Handbook of Zoology: Phylum Bryozoa 9783110586312, 9783110585407

Table of contents :
Preface
Contents
List of contributing authors
1 General introduction
2 Fossil record and evolution of Bryozoa
3 Morphology of bryozoans
4 Sexual reproduction in Bryozoa
5 Larval structure and metamorphosis
6 Behavior
7 Phylactolaemata
8 Cyclostomata (Stenolaemata)
9 Gymnolaemata
10 Ctenostomata
11 Gymnolaemata, Cheilostomata
Appendix
Index

Citation preview

Handbook of Zoology Phylum Bryozoa

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

Phylum Bryozoa Edited by Thomas Schwaha

DE GRUYTER

Phylum Bryozoa

Edited by Thomas Schwaha

DE GRUYTER

Scientific Editor PD Dr. Thomas Schwaha Universität Wien Department of Evolutionary Biology, Integrative Zoology Althanstraße (UZA I) 14 1090 Wien Austria [email protected]

ISBN 978-3-11-058540-7 e-ISBN (PDF) 978-3-11-058631-2 e-ISBN (EPUB) 978-3-11-058568-1 ISSN 2193-4231 Library of Congress Control Number: 2020941128 Bibliographic information published by the Deutsche Nationalbibliothek The Deutsche Nationalbibliothek lists this publication in the Deutsche Nationalbibliografie; detailed bibliographic data are available on the Internet at http://dnb.dnb.de. © 2021 Walter de Gruyter GmbH, Berlin/Boston Typesetting: Compuscript Ltd. Shannon, Ireland Printing and Binding: CPI books GmbH, Leck www.degruyter.com

Preface Despite almost 200 years of research on bryozoans, bryozoology, these animals remain comparatively little studied and many different aspects of their biology are still poorly understood. The mere numbers of several thousand Recent species and even more fossil ones just indicate how diverse bryozoans are, but only still little how these evolved in time and space. Most researchers are easily intrigued by this phylum of colonial animals and easily realize how little we still know about this fascinating group. In 2017 I was asked to edit the volume ‘Bryozoa’ for the Handbook of Zoology series, which I gladly accepted as the series is of high quality and can be used as good reference work for several phyla already. Such an undertaking is, of course, always a challenge. Based on my own background, it focuses on Recent bryozoans and only has one chapter dedicated to fossils. Recently, Bryozoan Paleobiology by Paul Taylor was just published this year and offers an excellent summary on this particular topic.  

https://doi.org/10.1515/9783110586312-202

This book tries to provide an overview of several general aspects of bryozoan biology and a more detailed insight into the various diverse groups. It brings together several authors of various disciplines. I hope my efforts in the compilation of this book will yield students, more experienced researchers and specialists a useful summary. I’m deeply grateful to Andreas Schmidt-Rhaesa for offering me to edit this volume and also all the authors involved in this long and intense project. Numerous colleagues also provided images or samples used for some of the chapters: Sebastian Decker (Vienna), Paul Taylor (London), Priska Schäfer (Kiel), Piotr Kuklinski (Sopot), Peter Batson (Dunedin), Joachim Scholz (Frankfurt), Matthew Dick (Sapporo), Phil Bock (Melbourne). I’m also indebted to Mary Spencer Jones for letting me dig through the Natural History Museum London collection, which provided several specimens used for some of the chapters. Thomas Schwaha August, 2020

Contents Preface

v

List of contributing authors

xv

Thomas Schwaha 1 General introduction 1 1.1 A brief introduction to bryozoans 1 Past summaries on bryozoans 1.2 2 Topics in bryozoan research 1.3 5 Terminological issues 1.4 5 1.5 Phylogeny and systematics of bryozoans 6 1.5.1 Relationship of bryozoans to other phyla 6 The morphological perspective 1.5.1.1 6 The molecular perspective 1.5.1.2 7 1.5.2 Internal systematics and phylogeny of bryozoans 7 Outline of this book 1.6 8 Literature 8 Andrej Ernst 2 Fossil record and evolution of Bryozoa 11 2.1 Introduction 11 2.2 Methods for studying fossil bryozoans 12 2.3 Development and evolution of bryozoans in the Palaeozoic 12 Morphology of the Palaeostomata 2.3.1 14 Definition of Palaeostomata 2.3.1.1 14 Morphology of Palaeostomata 2.3.1.2 14 Overview of the orders of the 2.3.1.3 Palaeostomata 21 2.4 Evolutionary history of Bryozoa 23 2.4.1 Ordovician 23 2.4.2 Silurian 27 2.4.3 Devonian 27 2.4.4 Carboniferous 29 2.4.5 Permian 31 2.4.6 Triassic 33 2.4.7 Jurassic 35 2.4.8 Cretaceous 35 2.4.9 Crisis at the K/T boundary 37 2.4.10 Cenozoic 37 2.5 Bryozoans in reefs and other organic build-ups 38

2.6 Evolutionary patterns in bryozoans 39 2.6.1 Macroevolutionary trends in bryozoans 40 Growth and growth forms 2.6.1.1 40 2.6.1.2 Feeding 41 2.6.1.3 Reproduction 41 2.6.1.4 Defense 41 Homeomorphy and convergence 2.6.2 42 Literature 43 Thomas Schwaha 3 Morphology of bryozoans 57 3.1 A short glimpse into morphological and anatomical research of bryozoans 57 3.2 Outline of this chapter 57 3.3 General body organization and features 57 3.4 Body cavity 59 3.5 Coelomic partitions or canals 60 3.5.1 Phylactolaemata 61 3.5.2 Myolaemata 62 3.6 Body walls 62 3.6.1 Phylactolaemata 62 3.6.2 Cyclostomata 63 3.6.3 Gymnolaemata 65 3.7 Communication areas and pores 65 3.8 Lophophore 67 3.8.1 General structure 67 3.8.2 Tentacles and ciliation 67 3.9 Digestive system 69 3.9.1 General features of the digestive tract 69 3.9.2 Cytological features of the digestive epithelium 71 3.10 Funicular system 72 Nervous system 3.11 77 Tentacle innervation 3.11.1 79 3.11.1.1 Phylactolaemata 79 3.11.1.2 Myolaemata 83 Peripheral innervation 3.11.2 83 3.11.2.1 Tentacle sheath, apertural, and body wall innervation 83 Visceral innervation 3.11.2.2 85 Muscular system 3.12 86 3.12.1 Muscles associated with the body wall 86 3.12.2 Apertural muscles of bryozoans 86

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3.12.2.1

Functional aspects of the apertural muscles 90 Lophophoral muscles 3.12.3 90 Tentacle muscles 3.12.3.1 90 Lophophoral base muscles 3.12.3.2 91 Tentacle sheath muscles 3.12.4 93 Digestive tract muscles 3.12.5 93 Retractor muscles 3.12.6 93 Excretory system 3.13 95 3.14 Coelomocytes 95 Literature 96 Andrew N. Ostrovsky 4 Sexual reproduction in Bryozoa 101 4.1 Generalities of bryozoan reproduction 101 4.2 Reproductive patterns 101 4.3 Colony sexual structure and sexual polymorphism 102 Gonado- and gametogenesis 4.4 102 4.5 Gamete release and fertilization 109 Embryonic incubation 4.6 110 4.7 Matrotrophy and evolution of reproductive patterns 116 Life histories 4.8 118 Acknowledgments 118 Literature 119 Alexander Gruhl 5 Larval structure and metamorphosis 123 5.1 Introduction 123 Embryonic development 5.2 123 Larval morphology 5.3 128 5.3.1 General 128 Nervous and sensory system 5.3.2 129 5.3.3 Musculature 131 5.3.4 Digestive tracts and particle feeding 131 5.3.5 Differences between larval types 133 5.4 Larval behavior, physiology, and ecology 135 5.5 Metamorphosis 136 Literature 138 Judith E. Winston & Alvaro E. Migotto 6 Behavior 143 6.1 Introduction 143 6.2 Feeding 143 6.2.1 Food 143 6.2.2 Mechanics of feeding 146

Flow and feeding 148 Morphology of feeding 150 Lophophore size 150 Individual behavior 150 Tentacle sheath function 151 Mouth and gut morphology 153 Use of tentacles in feeding 153 Protrusion, expansion, and retraction 153 Rejection of particles (individual and colony) 154 Particle handling by cilia and 6.2.5.6 tentacles 156 Individualized behavior patterns 6.2.6 157 Group and colonial feeding patterns 6.2.7 157 Colony integration 6.2.7.1 163 Reproductive behavior 6.3 163 6.3.1 Male zooids (androzooids) 165 6.4 Behavior of avicularia and vibracula 165 B zooids 6.4.1 168 Sessile avicularia 6.4.2 168 Bird’s head avicularia 6.4.3 168 Avicularian behavior 6.4.4 169 Vibraculan behavior 6.4.5 170 6.4.6 Nanozooids 172 Motility of zooids and colonies 6.5 172 6.6 Competition 175 6.6.1 Overgrowth 175 Feeding interference 6.6.2 177 Fighting buds 6.6.3 177 6.6.4 Molting 177 Chemical defenses 6.6.5 177 6.7 Interactions with other organisms 178 6.7.1 Fouling 180 6.7.2 Commensalism 180 6.7.3 Mutualism 180 6.7.4 Facilitation 180 6.8 Discussion 180 Acknowledgments 181 Literature 181

6.2.3 6.2.4 6.2.4.1 6.2.5 6.2.5.1 6.2.5.2 6.2.5.3 6.2.5.4 6.2.5.5

Thomas Schwaha 7 Phylactolaemata 189 7.1 Short history of research 189 7.1.1 The Japanese freshwater bryozoan research 189 7.2 Life of freshwater bryozoans 189 7.2.1 Distribution and dispersal 189 7.2.2 Current topics in phylactolaemate research 190 7.2.3 General colony morphology 191

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Morphological characters of zooids 7.2.4 192 7.2.4.1 Horseshoe-shaped lophophore and associated structures 192 7.2.4.2 Epistome 192 Body-wall musculature 7.2.4.3 192 Vestibular pore 7.2.4.4 192 7.2.5 Statoblasts 195 Forms of statoblasts 7.2.5.1 195 Systematic account 7.3 205 Cristatellidae Allman, 1856 7.3.1 205 General colony morphology 7.3.1.1 205 Zooidal characteristics 7.3.1.2 205 7.3.1.3 Statoblasts 207 Fredericellidae Allman, 1856 7.3.2 207 General colony morphology 7.3.2.1 207 Zooidal characteristics 7.3.2.2 207 7.3.2.3 Statoblasts 207 7.3.3 Lophopodidae Rogick, 1935 207 General colony morphology 7.3.3.1 211 Zooidal characteristics 7.3.3.2 211 7.3.3.3 Statoblasts 211 Pectinatellidae Lacourt, 1968 7.3.4 211 General colony morphology 7.3.4.1 211 Zooidal characteristics 7.3.4.2 213 7.3.4.3 Statoblasts 213 7.3.5 Plumatellidae Allman, 1856 213 General colony morphology 7.3.5.1 213 Zooidal characteristics 7.3.5.2 213 7.3.5.3 Statoblasts 214 7.3.6 Stephanellidae Lacourt, 1968 216 General colony morphology 7.3.6.1 216 Zooidal characteristics 7.3.6.2 216 7.3.6.3 Statoblasts 216 7.3.7 Tapajosellidae Wood & Okamura, 2017 216 7.4 Evolution and phylogeny of phylactolaemate bryozoans 216 7.4.1 Statoblast evolution 218 7.4.2 Ectocyst and colonial evolution 218 Literature 218 Andrej Ernst 8 Cyclostomata (Stenolaemata) 225 8.1 Introduction 225 8.2 Morphology of Cyclostomata 225 8.2.1 Soft part morphology 226 8.2.1.1 Membranous sac and tentacle protrusion 226 8.2.1.2 Skeletal organization (free-walled, fixed-walled) 227

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230 Feeding morphology Morphology of mineralized skeleton 231 8.2.2.1 Biomineralization and skeletal ultrastructure of cyclostomes 231 Early stage of colony development 8.2.2.2 231 Internal zooecial structures 8.2.2.3 232 Extrazooidal structures 8.2.2.4 233 Polymorphism in cyclostomes 8.2.3 233 8.2.4 Sexual reproduction and brood chamber morphology 234 8.2.4.1 Polyembryony 234 Brood chambers of Cyclostomata 8.2.4.2 234 Colonial forms of Cyclostomata 8.2.5 236 8.3 Evolutionary trends in Cyclostomata 238 Systematics of Cyclostomata 8.4 240 8.4.1 Suborder Paleotubuliporina Brood, 1973 240 Family Corynotrypidae Dzik, 1981 241 Family Sagenellidae Brood, 1975 241 Family Crownoporidae Ross, 1967 241 Family Flabellotrypidae Dzik, 1992 241 Family Diploclemidae Gorjunova, 1992 241 Suborder Tubuliporina Milne-Edwards, 8.4.2 1838 241 Family Stomatoporidae Pergens & Meunier, 1886 243 Family Oncousoeciidae Canu, 1918 243 Family Tubuliporidae Johnston, 1837 243 Family Multisparsidae Bassler, 1935 243 Family Celluliporidae Buge & Voigt, 1972 243 Family Plagioeciidae Canu, 1918 244 Family Terviidae Canu & Bassler, 1920 244 Family Spiroporidae Voigt, 1968 244 Family Annectocymidae Hayward & Ryland, 1985 244 Family Diaperoeciidae Canu, 1918 244 Family Entalophoridae Reuss, 1869 245 Family Pustuloporidae Smitt, 1872 245 Family Frondiporidae Busk, 1875 245 Family Theonoidae Busk, 1859 245 Family Actinoporidae Vigneaux, 1949 245 Family Siphoniotyphlidae Voigt, 1967 246 Family Hastingsiidae Borg, 1944 246 Family Semiceidae Buge, 1952 246

8.2.1.3 8.2.2

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Family Cinctiporidae Boardman, Taylor & McKinney, 1992 246 Family Eleidae d’Orbigny, 1852 246 Suborder Articulata Busk, 1859 8.4.3 249 Family Crisiidae Johnston, 1838 249 Family Crisuliporidae Buge, 1979 249 Suborder Cerioporina von Hagenow, 8.4.4 1851 249 Family Cerioporidae Reuss, 1866 249 Family Leiosoeciidae Canu & Bassler, 1920 251 Family Densiporidae Borg, 1944 252 Family Clausidae d’Orbigny, 1854 252 Family Cavidae d’Orbigny, 1854 252 Family Sulcocavidae Viskova, 1972b 252 Family Corymboporidae Smitt, 1866 252 Family Pseudocerioporidae Brood, 1972 252 Suborder Cancellata Gregory, 1896 8.4.5 254 Family Horneridae Smitt, 1867 254 Family Stigmatoechidae Brood, 1972 254 Family Petaloporidae Gregory, 1899 254 Family Crisinidae d’Orbigny, 1853 254 Family Crassodiscoporidae Brood, 1972 254 Family Cytididae d’Orbigny, 1854 254 Family Canaliporidae Brood, 1972 255 Suborder Rectangulata Waters, 1887 8.4.6 255 Family Lichenoporidae Smitt, 1867 255 Family Alyonushkidae Grischenko, Gordon & Melnik, 2018 255 Family Anyutidae Grischenko, Gordon & Melnik, 2018 255 Literature 257 Thomas Schwaha 9 Gymnolaemata 265 9.1 Gymnolaemate bryozoans 265 9.2 Characters of Gymnolaemata 265 9.2.1 Ontogenetic characters 265 9.2.2 Zooidal characters 265 9.2.3 Reproductive characters 265 9.3 Phylogeny of Gymnolaemata 266 Literature 268 Thomas Schwaha 10 Ctenostomata 269 10.1 Short history of research 269 10.2 General overview: what is a ctenostome? 269

Diversity of ctenostomes 10.3 270 10.4 Ctenostome colonies, zooids, and terminology 270 Ctenostome polymorphism 10.5 271 Dormant buds or hibernacula 10.6 273 Systematic account 10.7 274 Ctenostomata vs. Euctenostomata 10.7.1 274 10.7.2 Benedeniporoidea Jebram, 1973 274 Euctenostomata Jebram, 1973 10.7.3 275 Alcyonidioidea Johnston, 1847 10.7.4 275 Alcyonidiidae Hincks, 1880 10.7.4.1 275 Clavoporidae Soule, 1953 10.7.4.2 277 Flustrellidridae Bassler, 1953 10.7.4.3 277 Pherusellidae Soule, 1953 10.7.4.4 277 Pachyzoidae d’Hondt 1983 10.7.4.5 280 Arachnidioidea Hincks, 1880 10.7.5 280 10.7.5.1 Arachnidiidae Hincks, 1880 280 Immergentiidae Silén, 1946 10.7.5.2 280 10.7.5.3 Nolellidae Harmer, 1915 285 10.7.5.4 Aethozoidae, d’Hondt 1983 (emend Reverter-Gil et al. 2016) 285 Monobryozoidae, Remane 1936 10.7.5.5 285 Hislopioidea Jullien, 1885 10.7.6 289 Hislopiidae Jullien, 1885 10.7.6.1 289 10.7.7 Paludicelloidea Allman, 1856 289 10.7.8 Vesicularioidea Johnston, 1847 291 10.7.8.1 Buskiidae Hincks, 1880 292 Terebriporidae D’Orbigny, 1847 10.7.8.2 292 Spathiporidae Pohowsky, 1978 10.7.8.3 292 10.7.8.4 Vesiculariidae, Hincks 1880 292 Victorelloidea Hincks, 1880 10.7.9 296 10.7.9.1 Victorellidae Hincks, 1880 296 Sundanellidae Jebram, 1973 10.7.9.2 296 Pottsiellidae Braem, 1940 10.7.9.3 299 Walkerioidea Hincks, 1880 10.7.10 299 10.7.10.1 Aeverrilliidae Jebram, 1973 299 10.7.10.2 Bathyalozoidae d’Hondt 1976 300 10.7.10.3 Farrellidae d’Hondt, 1983 301 10.7.10.4 Harmeriellidae, d’Hondt 1983 301 10.7.10.5 Hypophorellidae Prenant and Bobin, 1956 301 10.7.10.6 Jebramellidae Vieira et al., 2014 301 10.7.10.7 Mimosellidae Hincks, 1877 301 10.7.10.8 Triticellidae G. O. Sars, 1874 301 10.7.10.9 Walkeriidae Hincks, 1880 305 10.7.11 Problematic taxa 305 10.7.11.1 Panolicella Jebram, 1985 305 10.7.11.2 Pierrella Wilson and Taylor, 2013 305 10.7.11.3 Penetrantiidae Silén, 1946 307

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H  aywardozoon, Platypolyzoon, and Anguinella 308 10.8 Phylogeny and evolution of ctenostome bryozoans 308 Past phylogeny and interpretations 10.8.1 308 10.8.2 Perspectives 309 10.8.3 Remarks on the origin and evolution of solitary forms 309 Literature 310 10.7.11.4

