harring, 1913 (Fig. 4.55 Q–S) Loricate. Lorica compressed dorsoventrally, composed of firmly joined dorsal and ventral plate, or a laterally compressed plate with ventral slit; lorica spines may be present. Head usually with retractile or non-retractile, semi-circular shield or membranous cap. Foot with 3‒4 pseudosegments. Toes slender, acutely pointed, occasionally partly or com-

pletely fused. Corona Euchlanis-like, frontal, a single ciliary band. Two lateral eyespots, or eyes absent. Trophi malleate, occasionally modified. Oviparous. Five genera: Colurella Bory de St. Vincent, 1824 (21 spp.), Halolepadella De Smet, 2012 (1 sp.), Lepadella Bory de St. Vincent, 1826 (subgenera Heterolepadella Bartoš, 1955; Xenolepadella Hauer, 1926, 80 spp.), Paracolurella Myers, 1936 (2 spp.), and Squatinella Bory de St. Vincent, 1826 (6 spp.). Littoral, benthic-periphytic, occasionally in plankton and psammon. Freshwater, rarely in brackish or marine waters. Lepadella astacicola, L. borealis, L. branchicola, L. lata, and L. parasitica live in the gill cavities of freshwater crayfish (Astacus spp.). Family Lindiidae Harring & Myers, 1924 (Fig. 4.56 B) Illoricate. Body vermiform or fusiform, head, trunk, and foot generally distinct; tail usually present. Foot short, a single rudimentary pseudosegment or 2–3 pseudosegments. Two small, short toes. Corona Notommata type, a simple narrow ciliated field, ventral or strongly oblique; 2 lateral ciliary tufts, usually on auricles. Cerebral eye present, rarely absent. Trophi cardate, usually with complicated accessory trophi elements. Oviparous and ovoviviparous. A single genus, Lindia Dujardin, 1841 (subgenus Halolindia Harring & Myers, 1924, Neolindia Segers, 2002) with 16 species. Benthic-periphytic, among cyanobacteria, littoral, also in psammon and Sphagnum. Mainly freshwater, 4 brackish-marine species. Family Microcodonidae Hudson & Gosse, 1886 (Fig. 4.56 C) Illoricate. Body conical, tapering to long foot with single lanceolate toe. Corona large, heart-shaped; paracingulum with dorsal and ventral interruption; mouth opening central, bordered by long stiff membranelles forming pseudotrochus. Single, large red cerebral eyespot. Trophi virgate, manubria and unci reduced; epipharynx plate-shaped, often colored red to purple. Oviparous. A single genus, Microcodon Ehrenberg, 1830 (1 sp.). Semi-planktonic in vegetation rich littoral, occasionally in Sphagnum. Family Mytilinidae Harring, 1913 (Figs. 4.50 I and 4.56 D, E) Loricate. Lorica thick or thin, often ornamented, elongate, in cross section mostly triangular or rhomboid; ventral plate and dorsolateral plates firmly fused; often with longitudinal dorsal furrow flanked by lateral ridges, or single,

4.6 Systematics 

 269

Fig. 4.56: Monogononta. (A) Itura viridis, (B) Lindia pallida, (C) Microcodon clavus, (D) Mytilina brevispina, (E) Lophocharis rubens, (F) Rousseletia corniculata, (G) Taphrocampa selenura, (H) Cephalodella misgurnus, (I) Monommata maculata, (J) Notommata pachyura, (K) Sphyrias lofuana, (L) Enteroplea lacustris, (M) Tylotrocha monopus, (N) Bryceella stylata, (O) Wulfertia ornata, (P) Proalinopsis caudatus, (Q) Proales reinhardti, (R) Proales doliaris, and (S) Scaridium longicaudum. (After different authors.)

270 

 4 Rotifera

rarely 3 ridges; anterior lorica margin often with spines. Foot short, 2‒3 pseudosegments. Two toes. Corona Euchlanistype. Single cerebral eye. Trophi malleate. Oviparous. Two genera: Lophocharis Ehrenberg, 1838 (6 spp.), Mytilina Bory de St. Vincent, 1826 (12 spp.). Benthic-periphytic, occasionally in plankton; freshwater, rarely in brackish environments. Family Notommatidae Hudson & Gosse, 1886 (Figs. 4.50 H and 4.56 F–M) A taxonomically unsatisfactory assemblage of diverse taxa (see Nogrady et al., 1995). Illoricate to partly loricate. Body plan various, mostly fusiform, with offset head, trunk, and foot; often with tail. Foot usually short, composed of single or few pseudosegments. Toes short to long. Corona Notommata, Dicranophorus, and Asplanchna type, often with ciliated auricles. Eyespots present or absent, frontal or cerebral. Trophi virgate. Oviparous. The family comprises 17 genera: Cephalodella Bory de St. Vincent, 1826 (191 spp.), Dorystoma Harring & Myers, 1922 (1 sp.), Drilophaga Vejdovsky, 1883 (3 spp.), Enteroplea Ehrenberg, 1830 (1 sp.), Eosphora Ehrenberg, 1830 (6 spp.), Eothinia Harring & Myers, 1922 (7 spp.), Monommata Bartsch, 1870 (18 spp.), Notommata Ehrenberg, 1830 (53 spp.), Pleurotrocha Ehrenberg, 1830 (8 spp.), Pleurotrochopsis Berzins, 1973 (2 spp.), Pseudoharringia Fadeew, 1925 (2 spp.), Pseudoploesoma Myers, 1938 (1 sp.), Resticula Harring & Myers, 1924 (7 spp.), Rousseletia Harring, 1913 (1 sp.), Sphyrias Harring, 1913 (1 sp.), Taphrocampa Gosse, 1851 (3 spp.), Tylotrocha Harring & Myers, 1922 (1 sp.), and Pleurata Nogrady & Pourriot, 1995 (6 spp.). Position of Pourriotia werneckii (formerly Proales werneckii) and P. carcharodonta uncertain (De Smet 2003, 2009). Most species benthic-periphytic or semi-planktonic in littoral of freshwater habitats, rarely in brackish waters; few marine species. Cephalodella crassipes lives in the branchial chambers of freshwater crayfish (Astacus spp.). Several species are ectoparasites or endoparasites. Ectoparasitic: Cephalodella parasitica on oligochaetes; Drilophaga bucephalus and D. delagei on oligochaetes and leeches. Endoparasitic: Cephalodella volvocicola in Volvox colonies; C. edax and Pleurata uroglenae in Uroglena volvox; Pourriotia spp. in Vaucheria spp. and Dichotomosiphon tuberosus. Family Proalidae Harring & Myers, 1924 (Figs. 4.50 D and 4.56 N–R) A taxonomically unsatisfactory assemblage of diverse taxa. Illoricate to semi-loricate. Body cylindrical, fusiform, or swollen. Head, trunk, and foot usually clearly defined; foot short to very long. Two toes or single. Corona simple, mostly supraoral, with oblique buccal field and