Silviu O. Martha, Leandro M. Vieira, Javier Souto-Derungs, Andrei V. Grischenko, Dennis P. Gordon, Andrew N. Ostrovsky Gymnolaemata, Cheilostomata 11 317 General morphology 11.1 317 General structure 11.1.1 317 Structure of frontal walls and mechanism of 11.1.2 polypide eversion 319 11.1.3 Polymorphism 323 11.1.4 A short summary of cheilostome phylogeny 324 11.2 Ecology of cheilostome bryozoans 324 11.2.1 Habitat and substratum selection 324 11.2.2 Diet of cheilostomes 325 11.2.3 Predation on Cheilostomata 325 11.3 Systematics of Cheilostomata 326 11.3.1 Suborder Membraniporina Ortmann, 1890 326 11.3.1.1 Superfamily Membraniporoidea Busk, 1852b 326 Family Membraniporidae Busk, 1852b 326 Family Electridae Stach, 1937 (1851) 327 Family Sinoflustridae Gordon, 2009a 328 11.3.2 Suborder Inovicellina Jullien, 1888 328 11.3.2.1 Superfamily Aeteoidea Smitt, 1868a 328 Family Aeteidae Smitt, 1868a 328 11.3.3 Suborder Scrupariina Silén, 1941 329 11.3.3.1 Superfamily Scruparioidea Gray, 1848 329 Family Scrupariidae Gray, 1848 329 Family Eucrateidae Johnston, 1847 330 Family Leiosalpingidae d’Hondt & Gordon, 1996 331 11.3.4 Suborder Tendrina Ostrovsky, 2013 331 11.3.4.1 Superfamily Tendroidea Vigneaux, 1949 332 Family Tendridae Vigneaux, 1949 332

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Suborder Thalamoporellina Ostrovsky, 2013 332 Superfamily Thalamoporelloidea Levinsen, 11.3.5.1 1902 332 Family Thalamoporellidae Levinsen, 1902 332 Family Steginoporellidae Hincks, 1884a 332 11.3.6 Suborder Belluloporina Ostrovsky, 2013 333 Superfamily Belluloporoidea Ostrovsky, 11.3.6.1 2013 333 Family Belluloporidae Ostrovsky, 2013 333 Suborder Flustrina Smitt, 1867 11.3.7 334 Superfamily Flustroidea Lamoroux, 11.3.7.1 1816 334 Family Flustridae Lamouroux, 1816 334 11.3.7.2 Superfamily Buguloidea Gray, 1848 335 Family Bugulidae Gray, 1848 335 Family Beaniidae Canu & Bassler, 1927 336 Family Candidae d’Orbigny, 1851 337 Family Epistomiidae Gregory, 1893 338 Family Euoplozoidae Harmer, 1926 338 Family Jubellidae Reverter-Gil & FernándezPulpeiro, 2001 338 Family Rhabdozoidae MacGillivray, 1887a 338 11.3.7.3 Superfamily Calloporoidea Norman, 1903a 339 Family Calloporidae Norman, 1903a 339 Family Antroporidae Vigneaux, 1949 340 Family Bryopastoridae d’Hondt & Gordon, 1999 340 Family Chaperiidae Jullien, 1888 341 Family Cupuladriidae Lagaaij, 1952 342 Family Cymuloporidae Winston & Vieira, 2013 343 Family Doryporellidae Grischenko, Taylor, & Mawatari, 2004 343 Family Ellisinidae Vigneaux, 1949 343 Family Farciminariidae Busk, 1852b 344 Family Foveolariidae Gordon & Winston, 2005 in Winston (2005) 345 Family Heliodomidae Vigneaux, 1949 345 Family Hiantoporidae Gregory, 1893 346 Family Mourellinidae Reverter-Gil, Souto, & Fernández-Pulpeiro, 2011 347 11.3.5

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11.3.7.4 11.3.7.5 11.3.7.6 11.3.7.7 11.3.7.8 11.3.7.9 11.3.7.10 11.3.7.11

Family Pyrisinellidae† Di Martino & Taylor, 2012 348 Family Quadricellariidae† Gordon, 1984 348 Superfamily Cellarioidea Lamouroux, 1816 348 Family Cellariidae Lamouroux, 1816 348 Family Membranicellariidae Levinsen, 1909 349 Superfamily Lunulitoidea† Lagaaij, 1952 350 Family Lunulariidae Levinsen, 1909 350 Family Otionellidae Bock & Cook, 1998 350 Family Selenariidae Busk, 1854 350 Superfamily Microporoidea Gray, 1848 351 Family Microporidae Gray, 1848 351 Family Alysidiidae Levinsen, 1909 352 Family Aspidostomatidae Jullien, 1888 352 Family Calescharidae Cook & Bock, 2001 352 Family Calpensiidae Canu & Bassler, 1923 353 Family Chlidoniidae Busk, 1884 353 Family Onychocellidae† Jullien, 1882b 353 Family Poricellariidae† Harmer, 1926 354 Family Setosellidae Levinsen, 1909 355 Superfamily Monoporelloidea Hincks, 1882b 355 Family Monoporellidae Hincks, 1882b 355 Family Macroporidae† Uttley, 1949 356 Superfamily Adeonoidea Busk, 1884 357 Family Adeonidae Busk, 1884 357 Family Inversiulidae Vigneaux, 1949 357 Superfamily Arachnopusioidea Jullien, 1888 357 Family Arachnopusiidae Jullien, 1888 357 Family Exechonellidae† Harmer, 1957 358 Superfamily Bifaxarioidea Busk, 1884 359 Family Bifaxariidae Busk, 1884 359 Family Mixtopeltidae Gordon, 1994a 360 Superfamily Catenicelloidea Busk, 1852b 360 Family Catenicellidae Busk, 1852b 360

11.3.7.12 11.3.7.13 11.3.7.14 11.3.7.15 11.3.7.16 11.3.7.17 11.3.7.18

Family Eurystomellidae Levinsen, 1909 360 Family Petalostegidae Gordon, 1984 360 Family Savignyellidae Levinsen, 1909 361 Superfamily Celleporoidea Johnston, 1838 362 Family Celleporidae Johnston, 1838 362 Family Colatooeciidae Winston, 2005 363 Family Hippoporidridae Vigneaux, 1949 363 Family Phidoloporidae† Gabb & Horn, 1862 363 Superfamily Chlidoniopsoidea Harmer, 1957 365 Family Chlidoniopsidae Harmer, 1957 365 Superfamily Conescharellinoidea Levinsen, 1909 365 Family Conescharellinidae Levinsen, 1909 365 Family Batoporidae† Neviani, 1901 366 Family Lekythoporidae Levinsen, 1909 366 Family Orbituliporidae† Canu & Bassler, 1923 366 Superfamily Cribrilinoidea Hincks, 1879 367 Family Cribrilinidae Hincks, 1879 367 Family Euthyroididae Levinsen, 1909 368 Family Polliciporidae Moyano G., 2000 368 Superfamily Didymoselloidea Brown, 1952 368 Family Didymosellidae Brown, 1952 368 Superfamily Euthyriselloidea Bassler, 1953 369 Family Euthyrisellidae Bassler, 1953 369 Family Clathrolunulidae Gordon & Sanner, 2020 369 Family Neoeuthyrididae Gordon & Sanner, 2020 369 Superfamily Hippothooidea Busk, 1859b 369 Family Hippothoidae Busk, 1859b 369 Family Chorizoporidae Vigneaux, 1949 370 Family Haplopomidae Gordon in De Blauwe, 2009 370

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Family Pasytheidae Davis, 1934 (1881) 371 Family Trypostegidae Gordon, Tilbrook, & Winston in Winston, 2005 372 Family Vitrimurellidae Winston, Vieira, & Woollacott, 2014 372 11.3.7.19 Superfamily Lepralielloidea Vigneaux, 1949 372 Family Lepraliellidae Vigneaux, 1949 372 Family Atlantisinidae Berning, Harmelin, & Bader, 2017 373 Family Bryocryptellidae Vigneaux, 1949 374 Family Dhondtiscidae Gordon, 1989a 375 Family Hincksiporidae Powell, 1968 375 Family Jaculinidae Zabala, 1986 375 Family Metrarabdotosidae† Vigneaux, 1949 376 Family Romancheinidae Jullien, 1888 376 Family Sclerodomidae Levinsen, 1909 377 Family Tessaradomidae Jullien in Jullien & Calvet, 1903 377 Family Umbonulidae Canu, 1904b 378 11.3.7.20 Superfamily Mamilloporoidea Canu & Bassler, 1927 380 Family Mamilloporidae Canu & Bassler, 1927 380 Family Ascosiidae Jullien, 1882a 381 Family Cleidochasmatidae Cheetham & Sandberg, 1964 381 Family Crepidacanthidae Levinsen, 1909 381 11.3.7.21 Superfamily Pseudolepralioidea Silén, 1941 382 Family Pseudolepraliidae Silén, 1941 382 11.3.7.22 Superfamily Schizoporelloidea Jullien, 1882a 383 Family Schizoporellidae Jullien, 1882a 383 Family Acoraniidae López‐Fé, 2006 384 Family Actisecidae Harmer, 1957 384 Family Buffonellidae Jullien, 1888 384 Family Calwelliidae MacGillivray, 1887 384 Family Cheiloporinidae Bassler, 1936 386

11.3.7.23 11.3.7.24

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Family Cryptosulidae Vigneaux, 1949 387 Family Cyclicoporidae Hincks, 1884b 387 Family Echinovadomidae Tilbrook, Hayward, & Gordon, 2001 387 Family Eminooeciidae Hayward & Thorpe, 1988c 387 Family Escharinidae Tilbrook, 2006 388 Family Fatkullinidae Grischenko, Gordon, & Morozov, 2018 389 Family Fenestrulinidae Jullien, 1888 390 Family Gigantoporidae Bassler, 1935 390 Family Hippaliosinidae Winston, 2005 390 Family Hippopodinidae Levinsen, 1909 391 Family Lacernidae Jullien, 1888 392 Family Marcusadoreidae Winston, Vieira, & Woollacott 2014 393 Family Margarettidae Harmer, 1957 394 Family Mawatariidae Gordon, 1990 395 Family Microporellidae Hincks, 1879 395 Family Myriaporidae Gray, 1841 395 Family Pacificincolidae Liu & Liu, 1999 395 Family Petraliidae Levinsen, 1909 395 Family Phoceanidae Vigneaux, 1949 396 Family Phorioppniidae Gordon & d’Hondt, 1997 397 Family Porinidae d’Orbigny, 1852 397 Family Robertsonidridae Rosso in Rosso et al., 2010 398 Family Stomachetosellidae† Canu & Bassler, 1917 399 Family Tetraplariidae Harmer, 1957 399 Family Teuchoporidae† Neviani, 1896b 400 Family Vicidae Gordon, 1988 400 Superfamily Siphonicytaroidea Harmer, 1957 400 Family Siphonicytaridae Harmer, 1957 400 Superfamily Smittinoidea Levinsen, 1909 401 Family Smittinidae Levinsen, 1909 401 Family Bitectiporidae† MacGillivray, 1895 402 Family Lanceoporidae Harmer, 1957 (1927) 402

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Family Powellithecidae Di Martino, Taylor, Gordon, & Liow, 2016 402 Family Watersiporidae Vigneaux, 1949 403 11.3.7.25 Superfamily Urceoliporoidea Bassler, 1936 404 Family Urceoliporidae Bassler, 1936 404 Family Prostomariidae† MacGillivray, 1895 404 11.3.8 Suborder Skylonina Viskova in Viskova & Morozova, 1988 404 Superfamily Skylonoidea Sandberg, 11.3.8.1 1963 404 Family Bicorniferidae† Keij, 1977 404 Literature 405

Appendix Glossary of common terms in Cheilostomata 425 Systematic classification of Cheilostomata used in this chapter 431 Index

435

List of contributing authors Dr. Andrej Ernst University of Hamburg Institute of Geology Bundesstr. 55 20146 Hamburg Germany [email protected] Dr. Andrei V. Grischenko Perm State National Research University Invertebrate Zoology and Aquatic Ecology Biological Faculty Bukirev Street, 15, GSP 614990 Perm Russia [email protected] A.V. Zhirmunsky National Scientific Center of Marine Biology, Far East Branch, Russian Academy of Sciences Palchevskogo Street 17 690041 Vladivostok Russia Dr. Alexander Gruhl Max Planck Institute for Marine Microbiology Department of Symbiosis Celsiusstr. 1 28359 Bremen Germany agruhl@mpi-bre­men.de Dr. Dennis P. Gordon National Institute of Water and Atmospheric Research Coasts and Oceans 301 Evans Bay Parade 6021 Wellington New Zealand [email protected] Dr. Silviu O. Martha Talstraße 2 72135 Dettenhausen Germany [email protected] Dr. Alvaro E. Migotto University of Sao Paulo Center for Marine Biology Sao Sebastao SP11600-000 Brazil [email protected]

Dr. Andrew N. Ostrovsky University of Vienna Department of Paleontology & St. Petersburg State University Department of Invertebrate Zoology Althanstraße 14, 1090 Wien & Universitetskaya emb. 7/9 St.Petersburg, 199034 Russia [email protected] PD Dr. Thomas Schwaha Universität Wien Department of Evolutionary Biology, Integrative Zoology Althanstraße (UZA I) 14 1090 Wien Austria [email protected] Dr. Javier Souto-Derungs University of Vienna Department of Paleontology Althanstraße 14 1090 Wien Austria [email protected] Dr. Leandro M. Vieira Universidade Federal de Pernambuco Departamento de Zoologia 1235 Av. Prof. Moraes Rego 50670-901 Recife PE Brazil [email protected] Dr. Judith E. Winston Smithsonian Marine Station 701 Seaway Drive Fort Pierce, FL 34949 USA [email protected]

Thomas Schwaha

1 General introduction 1.1 A brief introduction to bryozoans Bryozoa is a phylum of colonial aquatic suspension feeders. To date, bryozoans comprise over 6,500 Recent and over 15,000 fossil species (Bock & Gordon 2013, see also http://bryozoa.net/diversity.html). Almost all species live in marine habitats, with comparatively few species living in fresh or brackish waters. Colonies are predominantly sessile, with a few exceptions, which are either creeping or unattached and capable of moving on the substrate as solitary or colonial forms. The bathymetric distribution of bryozoans ranges from the intertidal to the shallow continental shelf and further to the abyssal zone (Ryland 1970). Some of the deepest recordings of bryozoans exceed over 5,000 m depth (d’Hondt & Hayward 1981). Bryozoans can be found on any kind of substrate and habitats. Most obvious to any common observer, snorkeler, or diver are the large and prominent cheilostome colonies that can be easily observed (Fig. 1.1). For example, these can form large encrusting sheets on stones or other hard substrates or erect branching colonies that form large bushy to tuft-like or frondose to foliate colonies. Bryozoans often show conspicuous bright colorations, e.g. orange, yellow, purple, or many others (Fig. 1.1). Several species are specialized to live in soft to muddy bottoms (e.g. Hirose 2011), in interstitial habitats (Winston & Hakansson 1986, Cook 1988) or are epizooic, living on other animals. Epizooic species are frequently encountered on various parts of crustaceans, pycnogonids, mollusc shells, echinoderms, ascidians, or even vertebrates such as sea turtles or sea snakes (e.g. Ryland 1970, Key et al. 1995, 1996a,b). Coloniality is not a unique feature and shared with some other benthic organisms such as certain cnidarians or ascidians, but unique for bryozoans is that essentially all members of the phylum are colonial. Colonies are modular formations, and each single module or individual is termed zooid in bryozoans. Zooids carry a ciliated tentacle crown or lophophore that is protruded in the open water column and used for suspension feeding (Mukai et al. 1997; Fig. 1.2). Besides coloniality, a distinct and unique feature of all bryozoans is a defensive mechanism: the retraction of the lophophore and associated structures (the polypide) https://doi.org/10.1515/9783110586312-001

into the body wall (or cystid). Body walls are calcified/mineralized in two clades of bryozoans, which evolved independently from uncalcified ancestors (Todd 2000, Ernst & Schäfer 2006). Calcified skeletons also account for the large amount of fossil species, which date back to the Ordovician (see chapter 2). Colonies are formed by asexually produced reproductive stages or by sexually produced larvae. In the latter case, the founding zooid of a colony is termed ancestrula, which forms after settling and metamorphosis of the larva (Reed 1991). Owing to its smaller size, the ancestrula is often distinguishable in colonies (Fig. 1.3). The ancestrula forms several asexually produced buds that increase the size and spreading of the colony. Naturally, the number and growth direction of these buds depend on the species-specific structure and arrangement of the zooids in the colony. Colony development is referred to as astogeny in bryozoans (Ryland 1970). Polypides of bryozoans originate by the formation of a bud; even during metamorphosis of larvae, the formation of the polypide is essentially a budding process (cf. Nielsen 1971). The first indications of polypide buds are discernible as thickening of the epidermis of the body wall and a second adjacent peritoneal (mesodermal) layer. Early buds thus consist of an inner (epidermal) budding layer and an outer (peritoneal) budding layer, which subsequently will form a two-layered vesicle protruding into the zooidal cavity (cf. Schwaha & Wood 2011). This vesicle gradually develops into the polypide in ontogeny. Polypide longevity is rather short and lasts from a week to a month, sometimes also two months (Gordon 1977). After that, it degenerates and the soft tissues are resorbed to form a so-called brown body, a characteristic feature of all bryozoans. With the exception of the Phylactolaemata, polypide recycling is possible in all bryozoans (Taylor & Waeschenbach 2015). In this case, a new polypide can form in the vacant zooid. Knowledge on bryozoan food is not extensive and most data are empirically provided by food given to artificial cultures (cf. Ryland 1976, see also chapter 6). Ingested food predominantly seems to be phytoplankton or bacteria, which are then digested. Some of the large phylactolaemate species often contain entire rotifers in their stomach. In contrast, bryozoans are preyed upon by several organisms, most notably by nudibranch mollusks that rasp of colony pieces or pycnogonids that suck out zooids.

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 1 General introduction

Fig. 1.1: General overview of bryozoans photographed in their natural habitat. (A) Over 25 colonies of the worm-shaped colonies of the phylactolaemate Cristatella mucedo in the Neue Donau, Vienna, Austria. Photo provided by Andrew N. Ostrovsky. (B–D) Macrophotographies provided by Sebastian Decker (Vienna) showing some growth forms and amount of bryozoans in benthic communities. Ctenostome and cheilostomes are visible in B, whereas C has two large cheilostome encrusters forming orange to black sheets. Abundant, single colonies in D are displayed by asterisks. Abbreviations: che – cheilostome, cri – Cristatella, cte – ctenostome.

Other predators include fish that chip off colony pieces or echinoderms such as sea urchins that graze the sea floor (McKinney & Jackson 1989).