lateral parts of circumapical band. Eyespot(s) cerebral, frontal, lateral, or absent. Trophi malleate, virgate, or a modification of one of these types; epipharynx usually present. Oviparous. Four genera: Bryceella Remane, 1929 (3 spp.), Proales Gosse, 1886 (43 spp.), Proalinopsis Weber, 1918 (6 spp.), and Wulfertia Donner, 1943 (3 spp.). Benthic-periphytic, planktonic, and semi-planktonic, psammic. In freshwater, brackish, and marine environments. Several parasitic species; ectoparasitic: Proales christinae and P. gonothyraeae on hydroid polyps, P. paguri on Eupagurus bernhardus; endoparastic: P. gigantea in eggs of several species of freshwater snail, P. parasita in colonial algae (Volvox, Uroglena, Uroglenopsis). Family Scaridiidae Manfredi, 1927 (Fig. 4.56 S) Loricate or semi-loricate. Body fusiform to cylindrical, with stiffened longitudinal plates. Foot very long, 3 pseudosegments, with strong transversally striated muscles. Two very long, equal toes. Corona oblique, a dorsally interrupted band of cilia, and ventral trochus with stiff cilia surrounding mouth opening. Unci projecting through mouth opening. Eyes absent. Mastax with apical red spot. Trophi modified virgate; epipharynx weak, consisting of horseshoe-toothed anterior part and pair of elongate dorsal projections. Oviparous. A single genus Scaridium Ehrenberg, 1830 (7 spp.). Periphytic in freshwaters. Family Synchaetidae Hudson & Gosse, 1886 (Fig. 4.57 A–D) Illoricate and loricate. Body of different shape, rectangular, sac-shaped, bell-shaped, ovate. Anterior of genus Polyarthra with 6 serrated paddles. Head usually set off from trunk by fold or fissure, with or without ciliated lateral auricles. Rigid lorica ornamented. Foot absent, short, or long and annulated. Toes short, 2, single, or absent. Corona Asplanchna type, frontal, with sensory setae, tufts of cilia, ciliated palps, etc. Cerebral eye usually present, single or paired. Trophi virgate, hypopharynx muscle mostly prominent. Oviparous and ovoviviparous. Three genera: Ploesoma Herrick, 1885 (7 spp.), Polyarthra Ehrenberg, 1834 (11 spp.), and Synchaeta Ehrenberg, 1832 (39 spp.). Mostly planktonic, rarely semi-planktonic and periphytic in freshwaters. Genus Synchaeta with several marine species. Family Tetrasiphonidae Harring & Myers, 1924 (Fig. 4.57 M) Illoricate, but integument stiff. Body cylindrical, head, and trunk not separated by transversal folds. Foot short, 2 pseudosegments. Two fairly long, slender, acute

4.6 Systematics 

 271

Fig. 4.57: Monogononta. (A) Polyarthra major, (B) Synchaeta pectinata, (C) Synchaeta bicornis, (D) Ploesoma lenticulare, (E) Trichocerca weberi, (F) Trichocerca cylindrica, (G) Trichocerca rattus, (H) Ascomorphella volvocicola, (I) Macrochaetus multispinosus, (J) Trichotria truncata, (K) Conochilus hippocrepis, (L) C. natans, (M) Tetrasiphon hydrocora, (N) Octotrocha speciosa, (O) Sinantherina spinosa, (P) Limnias melicerta, (Q) Pompholyx sulcata, (R) Trochosphaera aequatorialis, (S) Filinia limnetica, (T) Atrochus tentaculatus, and (U) Collotheca coronetta. (After different authors.)