1.2 Past summaries on bryozoans Several previous summaries, reviews, and compendia are available on bryozoans that are still a valuable source today: Cori (1941) wrote the original chapter on bryozoans in Handbuch der Zoologie. Brien (1960) compiled a similar summary in the French series Traité de Zoologie in 1960. Almost at the same time Libbie Hyman’s summarizing chapter on bryozoans was published (Hyman 1959). Later,

two excellent summaries on bryozoans were provided by Ryland in 1970 and 1976. Biology of Bryozoans, a book summarizing several aspects of bryozoan biology in 16 chapters was published in 1977 (Woollacott & Zimmer 1977). Some specific aspects included in this book, such as interzooidal communication (Bobin 1977) or ageing in bryozoans (Gordon 1977), have not advanced since that time and are thus still up to date. A bryozoan volume of the series Treatise on Invertebrate Paleontology was released in 1983 (Robinson 1983), which gives a good summary on general features of the main clades. Some years later, McKinney and Jackson (1989) wrote a book on bryozoan evolution, which has some very good information on growth forms and ecological aspects of calcified marine bryozoans.



1.2 Past summaries on bryozoans 

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Fig. 1.2: Live bryozoans showing protruded lophophores. (A) Phylactolaemata, Stephanella hina. (B) Ctenostomata, Flustrellidra hispida. (C) Cyclostomata, Patinella radiata. (D) Disporellid cyclostome. (E) Cheilostomata, Bugula sp. (F&G) Cheilostomata, Electra pilosa.

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Fig. 1.3: Bryozoan ancestrulae and colonial growth. (A) Tubuliporid cyclostome showing ancestrular area (asterisk) where asexual buds initially formed to build an extensive colony. Note the direction of the tubular skeletal extensions of individual zooids. (B) Tubulipord ancestrula with few zooidal buds formed. (C) Ancestrula of the phylactolaemate Plumatella fungosa. (D) Calloporid cheilostome colony showing the small ancestrula in bottom of the image and growing edge on top. (E) Malacostegine cheilostome with ancestrular area (asterisk). Abbreviation: anc – ancestrula.

The first best summary on bryozoan reproduction was provided by Reed (1991) in the book series Reproduction of marine Invertebrates. Mukai et al. (1997) summarized

morphological data of bryozoans in the series Microscopic Anatomy of Invertebrates. Reproduction, especially of Gymnolaemata, was recently summarized by Ostrovsky



(2013). Two volumes on Australian Bryozoa, one covering general aspects of bryozoans and the second a systematic overview of Australian taxa, were released in 2018 (Cook et al. 2018a).

1.3 Topics in bryozoan research During over 160 years of bryozoology, one of the main research areas was taxonomy and systematics – ­naturally with a focus on the diverse skeletal forms. Likewise, paleontology has a long tradition in bryozoan research, and these two disciplines are still the strongest in current times. Scanning electron microscopy, used since the 70s of the last century, opened an entire new perspective and world in bryozoology. It remains the most frequently used tool for imaging specimens and analyzing the skeletal morphology of colonies and zooids. Studies on soft tissues are, in comparison, only few (see chapter 3), and currently, very few research groups work on this topic. Another particular strong emphasis lies in ecological studies that analyze settlement parameters, biotic and abiotic factors affecting bryozoans, growth, substrate preference, inter- and intraspecific competition, interactions with other organisms, etc. The behavior of bryozoans, including feeding, has been studied most diligently by Winston (see chapter 6), reproduction has been studied quite extensively in the past decades by Ostrovsky (2013, see also chapter 4). Embryology was studied only by few researchers (see chapter 5), but an important new study on cleavage pattern and blastomere fates was recently published (Vellutini et al. 2017). Mineralogy of bryozoans has been an important field in bryozoan research (recently summarized by Taylor et al. 2015) and still remains a very important one today (see below). Molecular phylogenetic studies have been applied for less than two decades, with some early works appearing in the beginning of the century (e.g. Dick et al. 2000, 2003). The first molecular studies on phylactolaemates appeared in 2005 and onwards (e.g. Wood & Lore 2005, Hirose et al. 2006, 2008), whereas the most important contributions to the other clades were provided by Waeschenbach et al. (2006, 2009, 2012, 2015). Bryozoans have little to no economic significance. Neither do they deliver any potential food source nor are they a threat to humans. They are an important part of benthic ecosystems, sometimes forming the most abundant group in certain areas. They are generally easily overlooked or not recognized as animals or mistaken, e.g. as corals. The most important human-related topics were recently summarized in volume 1 of Australian bryozoans (Cook et al. 2018a): 1) Bryozoans as biofoulers (Gordon

1.3 Topics in bryozoan research 

 5

2018). Many species frequently settle on many anthropogenic substrates such as ship hulls, which can increase drag and thus fuel consumption and economic costs. In addition, biofouled objects are vectors for bryozoan dispersal, which results in increasing number of invasive species and subsequent changes in natural species distributions and compositions. 2) Bryozoans as source of pharmacologically important substances (Prinsep 2018). Several natural compounds isolated from bryozoans have the potential for pharmacological applications. The earliest and best studied compounds of bryozoans are bryostatins, first isolated in the 80s of the 20th century (Pettit et al. 1982). Clinical trials have shown anticancer effects of bryostatins, but they are also being explored for treating HIV/AIDS or Alzheimer’s disease. 3) Bryozoans and ocean acidification (Smith 2018). Ocean acidification and climate change is a fact, which results in lowered pH and less available carbonate ions. This poses a serious threat on organisms with calcified skeletons such as bryozoans or corals. Experimental approaches on bryozoans aid in monitoring the impact and effects of ocean acidification. Severe gaps are still present in numerous fields of bryozoology: Sensory systems, communication, and colonial integration is a field that is almost not studied at all. Based on descriptive morphological studies, few sensory organs were detected in the tentacles. Modern techniques and functional approaches have not been conducted, however. Likewise, the colonial nervous system and metabolic exchange within a colony remain an open field. A third totally unexplored topic is physiology of bryozoans, with only few experimental observations (Ryland 1970).

1.4 Terminological issues Bryozoans were first regarded as Zoophytes by early naturalists, which grouped several “plant-like” animals such cnidarians or ascidians together with bryozoans. The first specific taxonomic name bestowed upon bryozoans was “Polyzoa” and dates back to Thompson (1830), who first recognized the distinctiveness of bryozoans from other groups of zoophytes. Almost simultaneously, the German naturalist Christian Gottfried Ehrenberg created the name Bryozoa for the same group of animals in 1831. Several decades later, Nitsche divided this clade into “Entoprocta” (Kamptozoa) and “Ectoprocta” (Bryozoa) based on the position of the anus in respect to the tentacle crown of these phyla (Nitsche 1869). Polyzoa was the most common used name for bryozoans in Britain until the 60s of the 20th century (e.g.

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Ryland 1962). Bryozoa is the most commonly used name nowadays and the valid name accepted by the International Bryozoology Association, which was founded in 1965 (Cheetham 2002, see also Cook et al. 2018b). Other names should thus be disregarded and not be used in the future. Polyzoa has also been used for uniting K ­ amptozoa, Cycliophora, and Bryozoa as a related clade (Hejnol et al. 2009), but confuses as it is an old synonym for b ­ ryozoans. The term Ectoprocta is particularly meaningless as a condition, with the anus being located outside the food collecting organ is the more usual situation found in tentaculated suspension feeders; i.e. a phoronid or a cycliophoran is also ectoproct. In contrast, the entoproct condition of kamptozoans is a characteristic feature of this phylum.

1.5 Phylogeny and systematics of bryozoans 1.5.1 Relationship of bryozoans to other phyla 1.5.1.1 The morphological perspective The phylogenetic placement of bryozoans remains quite uncertain. Traditional scenarios unite bryozoans with phoronids and brachiopods into a clade called Lophophorata (Hyman 1959). The latter clade or concept is based on similarities in the morphology of the tentacle crown or lophophore. Lophophorates are coelomate and the lophophore and each tentacle is supplied with a peritoneal lining, either as a separate coelomic compartment as found in phoronids and some brachiopods or is continuous with the lining of the remaining body cavity as in most bryozoans. Tentacle amount varies in the different phyla: in phoronids, it varies from 28 to several hundred with a maximum of about 1,500 (Emig 1982); brachiopods usually start with few tentacles in ontogeny that multiply in later stages and also can reach several hundred. Phylactolaemate bryozoans vary from 20 to 100 tentacles, whereas the predominantly marine ones range from 8 to 30. Lophophorates use upstream-feeding or so-called ciliary sieving (see Riisgard et al. 2010) for particle capture. The basic feeding mechanism is very similar, and also tentacle ciliation for creating feeding currents is identical. All lophophorates show lateral, latero-frontal,

and frontal cilia used for suspension feeding (Riisgard et al. 2010). It has been previously emphasized that tentacle ciliation is monociliated in phoronids and brachiopods versus multiciliated in bryozoans and that this difference might be of phylogenetic significance. In fact, this condition does not seem to be of phylogenetic value as multiciliation of bryozoans is a mere adaptation to smaller size and fewer cells in tentacle cross-section compared to phoronids or brachiopods. The latter two phyla show 40–80 monociliated cells in cross-sectioned tentacles, whereas bryozoans have 9–12 multiciliary ones (cf. Schwaha et al. 2020). Consequently, multiciliation is a necessity for creating proper feeding currents and thus a functional adaptation. In addition to an identical ciliation pattern, tentacle muscles consist of two longitudinal muscle bands in each tentacle in all three phyla (cf. Schwaha & Wanninger 2012). The general body organization of bryozoans is very similar to phoronids; brachiopods are characterized by two mineralized shells that superficially resemble those of bivalve molluscs. A trimeric body organization as previously suggested (e.g. Hyman 1959) is particularly not present in bryozoans and should conceptually be abandoned in these phyla. The digestive system is u-shaped in all lophophorates (when complete in brachiopods). In contrast to bryozoans, brachiopods and phoronids share a blood vascular system and metanephridial system. As coelomate organisms, the latter two are strongly interconnected systems with excretory processes occurring via ultrafiltration at podocytes lining the blood vessels, although metanephridial systems are also capable of removing excretory coelomocytes/phagocytes. Because of their small zooidal size, these two organ systems apparently became redundant and bryozoans lack a distinct blood vascular and nephridial system. Recently, new data of the nervous system of bryozoans and other lophophorates identified significant similarities in the general organization and tentacle innervation in the three lophophorate phyla, which was subsequently used as argument for lophophorate monophyly (cf. Temereva 2017). Bryozoans show a significant difference, however, in regard to the location of the nervous system, which is subepithelial in contrast to a basi-/intraepithelial organization in phoronids and brachiopods. But again – maybe a consequence of miniaturization? An alternative, old hypothesis was that bryozoans could be related to Kamptozoa (e.g. Nitsche 1869, Nielsen 1971). However, there is not any morphological support for such a relationship since they differ e.g. in



their general body organization (acoelomate vs. coelomate), feeding mechanism (downstream vs. upstream, incl. different tentacle ciliation and muscles) and solitary vs. colonial (colonial kamptozoans are later-branching and the solitary loxosomatids are ancestral). In fact, there are merely some similarities in larval morphology and metamorphosis of some species (e.g. Nielsen 2012). Metamorphosis in bryozoans is generally regarded as “catastrophic”; i.e. most organ systems of the larva degenerate and polypides are actually formed de novo by a budding process in the settled ancestrula (Mukai et al. 1997). Kamptozoan metamorphosis in the early branching loxosomatids involves mostly elongation of the anteriorposterior axis to form the stalk, reduction of the apical organ and foot and reorganization of the oral side of the larva to form the tentacle crown (cf. Nielsen 1971, 2012). Several molecular phylogenies reconstruct a closer relationship of phoronids and brachiopods with other phyla, such as nemerteans (e.g. Nesnidal et al. 2010) or annelids (Kocot et al. 2017). Irrespective of the monophyletic state of the Lophophorata, it remains difficult to link the three phyla to other lophotrochozoans, especially in regard to body plan evolution. Back in the 20th century, a close association of lophophorates to deuterostomes, particularly to the morphologically similar pterobranchs, was taken into account (e.g. Salvini-Plawen 1982). However, since the emergence of molecular phylogenetic analyses, it became clear that these phyla are lophotrochozoans and not related to any deuterostomes.

1.5 Phylogeny and systematics of bryozoans 

 7

phylogenomic data are just starting to emerge (Kocot et al. 2017). Currently, however, phylogenomic data on bryozoans is only available for a few cheilostomes, a group of late branching bryozoans. In the near future, new phylogenomic data from other, more early-branching groups will probably yield new trees that will shed new light on the phylogenetic position of bryozoans.

1.5.2 Internal systematics and phylogeny of bryozoans

Bryozoans are divided into three distinct clades: Phylactolaemata, Stenolaemata, and Gymnolaemata. Phylactolaemata is a small group of sole freshwater inhabitants characterized by a horseshoe-shaped lophophore and dormant buds called statoblasts. Less than 100 species have been described in this clade. Stenolaemata is one of the taxa that evolved calcified cystid walls and comprises over 11,000 species, including Recent and fossil ones. With the exception of the Cyclostomata, all the other taxa of stenolaemates were almost exclusively present in the Paleozoic and are now extinct (see chapter 2). Approximately 700 Recent species of Cyclostomata are described. Gymnolaemata is the dominant group that contains the paraphyletic “Ctenostomata” and the monophyletic Cheilostomata. Fossil and Recent species included, over 11,000 species have been described, but in contrast to stenolaemates, over 5,800 Recent species of gymnolaemates are currently described. Ctenostomes are a small group of ~350 species, whereas the bulk 1.5.1.2 The molecular perspective of gymnolaemates are cheilostomes (http://bryozoa. net/diversity.html). The latter is the second clade that Whereas morphologically the lophophorates are cur- evolved calcified cystid walls, hence their large fossil rently the most feasible concept, molecular phylogenetic record. analyses were hardly able to reconstruct lophophorates. Originally the term Gymnolaemata created by Allman A close relationship of the phoronids and brachiopods is (1856) included stenolaemates and gymnolaemates, but supported in most phylogenies (cf. Kocot 2016), but bry- it became accustomed that Gymnolaemata is restricted to ozoans generally drop out. Molecular analyses placed the clade of Cteno- and Cheilostomata. An outdated name bryozoans at various positions clustering with different for the current Gymnolaemata was Eurystomata (Marcus lophotrochozoan phyla (e.g. Hausdorf et al. 2007, 2010, 1938), which, however, was only used by very few authors Helmkampf et al. 2008, Hejnol et al. 2009, Mallatt et al. and has not been applied to any larger work on bryozoans. 2012), but none of the placements seemed to repeatedly Likewise, new descriptions of cteno- and cheilostomes persist. Only recent molecular phylogenetic studies were in the past decades almost exclusively used the name able to reconstruct lophophorates (Nesnidal et al. 2013, Gymnolaemata (see also Schwaha et al. 2020). 2014, Marlétaz et al. 2019). Addressing some of the subgroups of the large clades Most molecular phylogenies were based on fast-­ often has two distinct spellings that can be found disevolving mitochondrial genes or nuclear ribosomal genes. persed in publications: cyclostome vs. cyclostomate, ctenMore robust phylogenetic reconstructions involving ostome vs. ctenostomate, cheilostome vs. cheilostomate.

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 1 General introduction

Literature

Fig. 1.4: Phylogeny of the major bryozoan clades, after Schwaha et al. (2020), general topology redrawn from Taylor & Waeschenbach (2015).

The former version of these spellings is the more common and nowadays used term. The interrelationship of the three large clades is well supported, with Phylactolaemata being the sister-group of a clade of Stenolaemata and Gymnolaemata. The latter two are sister-groups (Waeschenbach et al. 2012, Taylor & Waeschenbach 2015; Fig. 1.4). Recently, the name “Myolaemata” was proposed for the clade comprising Stenolaemata and Gymnolaemata. Distinct characters of this taxon are: 1) myoepithelial pharynx with triradiate lumen used for suction feeding, 2) circular lophophore (even though this could also be a plesiomorphic character), 3) pylorus with cilia and specific mode of digestion (see chapter 3), 4) lophophoral coelomic cavity in the form of a simple ring canal (see chapter 3), 5) anal growth direction (opposed to the oral one of Phylactolaemata, cf. Jebram 1973), 6) zooidal polymorphism, and 7) polypide recycling (Schwaha et al. 2020).

1.6 Outline of this book The main aim of this book is to give a thorough insight in the phylum Bryozoa. Chapters 2–6 are general, comparative chapters on bryozoan biology that have not been recently summarized. Several topics such a phylogeny or biomineralization have thus not been included into this volume as they are subject to change in the near future in case of phylogeny or have been recently summarized (Taylor et al. 2015). Chapter 2 is a short introduction into bryozoan paleontology, because understanding bryozoan evolution requires a necessary background into this old phylum. The remaining book is focused on Recent Bryozoa and chapters 3–5 deal with morphology (predominantly of soft-tissues), reproduction and development. Chapter 6 deals with the behavior of bryozoans, whereas the remaining chapters 7–11 introduce the systematic groups.

Allman, G.J. (1856): A Monograph of the Fresh-Water Polyzoa. Ray Society Publications 28. Ray Society, London: 119 pp. Bobin, G. (1977): Interzooecial communications and the funicular system. In: Woollacott, R.M. & Zimmer, R.L. (eds.). Biology of Bryozoans. Academic Press, New York: 307–333. Bock, P. & Gordon, D.P. (2013): Phylum Bryozoa Ehrenberg, 1831. Zootaxa 3703: 67–74. Brien, P. (1960): Classe des Bryozoaires. In: Grassé, P.P. (ed.). Traité de Zoologie. Masson, Paris: 1053–1335. Cheetham, A.H. (2002): The founding and early history of the International Bryozoology Association, 1965–1974. In: Wyse Jackson, P.N. & Spencer-Jones, M.E. (eds.). Annals of Bryozoology: Aspects of the History of Research on Bryozoans. International Bryozoology Association, Dublin: 45–57. Cook, P.L. (1988): Bryozoa. In: Higgin, R.P. & Theil, H. (eds.). Introduction to the Study of Meiofauna. Smithsonian Institution Press, Washington, D.C.: 438–443. Cook, P.L., Bock, P.E., Gordon, D.P. & Weaver, H.J. (2018a): Australian Bryozoa, Volume 1&2. CSIRO Publishing, Melbourne. Cook, P.L., Gordon, D.P., Hayward, P.J., Bock, P.E. & Bone, Y. (2018b): Introducing bryozoans. In: Cook, P.L., Bock, P.E., Gordon, D.P. & Weaver, H.J. (eds.). Australian Bryozoa, Volume 1. CSIRO Publishing, Melbourne: 1–16. Cori, C.J. (1941): Ordnung der Tentaculata: Bryozoa. In: Kükenthal, W. & Krumbach, T. (eds.). Handbuch der Zoologie. De Gruyter, Berlin: 263–502. Dick, M.H., Freeland, J.R., Williams, L.P. & Coggeshall-Burr, M. (2000): Use of 16S mitochondrial ribosomal DNA sequences to investigate sister-group relationships among gymnolaemate bryozoans. In: Herrera Cubilla, A. & Jackson, J.B.C. (eds.). Proceedings of the 11th International Bryozoology Association Conference. Smithsonian Tropical Research Institute, Balboa: 197–210. Dick, M.H., Herrera Cubilla, A. & Jackson, J.B.C. (2003): Molecular phylogeny and phylogeography of free-living Bryozoa (Cupuladriidae) from both sides of the Isthmus of Panama. Mol Phylogenet Evol 27: 355–371. d’Hondt, J.-L. & Hayward, P.L. (1981): Nouvelles recoltes de Bryozoaires Cténostomes bathyaux et abyssaux. Cah Biol Mar 22: 267–283. Emig, C. (1982): The Biology of Phoronida. Adv Mar Biol 19: 1–89. Ehrenberg, C.G. (1831): Symbolae Physicae, seu Icones et Description Mammalium, Avium, Insectorum et Animalium Evertebratorum. Ex Officina Academica, Berlin. Ernst, A. & Schäfer, P. (2006): Palaeozoic vs. post-Palaeozoic Stenolaemata: Phylogenetic relationship or morphological convergence? Courier Forschungsinstitut Senckenberg 257: 49–64. Gordon, D.P. (1977): The ageing process in bryozoans. In: Woollacott, R.M. & Zimmer, R.L. (eds.). Biology of Bryozoans. Academic Press, New York: 335–376. Gordon, D.P. (2018): Bryozoans and biosecurity. In: Cook, P.L., Bock, P.E., Gordon, D.P. & Weaver, H.J. (eds.). Australian Bryozoa, Volume 1. CSIRO Publishing. Melbourne: 71–90. Hausdorf, B., Helmkampf, M., Meyer, A., Witek, A., Herlyn, H., Bruchhaus, I., Hankeln, T., Struck, T.H. & Lieb, B. (2007): Spiralian phylogenomics supports the resurrection of bryozoa comprising ectoprocta and entoprocta. Mol Biol Evol 24: 2723–2729.