272 

 4 Rotifera

toes. Dorsal antenna double, 2 fairly large, conical tubules with tuft of long sensory setae. Lateral antennae near posterior, very long, tubular with few very long setae. Corona oblique, ciliation very faint, anterolaterally 2 V-shaped, concave ciliated appendages; post-oral margin projecting as fairly prominent chin. Single cerebral eyespot. Connection between stomach and intestine narrow, surrounded by wreath of ca. 6 spherical glands. Vitellarium strongly elongate, cylindrical, wound around intestine, with 20–30 nuclei. Trophi unique. Rami elongate lyrate, recurved dorsally, with large alulae. Fulcrum short, lamellar. Unci with single tooth, and thin crescent-shaped basal lamella. Manubria complex, rod-shaped, curved, with appendages, connected to unci and rami. Accessory trophi elements present. Adults usually covered with firm jelly case. Oviparous. Monotypic family, with single species: Tetrasiphon hydrocora Ehrenberg, 1840 (1 sp.) Periphytic in soft acid wates, Sphagnum pools. Family Trichocercidae Harring, 1913 (Fig. 4.57 E–H) Loricate, rarely illoricate. Body ovate, cylindrical, fusiform, often twisted and asymmetrical, often with dorsal crest. Head usually with closing plates. Foot present and short, or absent. Toes if present 1‒2, short to long, seta-like, usually of different length and left longest; often bristles or substyli at base of toes. Single cerebral eyespot. Corona Asplanchna type, frontal, circumapical band weakly developed. Trophi virgate, strongly asymmetrical. The family comprises 3 genera: Ascomorphella Wiszniewski, 1953 (1 sp.), Elosa Lord, 1891 (2 spp.), Trichocerca Lamarck, 1801 (75 spp.). Oviparous. Planktonic and periphytic, rarely psammic; predominantly freshwater, a few marine species only. Ascomorphella volvocicola lives endoparasitic in colonies of Volvox aureus, V. globator, and Uroglena volvox. Family Trichotriidae Harring, 1913 (Figs. 4.50 J and 4.57 I, J) Loricate. Lorica very rigid, ornamented with pustules and spines; spines movable in Macrochaetus; often with plates covering head. Head, trunk, and foot clearly defined. Foot usually 2‒4 pseudosegments. Toes 2, often very long. Single cerebral eyespot. Corona frontal, weakly developed, incomplete ring of cilia. Trophi malleate. Oviparous. The family comprises 3 genera: Macrochaetus Perty, 1850 (11 spp.), Trichotria Bory de St. Vincent, 1827 (7 spp.), and Wolga Skorikov, 1903 (1 sp.). Periphytic in littoral vegetation and Sphagnum puddles; freshwater, occasionally in brackish waters.

Order Flosculariacea Harring, 1913 Family Conochilidae Harring, 1913 (Figs. 4.51 A and 4.57 K–L) Illoricate. Adult females free-swimming, solitary, or colonial (5 to >400 individuals per colony) and clustered within gelatinous mass produced by pedal glands. Body conical, long, unsegmented foot, toes absent. Intestine U-shaped. Corona Conochilus type, horseshoe-shaped, U-shaped, or circular. Antennae apically inside coronal field or dorsally outside coronal field. Two dorsal eyes beneath corona. Trophi malleoramate, weakly to very asymmetrical; left uncinal teeth longer. Oviparous, ovoviviparous. Two genera: Conochilus Ehrenberg, 1834 (type genus), and subgenus Conochiloides Hlava, 1904 (6 spp.), and Conochilopsis Segers & Wallace, 2001 (1 sp.). Planktonic, freshwater, rarely in brackish water. Literature: Segers & Wallace (2001). Family Flosculariidae Ehrenberg, 1838 (Figs. 4.51 C and 4.57 N, O) Illoricate. Adult females sessile, solitary in tubes of mucus, pellets, or rigid material, or free-swimming in spherical colonies. Larvae free-swimming, foot with ciliated cup. Body conical, foot unsegmented, long to very long, often with pedicel. Dorsal antenna small to very long, single or paired. Elongate, hook-shaped, etc. stiff lorica elements in neck region present or absent. Corona Hexarthra type, heart-shaped, circular, or 2- to 8-lobed. Trophi malleoramate, symmetrical. Oviparous. The family comprises 9 genera: Beauchampia Harring, 1913 (1 sp.), Floscularia Cuvier, 1798 (10 spp.), Lacinularia Schweigger, 1820 (8 spp.), Lacinularoides Meksuwan, Pholpunthin & Segers (1 sp.), Limnias Schrank, 1803 (6 spp.), Octotrocha Thorpe, 1893 (1 sp.), Pentatrocha Segers & Shiel, 2008, Ptygura Ehrenberg, 1832 (28 spp.), and Sinantherina Bory de St Vincent, 1826 (6 spp.). Attached to submerged aquatic vegetation, Sphagnum, and other substrates, and planktonic (Lacinularia, Sinantherina, Ptygura libera). Freshwater, rarely brackish, or marine. Family Hexarthridae Bartoš, 1959 (Fig. 4.51 D, E) Illoricate. Body conical, with 6 thick arm-like appendages: 1 dorsal, 1 ventral, 2 laterodorsal, and 2 lateroventral, bearing pinnate bristles arranged fanwise at their tips; bristles inserted singly or in pairs; ventral arm longest with series of lateral spines. Foot absent. Some species with 2 dorsocaudal club-shaped appendages bearing distal tuft of cilia or caudal spine. Dorsal antenna on prominence above dorsal arm; lateral antennae on ventral arms. Corona Hexarthra type, an undulate double band of cilia; in some species, with outward bend ventral lip. Two

4.7 Biogeography 

red eyespots on apical field ventrally near corona. Trophi malleoramate. Oviparous. A single genus, Hexarthra Schmarda, 1854 (13 spp.). Planktonic in freshwater, brackish, marine, and inland saline waters. Family Testudinellidae Harring, 1913 (Fig. 4.57 Q) Loricate. Body more or less oval, more or less compressed dorsoventrally; head and trunk differentiated, foot absent or long, cylindrical ending in ciliated cup; lorica hyaline, usually smooth; anterodorsal margin often with median expansion; anteroventral margin with median incision. Foot opening ventral, from median to terminal. Lateral and dorsal antennae more or less in line, usually near middle of lorica. Corona Hexarthra type, a simple circumapical band of cilia. Two eyespots, sometimes absent. Trophi malleoramate. Oviparous. The family comprises 3 genera: Anchistestudinella Bērziņš, 1973 (1 sp.), Pompholyx Gosse, 1851 (3 spp.), Testudinella Bory de St. Vincent, 1826 (45 spp.). Benthic-periphytic, semi-planktonic in littoral. Predominantly freshwater, but also in brackish and marine environment. Testudinella caeca and T. elliptica periphytic but also epizoic on Asellus aquaticus. Family Trochosphaeridae Harring, 1913 (Figs. 4.51 B and 4.57 R, S) Illoricate. Body spherical or sac-shaped, with or without 2 movable anterolateral setae and 1 or 2 posterior setae. Foot absent. Dorsal antenna reduced, posterior to dorsal gap. Lateral antennae occasionally on papillae. Corona simple, a ciliary band with dorsal gap and ciliated buccal field; ventrally drawn out into more or less distinct lip. Apical field bare, domed. Two anterior red eyespots. Trophi malleoramate. Oviparous, ovoviviparous. Three genera: Filinia Bory de St. Vincent, 1824 (15 spp.), Horaella Donner, 1949 (2 spp.), and Trochosphaera Semper, 1872 (2 spp.). Planktonic, freshwater, and occasionally brackish waters. Order Collothecacea Harring, 1913 Family Atrochidae Harring, 1913 (Figs. 4.52 A–C and 4.57 T) Illoricate. Body elongate conical, anterior part funnelshaped, distinctly offset or not by constriction. Sessile or creeping. Foot unsegmented, long with small adhesive disc, contained in gelatinous sheath, or very short without sheath and adhesive disc, or modified into ventral suckerlike attachment organ without sheath. Lateral antennae near base of coronal funnel. Corona a large buccal funnel

 273

without cilia, with or without lobes and/or marginal tentacles; opening of funnel more or less apical or oblique ventral. Eyespots absent or only obvious in juveniles. Ciliated corona, used for swimming, present in juveniles. Trophi uncinate. Oviparous and ovoviviparous. Three genera: Acyclus Leidy, 1882 (2 spp.), Atrochus Wierzejski 1893 (1 sp.), Cupelopagis Forbes, 1882 (1 sp.). Periphytic, freshwater. Predatory; Acyclus inquietus lives in colonies of Sinantherina socialis, feeding on eggs and larvae. Family Collothecidae Harring, 1913 (Figs. 4.52 D and 4.57 U) Illoricate. Sessile, rarely free-swimming; mostly with mucous or gelatinous case or tube. Body plan various, ovate, spherical, elongate conical, etc.; foot unsegmented, long without toes, tip mostly undifferentiated, or with small adhesive disc. Lateral antennae absent. Dorsal antenna reduced. Corona Collotheca type, funnel-shaped with vestibulum and infundibulum; a single ring of mobile, often strongly reduced cilia; margin of coronal funnel smooth, or with lobes, tentacles, or knobs bearing long motionless setae. Eyespots present, 2, rarely 3, often disappearing in adults, or absent. A ciliated corona, used for swimming, present in juveniles. Trophi uncinate. Oviparous and ovoviviparous. The family comprises 2 genera: Collotheca Harring, 1913 (45 spp.), and Stephanoceros Ehrenberg, 1832 (2 spp.). Predominantly benthic-periphytic, a few planktonic species. Freshwater, less frequent in brackish water.

4.7 Biogeography Due to the presence of resting stages (Hairston & Kearns 2002), all Rotifera, both Bdelloidea and Monogononta, have, in principle, the potential for cosmopolitan distribution (Artois et al. 2011). Nevertheless, data on their distribution is really scarce; thus, no reliable inference can be reliable for Bdelloidea (Ricci 1987, Ricci & Fontaneto 2009), whereas for Monogononta our biogeographical knowledge reflects the distribution of rotifer scientists (Fig. 4.58) more than that of rotifers themselves (Dumont 1983, Segers & De Smet 2008, Fontaneto et al. 2012a). The information for Seisonacea is even scantier (see chapter by Ahlrichs & Riemann). Notwithstanding such difficulties in analyzing biogeography of understudied animals with too few data on their distribution (Artois et  al. 2011), some Rotifera are known to have very limited distribution. For species that are locally common and that are easy to identify, such

274 

 4 Rotifera

Fig. 4.58: Global species richness of Monogononta at the finest resolution scale of the Biodiversity Information Standards (Taxonomic Database Working Group) records published between 1960 and 1992. The gray gradient of the scale bar indicates species richness. Black dots mark geographical units with no rotifer records; white circles mark geographical units with at least 100 species. (Modified from Fontaneto et al. 2012a.)

narrow distribution can be considered reliable (Segers & De Smet 2008). Hotspots of diversity and with high number of endemic species for Rotifera exist in Australia, China, North America, and tropical South America, whereas few endemics are present in Africa and in the Indian subcontinent (Segers & De Smet 2008). Environmental variables, linked to niche conservatism and geological history, are known to shape biogeographical patterns in larger organisms (Cox & Moore 2010), but no studies have successfully attempted yet to disentangle these variables at the global scale for Rotifera (Fontaneto et al. 2012a). At smaller scale, distribution and richness of Rotifera are known to be correlated to environmental variables such as altitude (Obertegger et al. 2010), temperature (Bērziņš & Pejler 1989), salinity (Kaya et al. 2010b), and trophic state (Obertegger & Manca 2011). Latitude is only a description of position, but it often correlates with ecological variables that are biologically relevant and shape the biogeographical patterns in species richness (Hawkins & Diniz-Filho 2004). As in other microscopic animals (Maraun et  al. 2007, Artois et al. 2011), it seems that no latitudinal gradient in species richness is present in Rotifera (Fontaneto et  al. 2012a). For Bdelloidea, polar areas such as Svalbard islands are at least as rich as temperate ones (Kaya et  al. 2010a); meanwhile, for Monogononta, although a global analysis revealed no latitudinal gradients (Fontaneto et al. 2012a), detailed studies focusing on specific taxa described latitudinal gradients in species richness, for example, in Brachionidae (Pejler 1977), in Lecanidae (Segers 1996), and in Trichocerca (Segers 2003). Also, the species composition of communities in Monogononta is known to change with latitude (Green 1972).