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Mukai, H., Terakado, K. & Reed, C.G. (1997): Bryozoa. In: Harrison, F.W. &, Woollacott, R.M. (eds.). Microscopic Anatomy of Invertebrates. Wiley-Liss, New York, Chichester: 45–206. Nesnidal, M.P., Helmkampf, M., Bruchhaus, I. & Hausdorf, B. (2010): Compositional Heterogeneity and Phylogenomic Inference of Metazoan Relationships. Mol Biol Evol 27: 2095–2104. Nesnidal, M., Helmkampf, M., Meyer, A., Witek, A., Bruchhaus, I., Ebersberger, I., Hankeln, T., Lieb, B., Struck, T. & Hausdorf, B. (2013): New phylogenomic data support the monophyly of Lophophorata and an Ectoproct-Phoronid clade and indicate that Polyzoa and Kryptrochozoa are caused by systematic bias. BMC Evol Biol 13:1–13. Nesnidal, M.P., Helmkampf, M., Bruchhaus, I., Ebersberger, I. & Hausdorf, B. (2014): Lophophorata monophyletic – after all. In: Wägele, J.W. & Bartolomaeus, T. (eds.). Deep Metazoan Phylogeny: The Backbone of the Tree of Life: New Insights from Analyses of Molecules, Morphology, and Theory of Data Analysis. De Gruyter, Berlin, Boston: 127–142. Nielsen, C. (1971): Entoproct life-cycles and the Entoproct/Ectoproct relationship. Ophelia 9: 209–341. Nielsen, C. (2012): Animal Evolution. Interrelationships of the Living Phyla. University Press, Oxford: 402 pp. Nitsche, H. (1869): Beiträge zur Kenntniss der Bryozen. 1. Beobachtungen über die Entwicklungsgeschichte einiger chilostomen Bryozoen. 2. Ueber die Anatomie von Pedicellina echinata SARS. Z Wiss Zool 20: 1–36. Ostrovsky, A.N. (2013): Evolution of Sexual Reproduction in Marine Invertebrates: Example of Gymnolaemate Bryozoans. Springer, Dordrecht, Heidelberg, New York, London: 356 pp. Pettit, G.R., Herald, C.L., Doubek, J.M. & Herald, D.L. (1982): Isolation and structure of bryostatin-1. J Am Chem Soc 104: 6846–6848. Prinsep, M. (2018): Bryozoans and biotechnology. In: Cook, P.L., Bock, P.E., Gordon, D.P. & Weaver, H.J. (eds.). Australian Bryozoa, Volume 1. CSIRO Publishing. Melbourne: 121–138. Reed, C.G. (1991): Bryozoa. In: Giese, A.C., Pearse, J.S. & Pearse, V.B. (eds.). Reproduction of marine Invertebrates VI Echinoderms and Lophophorates. The Boxwood Press, Pacific Grove, California: 85–245. Riisgard, H.U., Okamura, B. & Funch, P. (2010): Particle capture in ciliary filter-feeding gymnolaemate and phylactolaemate bryozoans – a comparative study. Acta Zool 91: 416–425. Robinson, R.A. (1983) Treatise on Invertebrate Paleontology: Part G: Bryozoa. Geological Society of America and University of Kansas, Boulder and Lawrence. Ryland, J.S. (1962): Biology and identification of intertidal Polyzoa. Field Studies 1(4): 33–51. Ryland, J.S. (1970): Bryozoans. Hutchinson University Library, London. Ryland, J.S. (1976): Physiology and ecology of marine bryozoans. In: Russel, F.S. & Yonge, M. (eds.). Advances in Marine Biology Vol 14. Academic Press, London: 285–443. Salvini-Plawen, L. (1982): A paedomorphic origin of oligomerous animals. Zool Scr 11: 77–81. Schwaha, T. & Wood, T.S. (2011): Organogenesis during budding and lophophoral morphology of Hislopia malayensis Annandale, 1916 (Bryozoa, Ctenostomata). BMC Dev Biol 11: 23. Schwaha, T. & Wanninger, A. (2012): Myoanatomy and serotonergic nervous system of plumatellid and fredericellid phylactolaemata (lophotrochozoa, ectoprocta). J Morphol 273: 57–67.

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 1 General introduction

Schwaha, T., Ostrovsky, A.N. & Wanninger, A. (2020): Key novelties in the evolution of aquatic colonial invertebrates: Evidence from soft body morphology of Bryozoa. Biological Reviews 95: 696–729. Smith, A.M. (2018): Bryozoans and ocean acidification. In: Cook, P.L., Bock, P.E., Gordon, D.P. & Weaver, H.J. (eds.). Australian Bryozoa, Volume 1. CSIRO Publishing, Melbourne: 139–144. Taylor, P.D. & Waeschenbach, A. (2015): Phylogeny and diversification of bryozoans. Palaeontology 58: 585–599. Taylor, P.D., Lombardi, C. & Cocito, S. (2015): Biomineralization in bryozoans: present, past and future. Biol Rev Camb Philos Soc 90: 1118–1150. Temereva, E.N. (2017): Morphology evidences the lophophorates monophyly: brief review of studies on the lophophore innervation. Invertebrate Zoology 14: 85–91. Thompson, J.V. (1830): On Polyzoa, a new animal discovered as an inhabitant of some Zoophites – with a description of the newly instituted genera of Pedicellaria and Vesicularia, and their species. Zoological Researches, and illustrations; or, Natural History of Nondescript or Imperfectly Known Animals. Vol. 1. King and Ridings, Cork: 89–102. Todd, J.A. (2000): The central role of ctenostomes in bryozoan phylogeny. In: Herrera Cubilla, A. & Jackson, J.B.C. (eds.). Proceedings of the 11th International Bryozoology Association Conference. Smithsonian Tropical Research Institute, Balboa: 104–135. Vellutini, B.C., Martin-Duran, J.M. & Hejnol, A. (2017): Cleavage modification did not alter blastomere fates during bryozoan evolution. BMC Biol 15: 33.

Waeschenbach, A., Telford, M.J., Porter, J.S. & Littlewood, D.T.J. (2006): The complete mitochondrial genome of Flustrellidra hispida and the phylogenetic position of Bryozoa among the Metazoa. Mol Phylogenet Evol 40: 195–207. Waeschenbach, A., Cox, C.J., Littlewood, D.T.J., Porter, J.S. & Taylor, P.D. (2009): First molecular estimate of cyclostome bryozoan phylogeny confirms extensive homoplasy among skeletal characters used in traditional taxonomy. Mol Phylogenet Evol 52: 241–251. Waeschenbach, A., Taylor, P.D. & Littlewood, D.T.J. (2012): A molecular phylogeny of bryozoans. Mol Phylogenet Evol 62: 718–735. Waeschenbach, A., Vieira, L.M., Reverter-Gil, O., Souto-Derungs, J., Nascimento, K.B. & Fehlauer-Ale, K.H. (2015): A phylogeny of Vesiculariidae (Bryozoa, Ctenostomata) supports synonymization of three genera and reveals possible cryptic diversity. Zool Scr 44: 667–683. Winston, J.E. & Hakansson, E. (1986): The interstitial Bryozoan fauna of the Capron Shoals, Florida. Am Mus Novit 2865: 1–98. Wood, T.S. & Lore, M. (2005): The higher phylogeny of Phylactolaemate bryozoans inferred from 18S ribosomal DNA sequences. In: Moyano, C., Cancino, J.M. & Wyse Jackson, P.N. (eds.). Bryozoan Studies 2005. A.A. Balkema Publishers, Leiden, London, New York, Philadelphia, Singapore: 361–367. Woollacott, R.M. & Zimmer, R.L. (1977): Biology of Bryozoans. Academic Press, New York: 566 pp.

Andrej Ernst

2 Fossil record and evolution of Bryozoa 2.1 Introduction The Phylum Bryozoa has a long and extensive fossil record (Fig. 2.1) counting ca 15,000 fossil species (Horowitz & Pachut 2000, Gordon et al. 2009). Their remnants are found in the majority of marine sediments of the Phanerozoic. They played important roles in various communities and were significant contributors to carbonate sedimentation and reefs. Bryozoans belong to the few phyla of which no fossilized representatives are known during the Cambrian explosion. They were repeatedly reported from the strata prior to the Ordovician, but which appeared to be findings of non-bryozoan nature (see reviews in Ross 1964, 1985, Taylor & Ernst 2004). Most recently, Landing et al. (2010, 2015) described fossils from the lower Tiñu Formation of the Upper Cambrian of Mexico, which they claimed were bryozoans. However, their assignment to bryozoans appears doubtful (Taylor et al. 2013). Confirmed records of earliest bryozoans are known from the Early Ordovician (Tremadoc) of China (Ma et al. 2015). Calcified bryozoans appeared in the Early Ordovician of China and diversified very fast (Hu & Spjeldnaes 1991, Xia et al. 2007, Ma et al. 2015). The majority of Palaeozoic bryozoan faunas were distributed in tropics, whereas the post-Palaeozoic bryozoans tend to occur in temperate or even cool-water environments (Taylor & Alison 1998, Taylor & Sendino 2010). The majority of Palaeozoic bryozoans belong to the Class Stenolaemata, although gymnolaemates (burrowing Ctenostomata) are also known throughout the Palaeozoic. Ma et al. (2014) introduced the suborder Palaeostomata for the free-walled (i.e. without calcified frontal walls, in contrast to those with calcified frontal walls and therefore called fixed-walled; see chapter 8) “Palaeozoic” stenolaemates including the orders Cystoporata, Esthonioporata, Trepostomata, Cryptostomata, Fenestrata, and the group of Timanodictyina, whose position is still under discussion. The authors also erected the Order Esthonioporata, elevating its status from suborder to order level. All the Palaeostomata except Timanodictyina appeared and diversified during the Great Ordovician Biodiversification Event (Taylor & Ernst 2004, Ernst 2018). The majority of palaeostomates (as well as cyclostomes) possessed a calcitic skeleton with a stable low-Mg content, allowing excellent preservation of fossil bryozoans. The representatives of the Order Esthonioporata Ma https://doi.org/10.1515/9783110586312-002

et al., 2014 may have possessed high-Mg calcite in their skeleton (Taylor & Wilson 1999). The sister group of Palaeostomata, the Order Cyclostomata, appeared in the Early Ordovician, too. They are also calcified bryozoans, but in contrast to Palaeostomata, cyclostomes survived the end-Palaeozoic and Triassic crises and radiated during the Mesozoic (Schäfer & FoisErikson 1987, Taylor & Ernst 2008). This group includes some few hundreds of species in the modern seas and gives clues for understanding the morphology of the Palaeostomata, which are not represented in modern faunas (Boardman 1971, Boardman et al. 1992, Ernst & Schäfer 2006). The gymnolaemate Order Ctenostomata appeared in the Ordovician, too. Borings belonging to endolithic ctenostomes were discovered in the Early Ordovician of Spain and Russia (Mayoral 1991, Mayoral et al. 1994, Taylor & Rozhnov 1996). Boring ctenostomes are known from Ordovician to present (e.g. Pohowsky 1974, 1978, Viskova & Pakhnevich 2010). Otherwise, the fossil record of ctenostomes is rather scarce. Normally, uncalcified ctenostome bryozoans (as other soft-bodied invertebrates) have little chance for preservation. Due to organic overgrowth of unmineralized organisms, their natural molds may be formed. This process is called bioimmuration (Vialov 1961). Such fossils usually appear as negative relief and often show exceptional details of the surface of the bioimmured organism (Taylor 1990b). Bioimmured ctenostomes are mainly known from the Jurassic and Cretaceous (Voigt 1977, Taylor 1978, 1990a,b, Todd 1994, Todd et al. 1997). Cheilostome bryozoans derived most certainly from ctenostome-grade gymnolaemates during the Jurassic (Pohowsky 1973, Taylor 1981a, 1986, 1994). This group diversified during the Cretaceous and especially in the post-Cretaceous time until the present (Taylor & Larwood 1990, Lidgard et al. 1993, Jablonski et al. 1997, Sepkoski et al. 2000). Phylactolaemata are the group of entirely freshwater bryozoans, which are regarded as earliest branch of all bryozoans (e.g. Fuchs et al. 2009, Waeschenbach et al. 2012, Taylor & Waeschenbach 2015). Phylactolaemates are not calcified; their bodies are covered by gelatinous or chitinous material (e.g. Ryland 1970, Wöss 2005, see also chapter 7); therefore their preservation potential is very low. However, phylactolaemates produce dormant stages called statoblasts that possess chitinous shells. Fossil

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Fig. 2.1: Ranges and approximate generic diversity of higher taxa of stenolaemates (left, blue) and gymnolaemates (right, red). Dark blue: Palaeostomata (modified after McKinney & Jackson 1989).

statoblasts are known from few records, with the oldest from the Permian (Vinogradov 1996) of Russia and Triassic of South Africa (Kohring & Hörnig 2002). The reason for the missing earlier records might be that statoblasts represent an adaptation of living in freshwater, whereas the earliest phylactolaemate ancestors were supposedly marine and therefore did not produce statoblasts (Taylor & Waeschenbach 2015). The present chapter gives an outline of the evolutionary history of bryozoans and their fossil record, as well as some detailed insights in the morphology and general taxonomy of Palaeostomata.

2.2 Methods for studying fossil bryozoans In contrast to cyclostomes and cheilostomes, the Palaeostomata display a great variety of internal characters that cannot be studied externally. Therefore, the most important method of their study is the use of oriented thin sections. Three orientations are generally needed: tangential (showing the character of autozooecial apertures, heteromorphs, and skeletal structures such as styles), longitudinal, and transversal, which reveal the shape of autozooecia and their internal structures (Figs. 2.2 and 2.5 A–C). Alternatively, the methods of acetate peels can be used for studying the internal morphology of palaeostomates (Boardman & Utgaard 1964). This method works on well preserved carbonate material and is not appropriate for silicified or dolomitized rocks. The idea of this method is based on the solution of carbonate by a weak acid with creation of a relief, which can be replicated by plastic softened by acetone. For the preparation of peels, the specimen is cut and polished as for the preparation of thin sections. Then its surface is put in weak acid (hydrochloric or formic in 1%–5% solution) for 20–40 seconds. After washing and drying,

the etched surface is doused by acetone and covered by a piece of acetone-soluble plastic. After evaporation of the acetone, the plastic is peeled (the name!) from the sample, with the relief replicated on its surface. There are advantages and disadvantages of both methods. Thin sections deliver highest quality, but they are fragile and their preparation destroys the sample or at least a large portion of it. Acetate peels are worse in quality, they often curve, and the high relief prevents production of high-magnification photographs. However, the samples are less affected by the preparation of peels (important in case of museum material), and it is possible to produce new peels from them. In fact, this method can be used to produce serial peels in distances of few microns and to arrange these images to a three-dimensional model using visualization software. External characters such as micro- and ultrastructure of autozooecial walls can be studied using scanning electron microscopy (SEM) (e.g. Taylor & Jones 1996). Useful are also analytic methods such as Raman spectroscopy (Taylor et al. 2008, Di Martino et al. 2016) or x-ray diffraction analysis (Fortunato et al. 2012). Post-Palaeozoic bryozoans (cyclostomes and cheilostomes) are studied almost exclusively by use of the SEM.

2.3 Development and evolution of bryozoans in the Palaeozoic Bryozoans of the Palaeozoic age are represented by the classes Gymnolaemata and Stenolaemata. The records of Palaeozoic Gymnolaemata are exclusively boring ctenostomes known since the Early Ordovician. Several genera were described from the Palaeozoic: Ropalonaria Ulrich, 1879 (Fig. 2.10 D), Vinella Ulrich, 1890, Condranema Bassler, 1952, Eliasopora Bassler, 1952, Casteropora Pohowsky, 1978, Bascomella Morningstar, 1922, and Orbignyopora Pohowsky, 1978. The genera Allonema



2.3 Development and evolution of bryozoans in the Palaeozoic 

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Fig. 2.2: Cutaway diagram of a segment of a branched colony of the Palaeostomata showing orientations of standard sections used for study (modified after Boardman 1984).

Ulrich & Bassler, 1904 and Ascodictyon Nicholson & Etheridge, 1877 were originally described as ctenostome bryozoans. However, their closer study showed that they have no apertures for lophophore protrusion and apparently represent encrusting bases of unknown articulated organisms (Wilson & Taylor 2014). Jarochowska and Munnecke (2014) suggested that Allonema Ulrich & Bassler, 1904 is a senior synonym for Wetheredella Wood, 1948, an encrusting problematicum widely distributed in the Palaeozoic. The Class Stenolaemata Borg, 1926 comprises bryozoans with complete interior vertical walls producing tubular, conical, or sac-shaped zooecia (Boardman 1983). The modern classification favors the division of Stenolaemata in two superorders: Palaeostomata and Tubuliporata. The latter comprises the Order Cyclostomata, which ranges from the Early Ordovician to the Recent. This clade is regarded as monophyletic (e.g. Brood 1973, Viskova 1992, Taylor & Waeschenbach 2015). Alternative hypotheses suggest the polyphyletic origin of cyclostomes (Boardman 1984, Ernst & Schäfer 2006). The Palaeostomata derived apparently from tubuliporate cyclostomes ( = Palaeotubuliporina Brood, 1973). In contrast to cyclostomes, the Palaeostomata are

exclusively free-walled, lacking calcified exterior walls (e.g. Boardman 1971, 1998, Boardman & Cheetham 1973). In contrast, cyclostomes may also possess a so-called fixed-walled skeletal organization having a calcified exterior wall (see chapter 8, this volume). Curiously, some Early Palaeozoic ceramoporids (Order Cystoporata) possess communication pores in their internal walls, a structure known from cyclostomes with calcified exterior walls (Fig. 2.9 E). The communication pores are necessary because exterior walls prevent fluid exchange and communication between autozooecia. In free-walled stenolaemates communication between zooecia occurs above the interior walls. Communication pores in cystoporates apparently evolved independently from those of the post-Palaeozoic cyclostomes (e.g. Boardman 1998). The Superorder Palaeostomata Ma et al., 2014 comprises fewer than 600 genera distributed from the Early Ordovician to the Late Triassic and includes the orders Cystoporata, Esthonioporata, Trepostomata, Cryptostomata, Fenestrata, and the group of Timanodictyina. The latter is recognized as a separate order by Gorjunova (1992, 1994), who postulated their origin from cystoporate bryozoans. Alternatively, timanodictyines are regarded

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being related to cryptostomes (Blake 1983a). The Palaeostomata are indeed the dominant bryozoan group in the Palaeozoic, representing more than 95% of total generic composition at that time.

2.3.1 Morphology of the Palaeostomata 2.3.1.1 Definition of Palaeostomata Ma et al. (2014) separated the Palaeozoic orders of freewalled stenolaemates into the Superorder Palaeostomata. They proposed the following definition: Hemispherical, massive, ramose, frondose, reticulate or encrusting, occasionally articulated colonies. Maculae evenly spaced, present on colony surfaces of broadly multiserial taxa, monticulate, flat or depressed. All skeletal walls above basal exterior colony wall interior in origin, calcified frontal exterior walls lacking. Autozooecia short to long, polygonal to rounded in cross-section, apertures sometimes with lunaria. Walls usually with a lamellar microstructure, occasionally granular, typically non-porous; pseudopores wanting. Basal diaphragms common to absent; hemiphragms, ring septa and cystiphragms present or absent. Styles often present. Mesozooecia and exilazooecia commonly present. Apparent brooding structures in some groups present. (Ordovician – Triassic)

Palaeostomates differ from cyclostomes in some important morphological characters. The key feature of the Superorder Palaeostomata is the absence of calcified exterior walls above the basal lamina (Borg 1926, Boardman 1983). Furthermore, palaeostomates possess various kinds of intrazooecial partitions (basal diaphragms, cystiphragms, hemiphragms, etc.), which are rare or absent in cyclostomes (Boardman 2001).

2.3.1.2 Morphology of Palaeostomata Palaeostomata are often known as “stony bryozoans” because of their heavy calcification. The majority of palaeostomates possess a well-developed external calcitic skeleton. In contrast to other Palaeozoic colonial animals such as tabulate and rugose corals or stromatoporoids, bryozoan colonies did not reach large sizes. The largest colonies of Palaeostomata are known from the Trepostomata and are mostly restricted to the Ordovician or Permian (e.g. Ross & Ross 1962, Håkansson & Madsen 1991, Key et al. 2005). Massive trepostome bryozoans from the Ordovician of Estonia reach sizes of 30 cm in diameter and 10 cm in height. Branched trepostomes from the Ordovician of the USA reached heights of up to 66 cm (Cuffey & Fine

2005). Late Palaeozoic fenestrates are known up to 30 cm in height (Wood et al. 1996). However, the majority of Palaeozoic bryozoans are smaller, having sizes of few centimeters or even smaller. As colonial (modular) organisms, bryozoans produce colonies by proliferation of new zooids and extrazooidal tissue (e.g. Ryland 1981, McKinney & Jackson 1989). The form of the colony (or growth form, growth habit) is determined by the position of new zooids in respect to those formed earlier including their shape, orientation, and rate of addition in each portion of the colony. Bryozoans are able to develop colonies of various forms that are of immense adaptive importance. Therefore, a significant number of attempts to use colony forms for palaeoecological studies, especially as indicators for depth and water energy, were undertaken (e.g. Stach 1936, Nelson et al. 1988, Hageman et al. 1997, 1998, Amini et al. 2004). In fact, observations on Mediterranean bryozoans from different habitats revealed distinct plasticity in colony forms according to the habitat (Harmelin 1973, 1976). Simplest colony forms are encrusting and represented by runners (Fig. 2.9 D) and sheets (Figs. 2.9 E and 2.14 D). The latter may be a single layer (Fig. 2.13 K) or multilayered (Figs. 2.4 B and 2.12 F). Massive colonies are especially common among early taxa of Esthonioporata, Cystoporata, and Trepostomata (Figs. 2.4 A, 2.9 G, and 2.10 I). Erect branched colonies occur among these three groups (especially Trepostomata) but are also typical for rhabdomesine cryptostomes (Figs. 2.5 A–C, N, O, 2.6 D–M, 2.10 B–C, 2.11 G–K, 2.12 G–J, 2.13 C–E, and 2.14 A–C). Many cyclostomes and cheilostomes develop erect branched colonies, too, which can be rigid or flexibly connected (Fig. 2.14 F, G). Among erect (arborescent) colonies, unilaminate and bilaminate (Figs. 2.4 F–H, 2.6 A–C, 2.9 J, 2.10 J, 2.11 D–F, and 2.13 B, M), or even trilaminate (Fig. 2.4 E), forms can be distinguished. Unilaminate erect colonies can be of various shape and are especially diverse in Palaeozoic fenestrates (Figs. 2.7 A–I, 2.9 I, 2.10 J, 2.11 B, 2.12 A, K–M, and 2.13 A, G–I). A special character of the colony surface are so-called maculae, areas of the colony surface with different morphology. They usually consist of larger zooecia (e.g. macrozooecia in trepostomes) or smaller polymorphs like mesozooecia and exilazooecia, as well as vesicular skeleton or extrazooidal skeleton. Many maculae are elevated above the colony surface and in that case are called monticulae (Fig. 2.4 I). Others are rather depressed or positioned at the level of the colony surface. Maculae are usually rounded or oval in shape, but also stellate ones are common, as for example in the cystoporate genus Constellaria or in the Devonian genus Stellatoides (Ernst et al. 2014), or in large trepostomes from the Permian of



2.3 Development and evolution of bryozoans in the Palaeozoic 

Greenland (Key et al. 2002, 2011). The major interpretation of maculae is their function in the regulation of feeding currents (see section 2.6.1.2). Maculae are common in the Palaeostomata as well as in branched post-Palaeozoic Cyclostomata. Palaeostomata mainly had low-Mg calcitic skeletons (Smith et al. 2006), whereas esthoniporines seem to have had high-Mg calcite (Taylor & Wilson 1999, Ma et al. 2014). The skeleton in Palaeostomates is usually laminated, normally in the exozone, or rather hyaline, amorphic in the endozone or in the cores of styles (Tavener-Smith 1969a,b, Armstrong 1970, Blake 1973). In their appearance, the laminated internal walls may be merged, without visible zooecial boundary, or serrated, with distinct boundary between adjacent zooecia (Boardman & Buttler 2005). Fenestrate and cryptostome bryozoans had extensive sheets of external laminated skeletal material, whereas the majority of cystoporates developed a special vesicular skeleton, which filled the space between autozooecia (Utgaard 1983). Such a vesicular skeleton is in less extent also present in other groups such as in trepostomes (Boardman & Buttler 2005), in ptilodictyines (Karklins 1983, Gorjunova & Lavrentjeva 1993), and in fenestrates (Morozova 2001). A vesicular skeleton is not known from rhabdomesines (Blake 1983b, Gorjunova 1985, 1996). Kenozooids and rootlets are known in fenestrates (Morozova 2001). Different types of styles are known in various groups of Palaeostomata. Their function is mainly regarded as protective or structural (Tavener-Smith 1969a, Armstrong 1970, Blake 1973, 1983b, Boardman & Cheetham 1973, Boardman & Buttler 2005). It is supposed that the function of acanthostyles might be to raise exterior membranous walls above zooecial apertures and skeletal surfaces in order to improve communication between zooids (Boardman 1983). Acanthostyles are the principal type of styles consisting of hyaline cores surrounded by laminated sheaths (e.g. Figs. 2.5 H, I and 2.6 M). In the Permian genus Dyscritellina, acanthostyles are as large as zooecia (Fig. 2.5 I). They usually protrude above the colony surface, whereas the others such as paurostyles, heterostyles, or aktinotostyles are partly or completely embedded in the skeleton (Blake 1983b). Different kinds of inhomogeneities such as tubules and spherules are known in the skeleton of mainly trepostomes, but also some cryptostomes (Boardman & Buttler 2005). Especially erydotrypellid and ulrichotrypellid trepostomes are characterized by the presence of such structures (Fig. 2.5 L–O). Mural (or zooecial) spines are spine-like structures developed inside of autozooecia and occur mainly in trepostomes, being rare in cryptostomes (Fig. 2.3 G, H). They are

 15

usually interpreted as ligament attachment points (Boardman 1960b, 1971, Farmer 1979). Structures related to brooding are known in some Palaeostomata, especially in fenestrates (Tavener-Smith 1966, Stratton 1981, Southwood 1985, Bancroft 1986a, 1988, Morozova 2001). They are mostly represented by different kinds of rounded chambers attached to autozooecia (Fig. 2.7 E, G). Some few apparent brooding structures are known in cystoporates (Utgaard 1973, Buttler 1991, Pachut & Horowitz 2013). Distinct reproductive structures are still unrevealed in trepostomes, rhabdomesines and ptilodictyines, although large autozooecia (macrozooecia) of Trepostomata may have served for brooding of larvae (Ulrich 1890, Astrova 1965). In contrast to the Cyclostomata, representatives of Palaeostomata reveal various internal morphological characters in form of lateral projections or chamber partitions. Functional morphology of these structures is often difficult to interpret as they are largely unknown among Recent stenolaemates. Autozooecial chambers in Palaeostomata are often portioned by horizontal structures called basal diaphragms (Boardman 2001). These structures seal off interiors of colonies from nutrition and further growth (Figs. 2.4 C and 2.5 B, D, F). Such structures are not known in modern cyclostomes and their interpretation remains difficult. However, they supposedly served as a “floor” of the living chambers of feeding zooids (Boardman 2001). Furthermore, some diaphragms in trepostome bryozoans are regarded as terminal, which capped living chambers. Some structures like the cap-like apparatus sensu Conti and Serpagli (1987), known from halloporine bryozoans, may also be a kind of terminal diaphragm (Fig. 2.3 I). In fenestrate bryozoans, centrally perforated terminal diaphragms are considered to represent secondary nanozooecia, polymorphic structures comparable to vibracula of cheilostomes (Bancroft 1986b). Various kinds of partitions of autozooecial chambers are mainly known in trepostome and cystoporate bryozoans, such as hemiphragms, ring-septa, cystiphragms (Fig. 2.5 K), funnel-shaped cystiphragms, and flask-shaped chambers (Fig. 2.12 C, D). The function of hemiphragms, ring-septa, and cystiphragms may be considered as portioning of the autozooecial chambers or possible attachment structure of retractor muscles (Boardman 1971), whereas the funnel-shaped cystiphragms and flaskshaped chambers are rather seen as polymorphs (Utgaard 1973, 1983, Boardman & McKinney 1976). Cryptostome, fenestrate, and some cystoporate bryozoans also have a kind of lateral structures, which are called hemisepta. These are shelflike, straight, or curved

16 

 2 Fossil record and evolution of Bryozoa

Fig. 2.3: (A–F) Indications of soft parts and fossilized brown deposits in autozooecial chambers of Palaeostomata (thin sections): A–C, Hemitrypa sp. (Fenestrata), Eifelian, Middle Devonian, Germany (A – tangential section, B, C – longitudinal sections); D, Cyclophaenopora robusta Spjeldnaes, 1984 (Cryptostomata, Rhabdomesina), oblique section showing encapsulated brow deposits (arrow). Vasalemma Formation, lower Katian, Upper Ordovician; Vasalemma Quarry, Estonia; E, F, Unidentified trepostome bryozoan, Pirgu Stage, Katian, Upper Ordovician; Vormsi island, Estonia. (G, H) Trepostome Leptotrypella provecta Boardman, 1960a, Emsian, Lower Devonian; Cantabrian Mountains, northwestern Spain. Tangential (G) and longitudinal (H) sections showing mural spines in autozooecial walls (arrows). (I) Cap-like apparatus, covering autozooecial apertures, Hallopora sp. (Trepostomata), Rakvere Stage, Katian, Upper Ordovician; Pechurki Quarry, Russia.



2.3 Development and evolution of bryozoans in the Palaeozoic 

 17

Fig. 2.4: Morphology of the Palaeostomata (Esthonioporata and Cystoporata). (A–D) Esthonioporate Revalotrypa gibbosa Bassler, 1911b. A, Longitudinal section of a fungi-shaped colony; B, C, longitudinal section showing autozooecial chambers and neozooecia; D, tangential section. Lower Ordovician, B III β, Kunda Stage; Putilovo Quarry at Volkhov, NW Russia. (E) Cystoporate Prismopora triangulata (White, 1878), branch transverse section. Horquilla Formation, Desmoinesian (late Moscovian), Pennsylvanian, Carboniferous; Cerros de Tule, Sonora, Mexico. (F) Cystoporate Sulcoretepora nitida (Ulrich, 1890). Oblique section of the branch. Tierra Blanca Member, Lake Valley Formation, Mississippian (Osagean); Sierra County, New Mexico, USA. (G) Cystoporate Cystodictya lineata Ulrich, 1884. Branch transverse section. Tierra Blanca Member, Lake Valley Formation, Mississippian (Osagean); Sierra County, New Mexico, USA. (H) Cystoporate Ramiporalia robusta Delvolve & McKinney, 1983, branch transverse section. Upper Viséan, Mississippian, Carboniferous; Roque Redonde, Montagne Noire, southern France. (I, J) Cystoporate Fistuliphragma eifelensis Ernst, 2008: I, Colony surface with a monticula; J, Autozooecial apertures with lunaria. Junkerberg Formation, Eifelian, Middle Devonian; Gondelsheim, Prüm Syncline, western Rhenish Massif, Germany.

18 

 2 Fossil record and evolution of Bryozoa



2.3 Development and evolution of bryozoans in the Palaeozoic 

projections that extend from the wall partway into autozooecial chambers (Karklins 1983). According to their position, hemisepta may be superior if they project from the proximal wall into the autozooecial chamber (Figs. 2.7 B and 2.12 I, J) or inferior if they originate from the mesotheca or distal wall (Figs. 2.6 K, L and 2.11 I). Lunaria are structures of a semicircular shape representing projections in form of hoods on the colony surface (Fig. 2.4 I, J). Lunaria originate at early ontogenetic stages of autozooecial development and are located at the proximal part of autozooecial chambers (Utgaard 1973, 1983). In tangential sections, lunaria are microstructurally different from autozooecial walls and commonly have a shorter radius of curvature (Fig. 2.13 L). In the majority of genera, lunaria are oriented toward the nearest macula (Fig. 2.4 I). Lunaria are an exceptional character of the Order Cystoporata. However, structures similar to lunaria were found in the Devonian rhabdomesine cryptostome Lunostoma Ernst et al., 2012c, which are apparently homeomorphic in origin. In post-Palaeozoic stenolaemates, similar structures are known in Jurassic and Recent lichenoporid bryozoans (Borg 1944, Utgaard 1968, Boardman 1984, Voigt 1993). Their origin is also most probably due to homeomorphy. Fenestrate and cryptostome bryozoans reveal a significant number of structures called allozooecia, amplozooecia, cavernozooecia, cyclozooecia, fossazooecia, leptozooecia, macrozooecia, metaxizooecia, minutozooecia, parazooecia, tectitozooecia, and aviculomorphs (Morozova 1973, 1974, 1987, 2001). In their majority, these are chamber- or tube-like structures (Fig. 2.7 F), which are often regarded as equivalents of avicularia and vibracularia of cheilostome bryozoans, or special kenozooids for improving stability (Morozova 2001). Some of them, like microzooecia, are thought to be of reproductive importance (Morozova 1973, 1974). Aviculomorphs are rounded chambers with triangular projections (Fig. 2.7 D) and occur in the Devonian genus Fenestrapora. They are regarded

 19

as analogous development to true avicularia of Cheilostomata (McKinney 1998, Morozova 2001). Notable is the genus Aviculofenestella Xia, 2002 with two species from the Bajocian (Middle Jurassic) of northern Tibet, which is characterized by the presence of aviculomorphs. The Jurassic age of this record appears to be doubtful; the age of the block from where this material was extracted may be most probably Middle Palaeozoic. Trepostome bryozoans are relatively poor in polymorphs. They have generally two types of tube- or prismatic-shaped heteromorphs called mesozooecia and exilazooecia (which are sometimes regarded being extrazooidal structures, see Boardman & Buttler 2005). These structures were called “unmature zooecia” by early authors, because of their smaller size compared to the autozooecia. The main difference is the presence of closely spaced diaphragms in mesozooecia (Fig. 2.5 B, D) and their absence in exilazooecia (Fig. 2.5 I). The presence of mesozooecia generally constitutes the Suborder Halloporina, while the Amplexoporina possess exilazooecia (Astrova 1965, 1978). The importance of both of these zooecia seems to be of structural character. It was shown that exilazooecia played a space-filling role in order to maintain regular spacing of autozooecia (Key et al. 2001; Fig. 2.12 E, F). Mesozooecia apparently had the same function (Astrova 1965, 1978, Boardman & Buttler 2005). Both mesozooecia and exilazooecia can give rise to new autozooecia during ontogenetic development. Esthonioporate bryozoans possess similar structures called “neozooecia” (Fig. 2.4 A–C). Autozooecial chambers in fossil bryozoans are known to bear so-called brown deposits, which are regarded as fossilized brown bodies remaining after degenerationregeneration cycles (e.g. Borg 1926, Gordon 1977, Boardman 1999, Ernst & Voigt 2002). Especially Early Palaeozoic trepostomes and cryptostomes are rich in brown deposits, which can indicate the position of soft-bodied structures (Fig. 2.3 A–F).

◂ Fig. 2.5: Morphology of the Palaeostomata (Trepostomata). (A–D) Parvohallopora ramosa (d’Orbigny, 1850): A, Branch transverse section; B, longitudinal section; C, tangential section; D, longitudinal section of a mesozooecium with closely spaced diaphragms. Furuberget Formation, Sandbian, Upper Ordovician; Mjøsa District, Norway. (E–G) Amplexopora crassiparietum Ernst & Nakrem, 2015: E, F, longitudinal section; G, tangential section. Steinsfjorden Formation, Brattstad Member, Sheinwoodian–Homerian, Wenlock, lower Silurian; Ødegårdsviken, Ringerike, Norway. (H, I) Dyscritella angularis (Trizna, 1948): H, tangential section showing autozooecial apertures, exilazooecia and acanthostyles; I, longitudinal section showing an autozooecial chamber with diaphragm and exilazooecium (arrow). Zechstein, Upper Permian; Gera, Germany. (J) Dyscritellina cf. aculeata Morozova in Morozova & Krutchinina, 1986, tangential section. Upper Permian; Canada. (K) Homotrypa niagarensis Ernst et al., 2019, longitudinal section showing cystiphragms. Reynales Formation, Hickory Corners Member, Aeronian, Llandovery, Lower Silurian; Hickory Corners, New York, USA. (L, M) Ulrichotrypa incrustata Ernst, 2001: L, tangential section; M, longitudinal section showing autozooecial wall with tubules. Zechstein, Upper Permian; Gera, Germany. (N, O) Microcampylus regularis Ernst, 2008: N, oblique section of the branch; O, tangential section showing tubules in autozooecial walls. Upper Nims Member, Junkerberg Formation, Eifelian, Middle Devonian; Brühlborn, Prüm Syncline, western Rhenish Massif, Germany.