Detailed lists of rotifer species exist for several parts of the world, and the geographical distribution of most species is rather well known (recent reviews in Segers 2007, 2008). Nevertheless, much work still needs to be performed, new species are often found even in well-studied areas, and the geographical ranges of most species are often widened. Biogeographical studies using phylogenetic information (=phylogeography) have been performed only for few species, mostly of the Brachionus plicatilis species complex (e.g., Gómez et  al. 1995, 2000, 2002a, 2002b, 2007, Mills et  al. 2007, Campillo et  al. 2011). Recently, additional model taxa have been used, including the B. calyciflorus species complex (e.g., Xiang et  al. 2011a, b), but also Testudinella clypeata (Leasi et  al. 2013) and several Synchaeta (Obertegger et  al. 2012). The consensus of these analyses is that rotifers, as other microscopic organisms with high potential for passive dispersal, are indeed widely distributed, but they still experience the constraints of geography in their distribution (Fontaneto 2011). Thus, the occurrence of refugia can be analyzed as in larger organisms (Gómez & Lunt 2006), biogeographical patterns exist, with evidence of enclave distribution, founder events, and localized genetic differentiation (De Meester et al. 2002).

4.8 Paleontology As for all the Platyzoa clade, the fossil record is almost non-existent. The few proper fossil records of rotifers have been found in amber and are limnoterrestial Bdelloidea (Poinar & Ricci 1992, Waggoner & Poinar 1993). More

4.9 Ecology 

numerous are the records for subfossil rotifers from the Holocene in peat bogs and other kind of deposits, both of Bdelloidea (e.g., Warner & Chengalath 1988, 1991) and of Monogononta (e.g., Swadling et al. 2001, Turton & McAndrews 2006). Resting eggs of rotifers are also known to occur as subfossil in different sediments (Van Geel 2001). Interestingly, some records of unidentified palynomorphs may actually be rotifer resting eggs (Van Geel 1998). Subfossil resting eggs may hatch after up to a century in the sediments (Piscia et  al. 2012), as it happens for ephippia of cladoceran crustaceans (Caceres 1998). Thus, they do not represent true fossils, but a genetic legacy from the past that can still “invade” the present time.

4.9 Ecology The rotifer species are conditioned by the classical abiotic and biotic variables, such as temperature, pH, ions, organic compounds, prey-predator relationships, competition, food supply, and parasites (see Wallace et  al. 2006). Many species play an important role in the food webs because of their large population size and rapid turnover rate. Rotifers serve as food for, e.g., copepods, oligochaetes, raptorial cladocerans, chironomids, chaoborids, fish larvae, and planktivorous and benthivorous fish (e.g., Schmid-Araya & Schmid 1995, Monakov 2003). Rotifers are ubiquitous components of the aquatic biocoenoses, both freshwater and saline. The great majority (~85%) of the species known to date, both bdelloids and monogononts, occur in freshwater environments; the others are inhabitants of athalassic, inland saline waters, and the true thalassic, brackish, and marine, environments (e.g., Fontaneto et al. 2006a, 2008). Most species are free-living, while others are sessile, epibiotic, ectoparasitic or endoparasitic; freeliving or sessile taxa may be solitary or colonial. Bdelloids are predominantly semi-aquatic, whereas monogononts are mostly found in truly aquatic environments. Species diversity of monogononts is smallest in the polar regions, whereas bdelloids are rarer in the subtropics and tropics (Donner 1956).

4.9.1 Feeding ecology The food and way of feeding depend on the structure and function of the corona and trophi. As a group, bdelloid rotifers are primary consumers and microphagous, feeding on bacteria, yeasts, and small algae, by filtering,

 275

scraping, or browsing (Melone et al. 1998a). Many species are exclusively feeding on bacteria and yeasts (Habrotrocha thienemanni) and some feed on chlorophytes in particular (Philodinavus paradoxus, Habrotrocha tridens); Macrotrachela fungicola feeds on a mushroom, and a single species, Abrochtha carnivora, preys on other bdelloids and monogononts (Ricci et al. 2001). The feeding habits of monogononts are enormously varied, as reflected by the variation in trophi and corona types. Both primary and secondary consumers, microphagous and macrophagous taxa, as well as omnivorous, herbivorous, and carnivorous species occur (see, e.g., Pourriot 1977, Monakov 2003). Monogonont rotifers with a well developed corona may be microphagous or macrophagous, consuming detritus, tripton, bacteria, yeasts, protozoans, and algae. They are usually planktonic, collecting food by currents generated through the coronal cilia, and grinded by the unci of the malleate or malleoramate trophi (e.g., Brachionidae, Conochilidae, Trochosphaeridae). Free-living herbivorous benthic-periphytic and semi-planktonic taxa mostly have a less developed corona and virgate trophi specialized for piercing and pumping. For example, Ascomorpha ovalis (Gastropodidae) feeds preferentially on dinoflagellates (Ceramium, Peridinium), by piercing the cells and sucking their content; it has special coronal palps to grasp the cell and to hold it in place when feeding (Stelzer 1998). Another species of the latter family, Gastropus hyptopus, grasps a cell of a Synura colony (xanthophyte) and swallows it whole. Members of the Notommatidae and Trichocercidae (e.g., Notommata copeus, Trichocerca rattus) move along the algal filaments of Spirogyra and Mougeotia, and feed by piercing the cells and sucking their content. Creeping species with in general ventrally situated mouth opening browse the epiphytic algae and bacteria (Proalidae, Lecanidae, Lepadellidae). In the sessile Collotheca spp. and Stephanoceros fimbriatus, the anterior part is transformed into a broad funnel or vestibulum, with arms that draw together to trap the prey (flagellates, ciliates) when it enters the infundibulum. Food selectivity is evident for several other taxa, e.g., Lindia spp., and Brachionus diversicornis preferentially feeds on cyanobacteria; Notommata collaris typically feeds on a single desmid genus (Closterium), whereas N. pachyura shows a preference for different genera (Closterium, Penium, Staurastrum), and Trichocerca elongata is feeding on Oedogoniales. Carnivory is quite common in Asplanchnidae, Dicranophoridae, Notommatidae, Synchaetidae, and Atrochidae. Among the prey eaten are protozoans, rotifers, nematodes, small crustaceans, and their juveniles; cannibalism has

276 

 4 Rotifera

been reported in Asplanchna (Gilbert 1976a, b, 1980). The prey may be grasped by the forcipate trophi (e.g., Dicranophorus) or swallowed after contact with the corona and help of the incudate trophi (Asplanchna) or its contents are sucked out using virgate trophi (Synchaeta). The sessile Cupelopagis vorax (Atrochidae) has a very large anterior funnel devoid of coronal cilia that envelops the prey, which is lacerated by the uncinate trophi. Selectivity for prey is also evident for the raptorial species. For example, Notommata glyphura, N. aurita, and Ploesoma hudsoni feed on other rotifers; N. pseudocerberus feeds exclusively on the ciliates Stentor niger and S. polymorphus, but avoids S. coerulaeus; Trichocerca capucina sucks out eggs of other planktonic rotifers; Dicranophorus isothes penetrates between the valves of cladocerans and eats the cladoceran from within. Although prey selectivity is evident in many species, populations of the same species may differ in their preferences (Gilbert 1980). Proales fallaciosa and Dicranophorus isothes are known as scavengers, feeding on dead oligochaetes, microcrustaceans, and macroinvertebrates. For the hosts and feeding habits of parasitic rotifer species, see Section 4.9.2.3, Symbiotic Associations.

4.9.2 Habitat 4.9.2.1 Freshwater and limnoterrestrial habitats Lentic free water. The euplanktonic rotifers of lentic waters are independent of substrate, and able to dwell at the surface or at determined depths. The rotifer fauna of the pelagial of lakes and large water bodies almost exclusively consists of euplanktonic species, while plankton of ponds and shallow water bodies is composed of euplanktonic and semi-planktonic species characteristic of the littoral and submerged vegetation. Many benthic species may be found swimming around in shallow waters as well. The rotifer species richness of the pelagic zone is lower than for the littoral region and psammon habitat. For instance, Muirhead et  al. (2006) estimated species richness for 3 temperate freshwater lakes in Poland, and found total species numbers ranging from 167 to 205 species, with 44–65 for the pelagic, 137–162 for the littoral, and 100–135 for the psammon habitat. Rotifer densities may vary from a few individuals per liter in oligotrophic waters to >10,000 ind L–1 in nutrient-enriched waters. Among the freshwater bdelloids, only a single species, Rotaria neptunia, is truly planktonic in eutrophied waters. Several others are often semi-planktonic in the littoral region or in shallow waters (e.g., Rotaria macrura, Philodina citrina, P. megalotrocha). All other euplanktonic

rotifers belong to the monogononts, in particular to the Asplanchnidae, Brachionidae, Conochilidae, Synchaetidae, and Trochosphaeridae. The rotifer communities are influenced by physical, chemical, and biological factors, whose relative role in structuring assemblages and controlling seasonal dynamics may vary within or between aquatic systems (Hunter & Price 1992). To mention are temperature, oxygen concentration, light intensity, and pH (e.g., Hofmann 1977), food quality and quantity (Dumont 1977), exploitative and interference competition (May & Jones 1989, MacIsaac & Gilbert 1991, Fussmann 1996), predation (Williamson 1983, Neill 1984), and parasitism (Ruttner-Kolisko 1977). The rotifer fauna can be divided into perennial, and seasonal taxa, with or without distinct maxima, and species with erratic occurrence. The seasonal succession and maxima of the different species is in general fairly constant, and characteristic of the water body, although variations related to the climatological conditions may occur. Seasonal succession in temperate regions is primarily driven by temperature, and related food supply (e.g., seasonal development of phytoplankton); the major determinant involved in the tropics is apparently the alternation of wet and dry seasons (e.g., Apstein 1904, Green 1960). Although most rotifers have a wide tolerance range for temperature (Bērziņš & Pejler 1989), many species show a distinct seasonality in occurrence (May 1983). Three main categories in response to water temperature can be distinguished: eurythermous species able to maintain a dense population over a wide range of temperatures (e.g., Keratella cochlearis, K. quadrata), and stenothermous species unable to maintain populations outside a well-defined range, i.e., warm stenothermous species preferring warm water (e.g., Anuraeopsis fissa, Pompholyx sulcata, Trichocerca pusilla), and cold stenothermous taxa exclusively found in winter and in cold hypolimnion (e.g., Kellicottia spp., Keratella hiemalis, Notholca spp.). Some species are eurytherm (Synchaeta kitina), but cold or warm adapted, showing high population development at low or high temperatures, respectively (May 1983). Light, food, temperature, oxygen, and predation are among the influences responsible, directly or indirectly, for the vertical distribution of planktonic rotifers. In stratified lakes, the vertical distribution of rotifers is strongly related to the season, and the thermal and oxygen gradients, with some species also showing diurnal vertical migrations (e.g., Larsson 1971, Ruttner-Kolisko 1980b, Armengol-Díaz et  al. 1993, Miracle & Armengol-Díaz 1995). Rotifer populations concentrate at the depths with pronounced gradients. For example, in Lake La Cruz, Armengol-Díaz et al. (1993) found that when stratification