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 2 Fossil record and evolution of Bryozoa

2.3 Development and evolution of bryozoans in the Palaeozoic 



2.3.1.3 Overview of the orders of the Palaeostomata Subdivision of Palaeostomata and phylogenetic relations within the clade is not consistently accepted at the current stage of research. There are two main views on the taxonomic division of Palaeozoic bryozoans, represented by Western and Soviet (now Russian) schools of bryozoology. A detailed history of the subdivision of the Palaeostomata is beyond the scope of the present review. The main differences concern the rank of some taxonomic units, for example the order rank of Rhabdomesina supported by Russian colleagues (suborder of the Cryptostomata according to Utgaard 1983) or the relation of Fenestellida (Fenestrata) to the Gymnolaemata (e.g. Ulrich 1890, Viskova & Morozova 1988, Gorjunova 1996, Morozova 2001, Xia 2002). At present, the following orders within the Superorder Palaeostomata are accepted: Esthonioporata, Cystoporata, Trepostomata, Fenestrata, and Cryptostomata (e.g. Ma et al. 2014, Taylor & Waeschenbach 2015). The group of Timanodictyida is regarded as suborder of the Order Cryptostomata (Blake 1983a). However, the position of the latter clade is controversial, Russian specialists consider both Cryptostomida and Timanodictyida as separate orders (e.g. Gorjunova & Lavrentjeva 1993, Gorjunova 1994, 1996). Esthonioporate bryozoans were usually considered to belong to Trepostomata or Cystoporata (e.g. Astrova 1978, Gorjunova 1996). Ma et al. (2014) erected the Order Esthonioporata, which they defined as follows: Hemispherical, massive and rarely encrusting colonies. Maculae common on colony surface. All mineralized walls above basal exterior colony walls are interior in origin. No differentiation recognized between endozone and exozone. Autozooecia long with polygonal cross-sections. Walls granular or with indistinct laminae. Diaphragms common or absent, thin, ring septa may develop. Styles often present, mesozooecia and exilazooecia absent, neozooecia present (Ordovician – Devonian).

The Order Cystoporata was erected by Astrova (1964), who extracted them from Cyclostomata. The following

 21

definition is modified after Utgaard (1983) and Gorjunova (1996): Hemispherical, massive, encrusting, branched, lenticulate (bifoliate), or reticulate colonies. Maculae common on colony surface. All mineralized walls above basal exterior colony walls are interior in origin. Differentiation between endozone and exozone distinct or absent. Autozooecia cylindrical or prismatic, often with polygonal cross-sections in endozones and circular apertures in exozones. Apertures often with lunaria. Walls granular or laminated. Diaphragms common or absent, thin, cystiphragms and hemiphragms may develop. Vesicular skeleton between autozooecia often developed. Styles often present, but mainly restricted to the vesicular skeleton. Communication pores present in some genera (ceramoporines). Apparent brooding structures known in few genera (Ordovician – Triassic).

The Order Trepostomata was erected by Ulrich (1882). The content of the order changed through time, and currently, the division in the suborders Amplexoporina and Halloporina is widely accepted (Astrova 1965). The current diagnosis is modified after Gorjunova (1996): Lamellar encrusting (mono- and multilamellar), hemispherical, massive, discoidal, branched ramose, and rarely lenticular (bifoliate). Maculae common on colony surface. All mineralized walls above basal exterior colony walls are interior in origin. Differentiation between endozone and exozone usually distinct. Autozooecia long or short, prismatic with polygonal cross-sections. Walls distinctly laminated, often irregularly thickened, in some genera containing spherules and tubules. Communication pores absent. Diaphragms common or absent, thin or thick; cystiphragms, ring septa, hemiphragms may develop. Styles often present, mesozooecia or exilazooecia often present. Apparent brooding structures unknown (Ordovician – Triassic).

The following orders were formerly summarized within the Order Cryptostomata Vine, 1884, which apparently had the most complex history of research. The earliest attempt of a division was made by McNair (1937), who suggested principal differentiation by colony form. Astrova and Morozova (1956) accordingly defined the Order Cryptostomata and established three suborders: Fenestelloidea, Ptilodictyoidea, and Rhabdomesoidea.

◂ Fig. 2.6: Morphology of the Palaeostomata (Cryptostomata). (A–C) Trigonodictya parvula Ernst & Carrera, 2012 (Ptilodictyina): A, longitudinal section; B, transverse section; C, tangential section of an aperture. Las Plantas Formation, Sandbian, Upper Ordovician; San Juan Province, Precordillera of Western Argentina. (D–F) Timanodictya sp. (Timanodictyina): D, transverse section; E, longitudinal section; F, tangential section. Toroweap Formation, Artinskian, Lower Permian; Nevada, USA. (G, H) Cyclophaenopora robusta Spjeldnaes, 1984: G, transverse section; H, tangential section. Vasalemma Formation, lower Katian, Upper Ordovician; Vasalemma Quarry, Estonia. (I) Moyerella parva Ernst et al., 2019 (Rhabdomesina), branch transverse section. Reynales Formation, Hickory Corners Member, Aeronian, Llandovery, lower Silurian; Hickory Corners, New York, USA. (J–M) Ascopora triseriata Schulga-Nesterenko, 1955 (Rhabdomesina): J, branch transverse section; K, L, longitudinal section showing axial zooecia and autozooecial chambers with hemisepta; M, tangential section showing autozooecial apertures, acanthostyles and paurostyles. Rod El Hamal Formation, upper Viséan-Westphalian, Pennsylvanian, Carboniferous; Wadi Araba area, Egypt.

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 2 Fossil record and evolution of Bryozoa

2.4 Evolutionary history of Bryozoa 



Elias and Condra (1957) constituted the Order Fenestrata (=Suborder Fenestelloidea Astrova & Morozova, 1956). Later, Viskova and Morozova (1988) raised this suborder to the rank of the Order Fenestellida. The Order Fenestrata is currently divided in two suborders: Phylloporinina and Fenestellina, following Lavrentjeva (1985). In contrast, Gorjunova (1996) and Morozova (2001) raised both suborders to orders Phylloporinida and Fenestellida within the Infraorder Fenestelloidea. The following diagnosis of the Order Fenestrata is modified, combining the diagnoses for Phylloporinida (Suborder Phylloporinina) and Fenestellida (Suborder Fenestellina) from Gorjunova (1996) and Morozova (2001): Arborescent unilaminate colonies of various shape: pinnate, freely dichotomizing (without connection), reticulate funnel- or fan-formed, reticulate screw-shaped, and rarely complex encrusting (Fig. 11G). Autozooecia usually short, box-shaped, with vestibules at proximal ends, in Phylloporinina long, tubular, often containing diaphragms (Fig. 7I). Hemisepta often present (Fig. 7B). Autozooecial apertures rounded to circular. Protective superstructures develop in some genera. Walls divided in internal granular and external laminated skeleton. Autozooecial apertures are usually arranged in rows and may be divided by longitudinal keels which can carry nodes. Heterozooecia include microzooecia, parazooecia, cavernozooecia, cyclozooecia, aviculomorphs, and metaxizooecia. Apparent brooding structures in form of spherical chambers attached to vestibules of autozooecial chambers common (Ordovician – Permian).

Shishova (1968) raised Rhabdomesoidea to the order rank (Order Rhabdomesida), and this opinion is still followed by Russian specialists (e.g. Gorjunova 1985, 1996). Later, Gorjunova (1987) raised Ptilodictyoidea to the rank of order (Order Cryptostomata). The main difference between ptilodictyines and rhabdomesines is their budding pattern. In ptilodictyines, zooecia bud from a mesotheca (or median wall), producing bifoliate branches (Figs. 2.6 A, B and 2.11 D, F), whereas in rhabdomesines, they originate from the more or less distinct median axis (e.g. Blake 1983a,

 23

Gorjunova 1985, Gorjunova & Lavrentjeva 1993; Figs. 2.6 G, I and 2.11 H, K). In some genera, the median part of the colony is composed of one or many heterozooids, like an axial tube (Fig. 2.12 H–J), or bundle of axial zooecia (Figs. 2.6 J, K, and 2.13 D, E). The Order Cryptostomata is currently divided into two suborders, Ptilodictyina and Rhabdomesina, whereas Blake (1983a) accepted the fenestrates and timanodictyines as the third and the fourth suborders of Cryptostomata. The modified diagnosis of the Order Cryptostomata Vine, 1884 in its currently accepted composition (Ptilodictyina and Rhabdomesina; consider Boardman 1983) is compiled after Blake (1983a), Gorjunova and Lavrentjeva (1993) and Gorjunova (1996): Arborescent colonies of various shape: bush-like, or reticulate (usually anastomosed). Branch transverse section circular or lens-shaped; secondary overgrowths rare. Clear distinction between endozone and exozone. Apertures common on all branch surfaces, arranged in regular longitudinal or spiral rows; apertures elliptical, subcircular, rhombic or rectangular. Autozooecia generally short, rarely elongate, usually with zooecial bend at endozonal-exozonal boundary. Hemisepta and mural spines present in some taxa; interzooecial pores absent. Vesicular skeleton may be developed (Ptilodictyina). Polymorphs represented by metazooecia, mesozooecia, tectitozooecia, allozooecia and axial zooecia; distinct brooding structures unknown. Striae, ridges, styles and stylets commonly well-developed on colony surface (Ordovician – Permian; an apparent rhabdomesine cryptostome Tebitopora reported from the Middle Triassic).

2.4 Evolutionary history of Bryozoa 2.4.1 Ordovician Although Ordovician bryozoan faunas are widely known and intensively studied worldwide, the earliest events in

◂ Fig. 2.7: Morphology of the Palaeostomata (Fenestrata). (A) Penniretepora sp. (Fenestellina). Las Llacerias Formation, Kasimovian, Pennsylvanian; Asturias, Cantabrian Mountains, northwestern Spain. (B) Laxifenestella kondrovensis (Schulga-Nesterenko, 1955) (Fenestellina), mid-tangential section of autozooecial chambers with hemisepta (arrows). Upper Viséan, Mississippian, Carboniferous; Roque Redonde, Montagne Noire, southern France. (C) Protoretepora sp. Bruten Yard, Western Australia; Noonkanbah Formation, Kungurian, Lower Permian. (D) Fenestrapora transcaucasica Morozova & Lavrentjeva, 1998 (Fenestellina). Thin section of the reverse surface showing aviculomorphs (arrows). Upper Nims Member of the Junkerberg Formation, middle Eifelian, Middle Devonian; Brühlborn near Rommersheim, Germany; (E) Colony obverse surface showing autozooecial apertures and a reproductive heterozooecium (arrow). Müllert Subformation of the Ahbach Formation, lowermost Givetian, Middle Devonian; Üxheim-Ahütte, Germany. (F) Thamniscus perplexus Ernst in Lisitsyn & Ernst, 2004 (Fenestellina), obverse colony surface with cyclozooecia (arrows). Zechstein, Upper Permian; Beeckerwerth near Duisberg, Germany. (G) Acanthocladia anceps (Schlotheim, 1820) (Fenestellina), branch with reproductive heterozooecia (arrows). Zechstein, Upper Permian; Beeckerwerth near Duisberg, Germany. (H) Chasmatopora rossae Ernst & Carrera, 2012 (Phylloporinina). Las Plantas Formation, Sandbian, Upper Ordovician; San Juan Province, Precordillera of Western Argentina. (I) Pseudohornera surculosa Lavrentjeva, 1985 (Phylloporinina). Sandbian, Kukruse stage (Kivioli Member), Upper Ordovician; Kiviõli, Estonia.

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 2 Fossil record and evolution of Bryozoa

bryozoan evolution are still unclear (e.g. Ross 1985, Taylor & Larwood 1990, Taylor & Ernst 2004, Ernst 2018). The earliest calcified bryozoans were apparently morphologically simple corynotrypids (Fig. 2.9 D), which originated from an unmineralized bryozoan (apparently ctenostome-like ancestor) with a simple, uniserial, encrusting colony form (Taylor & Larwood 1990, Gorjunova 1992, Taylor & Ernst 2004, Ma et al. 2015, Taylor & Waeschenbach 2015). Cyclostome bryozoans are considered as basal stenolaemates that mainly form uniserial encrusting colonies during the Ordovician, with the earliest record from the Dapingian of Russia (Gorjunovia Taylor & Rozhnov, 1996). Some more complex forms possessing pseudopores are known from the Ordovician and Silurian (Fig. 2.9 C, E). However, the oldest known bryozoans belong to the Palaeostomata, and they already show relatively advanced skeletal morphology (e.g. Taylor & Curry 1985, Hu & Spjeldnaes 1991, Cuffey et al. 2012). The oldest known bryozoan is the cryptostome Prophyllodictya simplex Ma et al., 2015 from the lower Nantzinkuan Formation, lower Tremadoc, which developed ramose (branching) colonies. Slightly younger faunas of the Fenxian Formation of South China contain additionally encrusting to dome-shaped Nekhorosheviella (Esthonioporata) and branched Orbiramus (Trepostomata) (Xia et al. 2007, Adachi et al. 2012). Early bryozoan faunas represented mainly by esthonioporines and trepostomes are known from the Floian of Baltoskandia (e.g. Koromyslova 2011, Fedorov et al. 2017). Apparently, bryozoans had a long period of earlier evolution that left no fossil evidence (e.g. Taylor & Waeschenbach 2015). Boring ctenostomes (Gymnolaemata) are known from the Early Ordovician, too (Mayoral 1991, Mayoral et al. 1994, Taylor & Rozhnov 1996). The genera Ropalonaria Ulrich, 1879 and Vinella Ulrich, 1890 were reported from the Late Ordovician of USA. A single example of a soft-bodied ctenostome preserved by bioimmuration is known from the Early Ordovician of Great Britain and Czech Republic (Fig. 2.9 A, B). The species Bolopora undosa Lewis, 1926 has been described from the basal Arenig (Early Ordovician) of the Ffestiniog district, North Wales. Initially, this fossil has been regarded as a cyclostome bryozoan. The same fossil was described by Prantl (1939) as the cyclostome bryozoan Berenicea vetera Prantl, 1939 from the Skiddavian (=Arenigian) Stage of Bohemia. Ross (1964) and then Hofmann (1975) claimed this fossil (Bolopora undosa) to be an alga. Study of the type material of Bolopora undosa, deposited at the Sedwick Museum in Cambridge, as well as material on Berenicea vetera Prantl, 1939 from the Klabava Formation (Early Ordovician, Floian), from Ejpovice in Czech Republic (provided by Michal Mergl, Prague),

showed that this fossil is rather a soft-bodied ctenostome bryozoan preserved by bioimmuration in phosphatic oncoids (Taylor & Ernst 2006). The genus Schallreuterella Hillmer, 1987 from the Late Ordovician (Hirnantian) of Sweden bears some superficial similarities to cheilostome bryozoans, especially in the shape of the aperture with proposed hinge line for an operculum and the shape of the autozooecia. These morphological features lead to the assumption of a huge gap in the fossil record between the Ordovician and Jurassic, the time of confirmed origination of cheilostomes (Hillmer 1991). In fact, Schallreuterella appears rather related to the Suborder Fenestellina (Order Fenestrata), a group that has often been regarded being ancestral to Cheilostomata because of their box-like zooecia and avicularia-like heteromorphs, as well as occasional findings of opercular structures (e.g. Morozova 1973, 1974, 1987, Wyse Jackson & Bancroft 1994). The alleged similarities between fenestrates and cheilostomes are due to evolutionary convergence. The majority of bryozoan faunas of the Ordovician occurred in tropical or subtropical climatic zones, whereas some others existed in temperate environments such as shelves of Gondwana. The sea level was high during most of the Ordovician, with extensive epicontinental seas, especially in the tropics (e.g. Haq & Schutter 2008, Pratt & Holmden 2008), and the climate was predominantly warm with high levels of CO2 (“green-house”), followed by a rapid cooling at the end of this period (Hirnantian glaciation). Bryozoans diversified rapidly during the Ordovician (Fig. 2.8). Within a short time interval, all orders except the group of Timanodictyina were established. Bryozoan diversity increased in several waves from the Tremadoc to the early Katian, with the peak in the late Sandbian (Taylor & Ernst 2004, Ernst 2018). This diversification event coincided with the beginning of the global flooding (e.g. Ross & Ross 1992, 1996). The reasons for the biodiversification of benthic suspension feeders, such as bryozoans, brachiopods, or echinoderms, are often explained by increasing phytoplankton availability (e.g. Servais et al. 2008, 2010, 2016). The level of bryozoan generic endemism was high during the Early and Middle Ordovician, whereas the bryozoan faunas in the Late Ordovician (Katian) became more cosmopolitan (Tuckey 1990a, Taylor & Ernst 2004). This loss of endemism is explained by the global warming in the late Katian (Boda Event), which enabled intermixing of temperate and tropical faunas (Jiménez-Sánchez & Villas 2010). The Late Ordovician was signed by a series of extinctions among bryozoans, the first and the strongest



2.4 Evolutionary history of Bryozoa 

 25

Fig. 2.8: Diversity dynamics of the Palaeostomata. (A) Generic diversity (raw data including Lazarus genera and singletons). (B) Generic originations (blue) and extinctions (red). Data set was compiled in January 2019 on the temporal distribution of 581 genera of Palaeostomata.

of them happened in the early Katian (Tuckey & Anstey 1992, Ernst 2018). The second extinction occurred in the late Katian, apparently caused by the Boda Event. Both first extinctions were interrupted by a slight recovery in bryozoan diversity. The third and the weakest extinction coincided with the global cooling in the Hirnantian. The diversity dynamic shows not only high extinctions rates, but also low rates of originations of new genera (Ernst 2018). Ordovician bryozoan faunas are dominated by trepostomes, cystoporates and ptilodictyine cryptostomes, whereas fenestrates and rhabdomesine cryptostomes are less diverse. Esthonioporine and trepostome bryozoans produced a variety of growth forms from which massive hemispherical, or dome-shaped, ones (Figs. 2.4 A, 2.9 G, and 2.10 I) are unmistakably characteristic for the Early

Palaeozoic (e.g. Taylor & James 2013). Dome-shaped trepostomes from Baltica reached significant sizes (Fig. 2.9 G). Some species of the esthonioporine genus Dianulites developed horn-shaped colonies (Fig. 2.9 H). Ptilodictyine cryptostomes developed exclusively ramose bilaminate colonies, which were lancet-shaped, bifurcating, or reticulate (Figs. 2.6 A–C and 2.9 J). In these colonies, autozooecia budded from a median lamina called mesotheca. This group of bryozoans declined significantly during the Silurian and much more during the Devonian (Fig. 2.11 D–F). In the Carboniferous, some few holdovers of ptilodictyines are known, and the last species of the suborder is known from the Early Permian of Canada (Ernst & Nakrem 2007). Fenestrate bryozoans of the Ordovician are mainly represented by the Suborder Phylloporinina, with only

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few species of the Suborder Fenestellina known from the Late Ordovician. Phylloporines developed typical fenestrate growth forms: branched dichotomous, pinnate, and reticulate (Fig. 2.7 H, I). A well-known example for the diverse phylloporine fauna is the so-called kukersite (e.g. Bekker 1921), a light-bright marine oil shale of the Sandbian age, which occurs mainly in Estonia and North West Russia (Fig. 2.9 I). The majority of the esthonioporates disappeared at the end of the period.