4.9 Ecology 

develops from winter to summer, some rotifers (Anuraeopsis fissa, Keratella quadrata, Polyarthra dolichoptera) show a downward migration following the thermocline to produce extremely dense populations near the oxicanoxic boundary, whereas others (Anuraeopsis miraclei, Filinia hofmanni) migrated upward following the oxycline. The vertical segregation of Cephalodella acidophila and Elosa woralii, dominant in the hypolimnion and the epilimnion of acid mining lakes, respectively, is attributed to specific differences in efficiency for using the autotrophic or mixotrophic form of Chlamydomonas acidophila as food. Elosa shows a higher efficiency in using the autotrophic form, whereas Cephalodella feeds on the mixotrophic form of the flagellate occurring in the deeper water under low light. The mixotrophic form proved a poor food source for Elosa (Weithoff & Wacker 2007, Hartwich et al. 2010). Active diurnal vertical migrations of rotifers have been described for several species (see, e.g., Miracle 1977, Magnien & Gilbert 1983). Migration tends to be different for different species and time of the year, and the same species may show upward nocturnal migration at one season, and the reverse at another, or no migration at all (e.g., George & Fernando 1970, Miracle 1977). The amplitude of diurnal migration ranges from 0.2 to 8 m. Variables triggering migration are temperature, oxygen concentration, competition for food resources, avoidance of predation, mechanical interference competition, etc. For example, George & Fernando (1970) found that light was controlling the migration and vertical distribution of Filinia terminalis, Keratella quadrata, and Polyarthra vulgaris. Food resource, both its quality and quantity, apparently determined the diurnal migrations to the surface at night of Synchaeta pectinata (8 m) and Trichocerca simoneae (6 m) in a Polish dystrophic lake (Karabin & Ejsmont-Karabin 2005). In a study on rotifer water layer preferences (0–2 m, and 5–35 m) in an oligotrophic mountain lake at midday and midnight during summer, Obertegger et  al. (2008) found the vertical distribution related to temperature, food availability, presence of predators, and exposure to UV radiation. Some species (Keratella quadrata, Synchaeta pectinata) showed a population maximum in the deeper layer during midday, and in the upper layer during midnight, whereas others always remained in the lower layer (Asplanchna priodonta, Filinia terminalis, Synchaeta kitina) or upper layer (Polyarthra dolichoptera, S. grandis). Migrating species apparently favored the higher temperatures in the upper layer, while non-migrating ones seemed restricted by, e.g., food supply. Positioning in the deeper layer during midday seemed a strategy for UV avoidance; moreover, the possession of photoprotective compounds probably played a role in UV tolerance as well. In a shallow (1.5 m) pond at

 277

Vermont, USA, Gilbert & Hampton (2001) found a possible predator avoidance-response cascade induced by notonectids. A reverse diurnal migration was noted for a single rotifer species, Polyarthra remata, but not for 6 other cooccurring rotifers, and the copepod Tropocyclops extensus, which is an important predator of Polyarthra. During the day when Tropocyclops is most abundant near the bottom, Polyarthra is most abundant near the surface. The diurnal migration of Tropocyclops itself to the deeper layers during the day was supposed to be an avoidance response to the notonectid Bueno, preying on the copepod during the day. Examining diurnal cycles of reproduction and vertical migration of Keratella crassa, Magnien & Gilbert (1983) found a differential migration of ovigerous and non-ovigerous females, the ovigerous ones reaching their lowest and highest positions in the water column about 4 and 6 h later than non-ovigerous females. Lotic free water. Swift flowing mountain waters cannot maintain downstream-directed rotifer plankton populations: species found in such habitat originate from vegetation, benthos, and hyporheos. In rivers and streams, however, rotifers usually form the dominant component of the potamoplankton (e.g., Shiel et al. 1982, Walz 1995, Zimmermann-Timm et al. 2007, Bertani et al. 2011). For example, the number of species amounts to 61 for the Middle Loire, France (Lair 2005), to 27, 53, and 74, respectively, for 3 Masurian streams (Ejsmont-Karabin & Kruk 1998), and up to 51 taxa for the freshwater reach of the river Scheldt, Belgium (Azémar et al. 2010). Planktonic loricate species usually predominate, followed by epibenthic and littoral species, and last by illoricate rotifers, which are the least diverse and abundant (Lair 2005, Azémar et al. 2010). However, land-use activity can influence this sequence, with the littoral and benthic species becoming dominant in streams draining agriculturally developed catchments (Ejsmont-Karabin & Kruk 1998). The loricate planktonic species appear better adapted to the current than the soft-bodied ones. Densities can be >10,000 ind L–1 (e.g., Zimmermann-Timm et  al. 2007). Brachionidae belonging to the genera Keratella and Brachionus usually dominate numerically. This can be explained by their ability of rapid reproduction and the capacity of several species to continue growing at currents of 0.2 m sec-1 (Lair 2005). The main factors controlling rotifer abundance are temperature and water transport time. Macrophytes. Aquatic macrophytes create an enormous increase in habitat complexity, food availability, and shelter for the fauna. As such, rotifers are known to show a greater diversity and mostly higher densities in stands of aquatic vegetation than in the adjacent pelagic zone (e.g., Pennak 1966, Duggan 2001, Duggan et  al.