2.4.2 Silurian The Silurian has been characterized as a period of low provinciality for marine invertebrates, with little biogeographic differentiation being observed in faunas from major continents. This situation was caused by the closing of the Iapetus Ocean, which ceased to be a barrier to larvae of marine animals (Cocks & Fortey 1982, Cocks 2001). This had consequences for animal diversity. Tuckey (1990b) studied the global bryozoan palaeobiogeography in the Silurian. He stated a low level of endemism for bryozoans during this period and postulated a correlation between provinciality and bryozoan diversity. Controversially, McCoy and Anstey (2010) came to the result that bryozoans are characterized by a high level of endemism in the Silurian, much higher than other groups of organisms. Buttler et al. (2013) showed that the provinciality was heterogeneous during the Silurian. During the Llandovery, bryozoans exhibited distinct provincialism, but this declined during the Wenlock, only to reemerge during the Ludlow. Late Silurian (Přidoli) faunas show a possible division into two provinces. Bryozoan diversity was relatively low during the Silurian, showing little recovery during the Wenlockian. The extinction of bryozoans at the end of the Silurian is evident and corresponds to the Mid-Přidolian bioevent. The causes of this event are supposed to be a change of the sedimentation regime and diminishment of suitable habitats (e.g. Kaljo et al. 1996, Calner 2008). Among

2.4 Evolutionary history of Bryozoa 

 27

the bryozoan groups, ptilodictyine cryptostomes experienced the highest decline. The phylloporines were largely replaced by fenestellines, which became much more diverse and abundant during the Silurian. Trepostome bryozoans suffered also significant losses during the Early Silurian, with slight recovery in the Wenlockian. The majority of Ordovician dome-shaped taxa disappeared (Fig. 2.10 I); the trepostomes in the Silurian are represented by branched ramose and encrusting species (Fig. 2.10 B–C, H). In contrast, the cystoporate diversity increased in the Silurian compared to the Ordovician. They produced mainly encrusting, globular, or branched ramose colony forms. Although not as abundant and diverse as in the Ordovician, bryozoans were important elements of the benthic communities during the Silurian and were locally rock-forming (Fig. 2.10 A–C, E, H). Well-studied Silurian bryozoan faunas are known from North America and Eurasia.

2.4.3 Devonian During the Devonian, several bioevents of smaller scale occurred (e.g. Walliser 1996). Bryozoans showed a slightly different sensitivity to these events than other fossil groups (Cuffey & McKinney 1979, Bigey 1988, Morozova et al. 2002). The Choteč Event at the Emsian–Eifelian boundary strongly affected brachiopods and trilobites, whereas corals and bryozoans did not show any noticeable changes in their diversity (Chlupáč & Kukal 1986, 1988, Ernst et al. 2012b, Ernst 2013). The bryozoan diversity increased steadily during the Early Devonian and reached its maximum in the Givetian (Fig. 2.8). In the subsequent Frasnian stage, the drop in bryozoan diversity is dramatic (Horowitz et al. 1996, Taylor & Larwood 1988). The Givetian/Frasnian bryozoan extinction most probably corresponds to the late Givetian Taghanic bioevents (e.g. House 2002, Boucot 1990). This crisis represents a series of faunal changes, with the impact on different groups shifted in time (e.g.

◂ Fig. 2.9: Ordovician bryozoans. (A) Apparent ctenostome Bolopora undosa Lewis, 1926 (=Berenicea vetera Prantl, 1939). Arenig, Lower

Ordovician; Ejpovice, Czech Republic (photo by Paul D. Taylor, London). (B) Same as in A, SEM photograph showing shallow zooids. (C) Cyclostome Kukersella borealis (Bassler, 1911b). Rakviere Stage, Upper Ordovician; Rakvere, Estonia. (D) Cyclostome Corynotrypa sp. Bromide Formation, Sandbian; Arbuckle Mountains, Oklahoma, USA (photo by Paul D. Taylor, London). (E) Cystoporate Ceramopora sp. Pin Formation, Upper Ordovician; Spity Valley, India (large communication pores inside of autozooecia). (F) Cyclostome Cuffeyella sp., Lorraine Group, Ordovician; Cincinnati, Ohio (photo courtesy Paul D. Taylor). (G) Trepostome Diplotrypa sp. Rakvere Stage, Upper Ordovician; Pechurki Quarry, NW Russia (polished slab). (H) Esthonioporate Dianulites detritus Eichwald, 1829. Kunda Stage, Darriwilian, Lower Ordovician; west side of Volkhov River, NW Russia. (I) Chasmatopora papillosa (Bekker, 1921) (Fenestrata, Phylloporinina). Sandbian, Kukruse stage (Kivioli Member), Upper Ordovician; Kohtla quarry, Estonia. (J) Stictoporellina sp. (Cryptostomata, Ptilodictyina). Oandu Stage, Upper Ordovician; quarry Slantsy, NW Russia.

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Baird & Brett 2008). The Taghanic event was considerable not only for bryozoans but also for other groups such as ammonoids (House 1996), trilobites (Chlupáč 1994) and corals (Oliver & Pedder 1994). It was apparently caused by transgression and marked the end of faunal provincialism, which had persisted since the Early Devonian (Johnson 1970). This may also explain its heavy impact on bryozoans because they seem to be sensitive to changes in provincialism, as shown for the Ordovician (Taylor & Ernst 2002, Tuckey 1990a). The increase of cosmopolitanism and reduction of available habitats as a result of the Taghanic transgression (with subsequent deepening of carbonate platforms) seem to bias bryozoan generic diversity. In contrast to the Taghanic event, the Frasnian/ Famennian bioevent did not greatly affect bryozoans (e.g. Bigey 1988, Morozova et al. 2002). Indeed, there are even signs of a slight recovery of bryozoan faunas during the early Famennian (Ernst 2013). Different causes for the Frasnian/Famennian bioevent (the so-called Kellwasser event) have been discussed, ranging from a worldwide anoxia (e.g. McGhee 1996, Schindler 1990, Walliser 1996) to an asteroid shower (Sandberg et al. 2002). The Hangenberg bioevent at the Devonian/Carboniferous boundary was apparently caused by glaciation resulting in a strong drop in sea level (e.g. Caplan & Bustin 1999, Kaiser et al. 2006, 2008). This resulted in a significant shift in the composition of many animal groups including bryozoans (Horowitz & Pachut 1993, Gutak et al. 2008, Tolokonnikova & Ernst 2010, Ernst 2013). An important factor affecting bryozoan diversity in the Late Devonian was strong endemism of some faunas (Tolokonnikova & Ernst 2010). Remarkably, many Devonian bryozoans, especially fenestrates, developed structures of obvious defensive character (e.g. McKinney et al. 2003). This morphological trend may reflect increasing pressure of predators during the mid-Palaeozoic predator revolution (e.g. Signor & Brett 1984, Sallan et al. 2011). Devonian bryozoan-bearing sediments are widely distributed worldwide and contain amazing fossils (Fig. 2.11

2.4 Evolutionary history of Bryozoa 

 29

A–C, G). This was a time for restructuring of the bryozoan fauna. During this interval, trepostomes and cystoporates lost their dominance, whereas fenestrates (Figs. 2.7 D, E and 2.11 B, G) and rhabdomesine cryptostomes (Fig. 2.11 H–K) became a dominant group (Cuffey & McKinney 1979, Bigey 1988, Ernst 2013). This constellation largely influenced the composition of bryozoan faunas in the Late Palaeozoic.

2.4.4 Carboniferous In the Mississippian, the phylum Bryozoa underwent rapid diversification and phylogenetic differentiation but showed a gradual decline in the Pennsylvanian (Ross 1981, Bancroft 1987). The Tournaisian-Viséan rise in bryozoan diversity was the last significant diversification event among the Palaeostomata (Tolokonnikova et al. 2014; Fig. 2.8). Brachiopods (e.g. Shen et al. 2006) and corals (Wang et al. 2006) show similar diversity patterns usually explained by fluctuations of sea level (Ross & Ross 1996), which was high in the Mississippian and low in the Pennsylvanian. The rapid diversification of bryozoans during the Viséan can be explained by filling of niches emptied by the Hangenberg extinction and the lowering of predator stress (Brett & Walker 2002, Sallan et al. 2011). The influence of the reduced predation on bryozoan diversification is suggested by a strong decrease of protective morphologies in Carboniferous bryozoans. Two major extinction events were identified during the Carboniferous: Mid-Tournaisian Event and Mid-Carboniferous Event (Walliser 1996). The Mid-Tournaisian Event has been connected with the worldwide occurrence of anoxic facies as a consequence of a rapid transgression (e.g. Becker 1993). On the contrary, the Mid-Carboniferous Event coincides with a strong regression documented by unconformities (Ramsbottom 1977). It is not known in detail how bryozoans reacted to these events, but generally a drop in diversity during Serpukhovian and Bashkirian is postulated (Ross & Ross 1996). The decline of bryozoan diversity

◂ Fig. 2.10: Silurian bryozoans. (A) Surface of bryozoan-rich limestone. Reynales Formation, Hickory Corners Member, Aeronian, Llandovery, Lower Silurian; Hickory Corners, New York, USA. (B, C) Trepostome bryozoans (Amplexopora crassiparietum Ernst & Nakrem, 2015) in the Steinsfjorden Formation, Brattstad Member, Silurian, Wenlock, Sheinwoodian-Homerian Ødegårdsviken, Ringerike, Norway (B, rock surface; C, thin section). (D) Boring ctenostome Rhopalonaria attenuata Ulrich & Bassler, 1904. Klinteberg beds, Wenlock, lower Silurian. Gothem Hammer 3, Gotland, Sweden. (E) Surface of bryozoan-rich limestone. Rochester Shale, Wenlock, lower Silurian; Niagara Street, New York, USA. (F) Bryoliths (pen 145 mm long). Wenlock, lower Silurian; Nors stenbrott (Blå Lagunan), Gotland, Sweden. (G) Thin section of the bryolith showing bryozoan-microbial crusts. (H) Underside of a stromatoporoid encrusted by cystoporate and trepostome bryozoans. Wenlock, lower Silurian; Ireviken, Gotland, Sweden. (I) Trepostome bryozoan Monotrypa sp. Polished slab of a hemispheric colony. Ninase Formation, Jaani Stage, Wenlock, lower Silurian; Kuriku, island Saarema, Estonia. (J) Phylloporina asperatostriata Hall, 1852 (Fenestrata, Phylloporinina). Rochester Shale, Wenlock, lower Silurian; Niagara River, New York, USA. (K) Phaenopora multifida (Hall, 1883) (Cryptostomata, Ptilodictyina). Reynales Formation, Hickory Corners Member, Aeronian, Llandovery, lower Silurian; Hickory Corners, New York, USA.

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may be also explained by worldwide changes in the availability of shelf habitats preferred by bryozoans (Taylor & Allison 1998). In the Pennsylvanian, the bryozoan diversity remains low, due to lower origination rates and raising extinctions. Significant influence of the climatic changes combined with the so-called Late Palaeozoic Ice Age is considered as rather negative effect of cooling on bryozoan faunas (e.g. Ross 1981). This glaciation event occurred in an interval from late Mississippian through Early Permian time (326–267 Ma), during which continental glaciers were widespread throughout the Southern Hemisphere (e.g. Frakes et al. 1992, Bishop et al. 2009). Among the groups of Palaeostomata, fenestrates were the most successful bryozoan group in the Carboniferous (Fig. 2.12 A, B, K–M), followed by rhabdomesine cryptostomes (Fig. 2.12 G–J) and cystoporates (Ross 1981, Bancroft 1987). In contrast, trepostomes were significantly reduced after the mid-Carboniferous crisis.

2.4.5 Permian Permian bryozoans display a high diversity worldwide (Figs. 2.5 H–J, L, M, 2.6 D–F, 2.7 C, F, G, and 2.13 A–M). Well-studied faunas are known from North America, Eurasia, and Australia (Ross 1995b, Gilmour & Morozova 1999, and references herein). The earliest reports on Permian bryozoans from Europe are known from the 19th century (e.g. Schlotheim 1820, Geinitz 1848, 1861, King 1850). Permian bryozoan faunas show remarkable inhomogeneity caused most probably by climatic zonation and progressive isolation of marine basins (Gilmour & Morozova 1999). Differences in the distribution and composition of faunas in tropical and extra-tropical basins were significant. Reid and James (2008, 2010) compared the bryozoan diversity of Gondwana and northern Eurasia. Boreal faunas show a higher overall diversity than those of East Gondwana (106 vs. 61 species from East Gondwana). Some Permian bryozoans developed gigantic colonies, as for example the cystoporate Evactinostella (Fig. 2.13 B) from the Callytharra Fm. of Western Australia with a

2.4 Evolutionary history of Bryozoa 

 31

branch thickness of 4 cm and height of 25 cm, or the trepostome Tabulipora from the Kim Fjelde Fm. in Greenland, which has a branch thickness of 7 cm and colony height of 20 cm (Ross & Ross 1962, Håkansson & Madsen 1991, Key et al. 2005). Remarkably, some other groups of suspension feeders such as bivalves and brachiopods also developed gigantic taxa during the Permian (Hayasaka & Hayasaka 1953, Isozaki & Aljinović 2009). However, the gigantism of Permian bivalves is explained by endosymbiotic algae (Isozaki 2006), whereas the isotopic characteristic of bryozoan skeletons revealed no signs of algal symbiosis (Key et al. 2005), supporting the hypothesis that the gigantism of the Permian bryozoans occurred due to ideal growth conditions in an eutrophic environment. Early Permian climate was influenced by the final stage of the Late Palaeozoic Ice Age. After a series of cooling events, a rapid warming started in the late Artinskian, accompanied by extensive marine flooding (e.g. Haig et al. 2017). This climatic upturn and creation of new habitats allowed wide distribution and diversification of bryozoan faunas in the Permian time (Fig. 2.8). In contrast, the Late Permian is better known for its devastating extinctions. Bryozoans experienced a series of massive declines just passing the narrow bottleneck at the end of the period. Two events were most substantial for bryozoans: Guadalupian-Lopingian and end-Permian extinction events. Both events are regarded almost equally in their impact on life on Earth (e.g. Jin et al. 1994, Stanley & Yang 1994). The Guadalupian-Lopingian (or end-Guadalupian) extinction is often regarded as an effect of global cooling (e.g. Hayasaka & Hayasaka 1953, Isozaki & Aljinović 2009). Significant declines were stated also for tabulate and rugose corals, fusulinids, and brachiopods (Rong & Shen 2002, Ross 1995a). All bryozoan groups experienced a catastrophic drop in diversity, especially fenestrates. Less than two dozens of bryozoan genera survived into the Wuchiapingian. The causes of end-Permian extinction may be diverse and complex, including climatic/ecological effects, or reduction of biotopes due to the completion of Pangea

◂ Fig. 2.11: Devonian bryozoans. (A) Surface of bryozoan-rich limestone. Birdsong Shale, Lochkovian, Lower Devonian; Tennessee, USA. (B) Fenestrate Fenestrapora tuberculata Ernst, 2016a. Müllert Subformation of the Ahbach Formation, lowermost Givetian, Middle Devonian; Üxheim-Ahütte, Germany. (C–F) Intrapora variabilis Ernst, 2008 (Cryptostomata, Ptilodictyina): C, colony on the limestone surface; D, branch transverse section; E, tangential section; F, longitudinal section. Junkerberg Formation, Eifelian, Middle Devonian; Blankenheim, western Rhenish Massif, Germany. (G) Fenestrate Schischcatella heinorum Ernst & Bohatý, 2009. Lower Givetian, Middle Devonian; Gerolstein Syncline, Eifel, western Rhenish Massif, Germany. (H–K) Vidronovella elegantula Ernst, 2012a (Cryptostomata, Rhabdomesina): H, I, longitudinal section showing autozooecia with hemisepta (arrow); J, tangential section showing autozooecial apertures and paurostyles; K, branch transverse section. Lebanza Formation, Pragian, Lower Devonian; Arauz Sur (Arroyo section), Province of Palencia, Cantabrian Mountains, northwestern Spain.

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(e.g. Erwin 1994, 1996, Gilmour & Morozova 1999), bolide impact (Becker et al. 2001), volcanism related to the Siberian Trap (Renne & Basu 1991, Bowring et al. 1998), global warming (Joachimski et al. 2012), and global anoxia associated with sea-level rise (e.g. Wignall & Hallam 1992, Isozaki 1997, Hallam & Wignall 1999). Powers and Bottjer (2009a,b) investigated the consequences of environmental stress on bryozoan faunas during the extinction events in the Permian and Triassic. They found that bryozoans largely disappeared from deep-water offshore settings during the Late Permian and from nearshore and offshore settings during the Late Triassic. The recolonization of these habitats was delayed after each mass extinction event. Marine environmental instability during the Late Permian and Late Triassic apparently resulted from some stressful deep-water phenomenon, whereas extinctions in the nearshore environments in the Late Permian may be linked to atmospheric perturbations. All stenolaemate taxa except for three trepostome genera got extinct during the end-Permian bioevent (Schäfer 1994, Gilmour et al. 1998). Unfortunately, the fossil record at the Permian-Triassic boundary is very scarce and does not allow exact reconstruction of the faunal transition into the Triassic. Only two trepostome genera, Dyscritella and Pseudobastomella, were found in the latest Permian stage, Changhsingian, whereas the last representatives of Cystoporata, Fenestrata, and Cryptostomata got extinct in the preceding Wuchiapingian. The reported finding of a fenestrate bryozoan Aviculofenestella Xia, 2002 in the Middle Jurassic of northern Tibet may have derived from the block of an allochtonous Palaeozoic rocks inside of the Jurassic sediments. The presence of apparent cystoporate and cryptostome genera in the Triassic suggests some Lazarus taxa, which were not delivered by the fossil record.