278 

 4 Rotifera

2001, Kuczyńska-Kippen et al. 2003, Kuczyńska-Kippen & Cerbin 2003, Kuczyńska-Kippen 2005). The rotifer assemblages are dominated by Lecanidae, Lepadellidae, Notomatidae, and Trichocercidae. The periphytic community comprises (1) plant browsers rarely leaving the substrate (e.g., Colurella colurus), (2) browsing species that leave the substrate frequently, and swim around in the spaces between the vegetation (e.g., Lecane lunaris), (3) sessile species, (4) planktonic species that browse occasionally (e.g., Brachionus calyciflorus), and (5) chiefly planktonic taxa (e.g., Filinia, Hexarthra, Kellicottia). Several species, especially sessiles, favor particular or few plant species and/or specific zones on the plants (e.g., Edmondson 1944, Wallace 1977a, b, Wallace & Edmondson 1986). For example, Stephanoceros fimbriatus is predominantly found on Utricularia, Cupelopagis vorax favors submersed plants with broad flat or very gently convex leaves (Potamogeton, Ranunculus), and Collotheca gracilipes selects leaflets of Elodea canadensis. Species richness and abundance of periphytic rotifers are generally related to macrophyte architecture, and greater among plants with a greater structural complexity providing a greater variety of habitats, and a larger quantity or variety of food (e.g., Duggan et  al. 2001, Kuczyńska-Kippen & Nagengast 2006, Lucena-Moya & Duggan 2011). An increasing structural complexity apparently does not affect species richness and abundance of the planktonic taxa. A greater architectural complexity likewise stands for more opportunities for refuge from predators. Kuczyńska-Kippen (2007) found several planktonic rotifers to seek anti-predator refuge in the macrophyte stands during the daytime. Walsh (1995) assessed habitatspecific predation susceptibilities for Euchlanis dilatata in the presence of Myriophyllum exalbescens, Elodea canadensis, and Ceratophyllum demersum and 2 predators (damselfly nymphs and Hydra). Rotifer survival was greatest on Myriophyllum in the presence of both predators, and conversely, the presence of the Elodea and Ceratophyllum increased rotifer susceptibility to predation by the damselfly nymphs, by increasing their foraging ability. Decreasing the macrophyte complexity by, for example, removing leaves, resulted in a lower survival of Euchlanis. In her study on rotifer body size and macrophyte architecture, Kuczyńska-Kippen (2005) found that the size distribution of the rotifer assemblages was directly related to the morphological and spatial structural complexity of the substrates. Densities of rotifers were also higher in the more heterogeneous habitats, probably by an increase of the potential refugia. Benthos. Rotifers living on and in the sediment of bottoms of both lentic and lotic habitats have hardly

been studied, because of difficulties with isolation of the specimens from the substrate (e.g., Carlin 1939, Pejler 1962, Donner 1970, Hoebel 1978, Anderson & De Henau 1980, Nalepa & Quigley 1983, Ricci & Balsamo 2000). The rotifer fauna is apparently less diverse and dominated by Dicranophoridae and Notommatidae (Cephalodella spp.), with some species, e.g., Atrochus tentaculatus, Mytilina crassipes, Paradicranophorus hudsoni characteristic of the habitat. Most species dwell at the sediment surface or mainly in the upper 1 cm of substrate. Densities are variable and low in oxygen poor environments; under favorable conditions a high density of 77,300 ind m–2 was found by Nalepa & Quigley (1983) in nearshore Lake Michigan. Arenal. The interstitial of sandy sediments on the shores and the bottom of lentic and lotic waters is inhabited by a psammon community, of which rotifers may form an important component. For example, Wiszniewski (1934a) recorded 82 species in Lake Wigry (Poland), Myers (1936) found 145 species in Lenape and Union Lakes (Virginia, USA), Turner & Palmer (1996) found 77 taxa in Goose Creek (USA), and Muirhead et al. (2006) recorded 119 species in Lake Mikolajskie (Poland). The rotifers can be divided into psammobiotic or almost exclusively found in sandy sediment (e.g., Lecane psammophila, Myersinella spp., Trichocerca taurocephala), psammophilic or living in sand preferentially, but also in other habitats (e.g., Lecane closterocerca, L. lunaris, Colurella colurus), and psammoxenic or accidental components of sand (e.g., Wiszniewski 1937, 1947). Three zones can be distinguished in the psammolittoral region with reference to their relative degree of saturation with water, and each inhabited by its own community of organisms (e.g., Wiszniewski 1934b, 1947). The hydropsammon is found in the hydroarenal or permanently submerged sand, the hygropsammon occurs in the hygroarenal or sand above and adjacent to the water level and saturated by wave action and capillarity and the eupsammon living in the moist, partially saturated euarenal situated above the hygroarenal. The hydropsammon experiences less stress in comparison to the other zones and usually shows a lower species richness and lower density of rotifers (Wiszniewski 1947, Pennak 1940,, Bielañska-Granjer 2004, Kalinowska et  al. 2012), whereas the hygropsammon in general yields the highest number and density of rotifers (e.g., Wiszniewski 1947, Evans 1982, BielañskaGranjer 2001, Bielañska-Granjer & Molanda 2008). Total rotifer density and density of the different species in general is very variable, differing over time, among sites and between depths in the sand (e.g., Evans 1982, Bielañska-Granjer 2001, Ejsmont-Karabin 2005); reported total densities range from 0.0 to >1,200 ind cm–3 or amount

4.9 Ecology 

to >1,500 ind 100 cm–2. The structure and density of the psammon rotifer assemblages apparently depend primarily on sand grain size (Ejsmont-Karabin 2004, BielañskaGranjer & Molanda 2008). In general, psammobiotic monogononts prefer the sand grain size fraction 0.5–1.0 mm, a fraction avoided by psammophiles and psammoxenes; bdelloids only prefer the smallest (