2.4.6 Triassic Most of the Palaeozoic stenolaemate orders were extinguished during or at the end of the Permian. The exception

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 33

was the Order Trepostomata, which reoccurred in the earliest Triassic and within a time span of over 50 million years radiated again to evolve a substantial number of species (Fig. 2.14 A–C). Early Triassic bryozoan faunas were restricted to Boreal areas (Powers & Pachut 2008), locally even being rock-forming (Baud et al. 2008). During the Triassic, bryozoans diversified and dispersed globally (e.g. Schäfer 1994). The youngest trepostomes are known from the Rhaetian, the group finally becoming extinct by the end of the Triassic. In total, 22–25 bryozoan genera, depending on taxonomic opinion, were recognized from the Triassic. Only three genera of the few holdovers from the Permian show continued fossil record; others represent apparent Lazarus taxa. Thirteen trepostome genera are known from the Triassic up to date. Only three Triassic palaeostomate bryozoans are regarded being non-trepostome: Tebitopora Hu, 1984 from the Ladinian-Carnian of China (rhabdomesine cryptostome), cystoporates Cystitrypa Schäfer & Fois-Erikson, 1987 from the Carnian of Italy, and Cyclotrypa sp. from the Norian of Iran (Schäfer et al. 2003). After this slight recovery, bryozoans experienced a decline (mid-Carnian bioevent). In the Rhaetian, the last Palaeostomata got extinct (Schäfer & Fois 1986, Schäfer & Fois-Erickson 1987, Schäfer 1994, Powers & Pachut 2008). In contrast, cyclostome bryozoans seem to have survived the end-Triassic extinction, although they reveal a gap of ca 20 Ma in the fossil record, which raised questions about their phylogenetic relations (Ernst & Schäfer 2006). The only two genera of cyclostomes are known from the preceding Permian Period: Corynotrypa Bassler, 1911a (Fig. 2.9 D) and Lagenosypho Spandel, 1898, which are both simple forms lacking calcified interior walls (Langer 1980, Taylor 1980, 1985a). The earliest Triassic cyclostomes are known from the Late Triassic and are represented by several tubuliporine species (Schäfer 1994). Stomatopora cf. dichotomoides (d’Orbigny, 1850) and Reptomultisparsa hybensis (Prantl, 1938) occur in the Rhaetian of the Nizhne-Tatry Mountains, Czech Republic (Taylor & Michalik 1991). Morphologically, they distinctly belong to the post-Palaeozoic stenolaemate bryozoans,

◂ Fig. 2.12: Carboniferous bryozoans. (A) Fenestrate Ptylopora laticarinata Kaisin, 1942. Mississippian; Belgium. (B) Surface of bryozoan-

rich limestone with coils of Archimedes and diverse fenestrate bryozoans. Upper part of the middle Mississippian; Sloan Valley, Kentucky, USA. (C, D) Trepostome Tabulipora sp., tangential (C) and longitudinal (D) sections showing autozooecial chambers with ring septa. Mississippian; Algeria. (E, F) Trepostome Hinaclema sakagamii Schastlivtseva, 1991: E, tangential section; F, longitudinal section. Mobarak Formation, Viséan, Mississippian; northern Iran. (G–J) Rhabdomeson progracile Wyse Jackson & Bancroft, 1995 (Cryptostomata, Rhabdomesina): G, tangential section; H, transverse section showing axial zooecium; I, J, longitudinal section showing axial zooecium and hemisepta (arrows). Upper Viséan, Mississippian, Carboniferous; Roque Redonde, Montagne Noire, southern France. (K) Fenestrate Polypora remota Condra, 1902. Valdeteja Formation, Bashkirian, Pennsylvanian; Asturias, Cantabrian Mountains, northwestern Spain. (L, M) Fenestrate Polypora dendroides M‘Coy, 1844: L, tangential section; M, longitudinal section. Linoproductus bed, Viséan, Mississippian, Carboniferous; Hook Head, Ireland.

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having a fixed-walled skeletal organization, whereas Reptomultisparsa hybensis possess the earliest known brood chambers within cyclostomes. A few other more or less distinct tubuliporines were recorded from the Carnian of Italy (Bizzarini & Braga 1985) and the Norian of Central Iran (Schäfer et al. 2003). Some few doubtful records of cerioporines are known from the Late Triassic (Carnian) of Hungary (Papp 1901, Vinassa de Regny 1901). However, these fossils are doubtful, whereas confirmed cerioporines are known from the Middle Jurassic (Aalenian) of England (Paul Taylor, pers. com., 2018).

2.4.7 Jurassic The Jurassic has marked an important stage in the development of bryozoans. In the Early Jurassic, bryozoan diversity remained low but peaked greatly in the Bathonian (e.g. Walter 1969, Henderson & Perry 1981, Taylor & Sequeiros 1982, Taylor & Ernst 2008). Key novelties such as brooding care (gonozooecia), free-walled skeletal organization, and development of communication pores (Taylor & Larwood 1990, Schäfer 1991) were considered as causes for this radiation. The absolute majority of Jurassic bryozoan localities lie within the European continent, whereas a few occurrences outside Europe are known, including those from North America and South America, Africa Australasia, and the Middle East (see overview in Taylor & Ernst 2008). Jurassic bryozoan-rich localities are generally associated with non-tropical areas, and the tropical bryozoan faunas are extremely rare (Taylor 1994, Wilson et al. 2015). The Middle Jurassic was a time of diversification and innovation in the phylum Bryozoa, especially among the cyclostomes (Taylor & Ernst 2008). Most species were encrusters of different configurations (up to 73%), whereas rare erect forms are known mostly from the Middle

2.4 Evolutionary history of Bryozoa 

 35

Jurassic. It is remarkable that reticulate (reteporiform) colonies, which were very abundant in the Palaeozoic and later in the Cenozoic, are completely unknown in the Jurassic. Within the European region, bryozoans were widely distributed and achieved high assemblage richness (up to 33 species pro assemblage in the Bathonian of Normandy). The majority of Jurassic stenolaemates belong to the suborders Tubuliporina and Cerioporina of the Order Cyclostomata (Fig. 2.14 D). The earliest representative of the Suborder Rectangulata appeared in the Middle Jurassic (Bajocian or Bathonian; Paul Taylor, pers. comm., 2018). In the Late Jurassic, a new group of bryozoans appeared: the Order Cheilostomata. Two earliest cheilostomes are known: Pyriporopsis portlandendis Pohowsky, 1973 from the Tithonian of England and P. pohowskyi Taylor, 1994 from Oxfordian or Kimmeridgian of Yemen. The ancestry of cheilostomes from soft-bodied ctenostomes (probably arachnidioidean forms) is widely accepted (Banta 1976, Taylor 1986). Soft-bodied Jurassic ctenostomes are known from bioimmurations by oysters (Voigt 1977, Taylor 1978, 1990a,b, Todd 1994). A contrasting hypothesis suggested by Dzik (1975) considers cheilostomes being derived from simple tubuliporines. The Jurassic cheilostomes apparently had calcitic skeletons, whereas the majority of later taxa produced aragonitic or bimineralic skeletons (Taylor et al. 2009a).

2.4.8 Cretaceous The majority of Cretaceous bryozoan faunas are known from Europe and the USA associated with deposits of the western Tethys Ocean. Some few records of bryozoans exist from the margin of eastern Asia in the NW Pacific (e.g. Dick et al. 2018), South America (Taylor et al. 2009b),

◂ Fig. 2.13: Permian bryozoans. (A) Surface of bryozoan-rich limestone with abundant fenestrates. Callytharra Formation, SakmarianArtinskian, Lower Permian; Bidgemia, Western Australia. (B) Cystoporate Evactinostella crucialis (Hudleston, 1883). Callytharra Formation, Sakmarian-Artinskian, Lower Permian; Dead Man’s Galley, Western Australia. (C–E) Streblascopora (Streblascopora) marmionensis (Bretnall, 1926) (Cryptostomata, Rhabdomesina): C, tangential section; D, longitudinal section; E, transverse section. Callytharra Formation, Sakmarian-Artinskian, Lower Permian; Jimba-Jimba, Western Australia. (F) Cystoporate Etherella tibetensis Ernst, 2016b. Zhongba Formation, Permian (upper Cisuralian-Guadalupian); southwestern Tibet. (G–I) Fenestrate Rectifenestella chapmani (Crockford, 1944): G, fragment of colony on the rock surface; H, I, tangential section showing autozooecial apertures and outline of autozooecial chambers in mid-tangential section. Callytharra Formation, Sakmarian-Artinskian, Lower Permian; Callytharra Springs, Western Australia. (J) Cystoporate Hexagonella hudlestonei Crockford, 1957. Callytharra Formation, Sakmarian-Artinskian, Lower Permian; Callytharra Springs, Western Australia. (K, L) Cystoporate Fistulipora elegantula Nikiforova, 1933; K, longitudinal section; L, tangential section showing autozooecial apertures with lunaria (arrows) and vesicular skeleton. Araxilevis-bed, Dzhulfian (= Wuchiapingian), Upper Permian; Abadeh, Iran. M. Cystoporate Goniocladia afghana (Termier & Termier, 1971). Chili Formation, Sakmarian-Artinskian, Lower Permian; Kalmard area, central Iran.

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Fig. 2.14: Post-Palaeozoic bryozoans. (A–C) Arcticopora lobatula (Schäfer & Grant-Mackie, 1994): A, longitudinal section; B, transverse section; C, tangential section. Otapirian, Rhaetian, Upper Triassic; New Zealand. (D) Cyclostome bryozoan on an oyster shell. Lower Kimmeridgian, Jurassic; Wierzbica Quarry near Radom, Poland. (E) Bryozoan-rich limestone (erratic boulder) of the Danian age, LimstenGeschiebe, Fischbeck near Hamburg (collection of the University of Hamburg, courtesy U. Kotthoff). (F) Modern cool-water carbonates, rich of bryozoans (e.g. Hornera sp.), bivalves, gastropods, and serpulids, depth of 100 m, Norwegian Shelf off Tromsø. (G) Modern cool-water carbonates, rich of bryozoans (mainly Cellaria sinuosa), balanids, gastropods, and bivalves, depth of 80 m, Trezen ar Skoden, Roscoff, Bretagne (samples courtesy P. Schäfer, Kiel).



New Zealand (Taylor & Gordon 2007), or Africa (e.g. Brood 1977, Taylor & Zaborski 2002). Tropical faunas in the Cretaceous are rare (Di Martino & Taylor 2013). Bryozoan faunas of North America are much less diverse than those in Europe (e.g. Taylor & Cuffey 1992, Taylor & McKinney 2006, Taylor 2008, McKinney & Taylor 2016). Possible explanations could be unfavorable environmental conditions like those in the Western Interior Seaway, an epicontinental sea within the North American continent (e.g. Taylor & Cuffey 1992, He et al. 2016, Eldrett et al. 2017) or prevailing surface currents that flowed away from North America and toward Europe preventing larval migration westwards (Barron & Peterson 1989, 1990). After the decrease of the cyclostome diversity in the Late Jurassic (apparently an artifact because of incomplete fossil record), a new pulse of diversification of cyclostomes occurred in the Early Cretaceous (Albian), which peaked in the Cenomanian and Maastrichtian (e.g. Taylor & Larwood 1990, Viskova 1992, Sepkoski et al. 2000). The cyclostome suborders Cancellata and Articulata appeared during the Cretaceous. The earliest representative of the Suborder Articulata, Crisidia inopinata Lagaaij, 1976, was recorded from the Maastrichtian of Netherlands. The earliest cancellate species apparently is Petalopora flavapetrensis Walter, 1993 from the Hauterivian of Switzerland. Cheilostome bryozoans remained at low diversity until the late Albian–early Cenomanian, when they began an explosive radiation that has continued to the present day (e.g. Taylor 1988, 2000b, Lidgard et al. 1993, Jablonski et al. 1997). This radiation apparently had intrinsic causes, especially the acquirement of larval brooding and origin of short-lived, non-feeding larvae. Expected consequences of this development are decreased gene flow within and between populations and increasing the likelihood of speciation, triggering the explosive radiation. The onset of the bryozoan radiation coincided with major sea-level rise, which provided extended habitat areas for bryozoans.

2.4.9 Crisis at the K/T boundary One of the most intriguing moments in Earth history, the mass extinction on the Cretaceous-Palaeogene (or Cretaceous-Tertiary, K-T), boundary is usually associated with the consequences of a meteorite impact (e.g. Alvarez et al. 1980, Robin et al. 1993), although different explanations such as volcanic activity of Deccan Traps are discussed (e.g. McLean 1985, Keller et al. 2009). Some authors were able to show that the extinction started long before

2.4 Evolutionary history of Bryozoa 

 37

the impact on the K-T boundary (e.g. Birkelund & Håkansson 1982). This event is best known to have wiped out the dinosaurs, but also most major benthic groups in marine habitats were subject to very substantial extinctions. Both groups of skeletonized bryozoans, cyclostomes and cheilostomes, were affected by the K-T crisis (e.g. Voigt 1985, Viskova 1992, Lidgard et al. 1993, MacLeod et al. 1997, Håkansson & Thomsen 1999, Sepkoski et al. 2000, McKinney & Taylor 2001, McKinney et al. 2001a). O’Dea et al. (2011) demonstrated that certain morphological features of cheilostome bryozoans (zooidal and colonial size and shape, avicularian density) indicate a time of increased environmental stability in the Danish Basin long before the end of Maastrichtian. However, the sediment layer of 20-cm thickness immediately prior to the K-T boundary is marked by strong ecological perturbations, as seen from the morphology of cheilostome bryozoans. A global phytoplankton crash is discussed as a profound aftermath of an extraterrestrial impact at the K-T boundary (e.g. Alvarez et al. 1980, Arthur et al. 1987), with obvious consequences for suspension feeders like bryozoans. Although McKinney et al. (1998) stated a distinct cyclostome spike in the early Danian, no significant evidence of such a phytoplankton crash could be found in bryozoans. Sogot et al. (2013, 2014) studied biogeographical and ecological patterns in bryozoans across the K-T boundary using zooidal size and proportion of runner-like colonies as proxies for the level of primary production. These authors were able to show that no significant changes occurred in the studied parameters of bryozoans at the K-T transition.

2.4.10 Cenozoic After the start of the cheilostome radiation in the Cretaceous, bryozoan diversity increased continuously (e.g. Taylor 2000b). Following the crisis at the K/T boundary, cheilostome diversity recovered by the Eocene (McKinney et al. 1998). In contrast, cyclostomes declined after the Campanian/Maastrichtian and remained at low diversity (e.g. Voigt 1985, Lidgard et al. 1993). An immense increase in cheilostome diversity and stagnancy in the evolution of cyclostomes mostly has intrinsic causes and relates to their previous evolutionary history (Jablonski et al. 1997). Cheilostomes developed several key novelties that gave them competitive advantages against cyclostome bryozoans (McKinney 1992, 1995, Lidgard et al. 1993, Jablonski et al. 1997, Sepkoski et al. 2000). Most important novelties were more rapid ontogenetic development of zooids along colony margins and generally larger zooids, which provided benefit in direct competition for space and food

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and were important contributors to the radiation of cheilostomes in the Cretaceous (McKinney 1992). Larger zooids imply larger lophophores and, consequently, more efficient feeding mechanisms (e.g. Winston 1977, 1978, McKinney & Jackson 1989, Okamura 1990). Cheilostomes with their larger lophophores grow faster, and they can feed on a wider range of food sources. In contrast, cyclostomes with smaller lophophores can feed only on smaller particles. Most diverse bryozoan faunas of the Cenozoic are known outside of the tropics (e.g. Taylor & Allison 1998, Taylor 2000b, Taylor & Di Martino 2014). However, recent studies show that bryozoan diversity in the tropics is significant, contrary to the ubiquitous opinion (e.g. Tilbrook et al. 2001, Taylor & Di Martino 2014, Di Martino et al. 2018). Several factors explain such an imbalance, especially taphonomic and monographic biases. Tropical bryozoan faunas have a larger proportion of small encrusting species, which have poor preservation potential and do not easily attract the attention of researchers. Bryozoans in the tropics are weakly calcified, in contrast to the robust erect species of non-tropical habitats. In addition, the tropical faunas are dominated by cheilostomes with largely aragonitic skeletons, which are easily destroyed by diagenesis (e.g. Smith et al. 2006, Taylor et al. 2009a). According to existing compilations (Horowitz & Pachut 1994, 1996), bryozoan diversity increased steadily from the Paleocene to the Pleistocene. The bryozoan diversity within assemblages also increased continuously (e.g. Lidgard et al. 1993), even if the average cyclostome diversity within assemblages showed a general decline through the Cenozoic, with a sharp drop in the late Neogene (Taylor 2000b). The observed peak of bryozoan diversity in Priabonian (Horowitz & Pachut 1996) seems to be an artifact of sampling because of well-studied faunas of this age in Europe and North America (Taylor 2000b). No striking mass extinction events are known for bryozoans in the Cenozoic except of the end-Danian (e.g. Voigt 1985, McKinney & Taylor 2001). Even the well-known Eocene-Oligocene event (Prothero 1994) does not show significant effect on the bryozoan record (Taylor 2000b). Locally, Mediterranean bryozoans were affected by the Messinian event in the late Miocene (Moissette & Pouyet 1987). The climatic variations seem to influence bryozoan diversity in the Cenozoic, as global cooling in the Pleistocene and Holocene affected the tropical bryozoan faunas. One of the most influential events in the Late Cenozoic was the formation of the Central America Isthmus (e.g. Lessios 2008). The final closure of the Isthmus seaway is estimated by 3–3.5 Ma (e.g. Knowlton & Weigt 1998, Coates & Stallard 2013, Jackson & O’Dea 2013). Geographic isolation and changes in primary production stimulated speciation and influenced modes of reproduction in bryozoans on

both sides of the Panama Isthmus (e.g. Jackson & Cheetham 1994, Cheetham et al. 1999, 2001, O’Dea & Jackson 2002, 2009, O’Dea 2003, O’Dea et al. 2004, 2008, Herrera-Cubilla et al. 2006). Otherwise, the collapse of primary productivity in the Caribbean due to closure of the Panama Isthmus caused mass extinction among erect and free-living cheilostomes but simultaneously led to the diversification of encrusting forms (Cheetham & Jackson 1996, Di Martino et al. 2018).

2.5 Bryozoans in reefs and other organic build-ups Although bryozoans were never as important reef-building organisms as corals, their contribution to the production of organogenic structures appears significant (e.g. Cuffey 1977, 1985, 2006). Bryozoans involved in reefs mainly played roles of principal frame-builders, sediment-binders, accessory reef encrusters, inhibitors of sediment movement, and sediment formers (Cuffey 1974, 1977). In total, three groups of bryozoans appear to be most important in reef constructions through geological time (Cuffey 1977, Fagerstrom 1987, Wood 1998, 1999): 1) massive, encrusting and branched trepostomes and cystoporates in the Ordovician and Silurian; 2) large frondose and branched fenestrates in the Carboniferous and Permian; and 3) massive and encrusting cheilostomes in the Cenozoic. In combination with other organisms, bryozoans produced significant organic build-ups starting as early as the Ordovician (e.g. Adachi et al. 2012, Cuffey et al. 2012). These constructions often represented consortia of microbes and bryozoans as well as other metazoans such as pelmatozoans, lithistid and stromatoporoid sponges, and tabulate corals (e.g. Cuffey et al. 2002, Webby 2002, Kröger et al. 2017, Hong et al. 2018). Scholz (2000) suggested the term “bryostromatolith” for biogenic constructions built by bryozoans and microbes and of typical laminated structure (Fig. 2.10 F–G). As principal frame-builders or sediment stabilizers, branched and encrusting trepostomes and cystoporates were involved in various types of reefs and bioherms during the Ordovician (e.g. McKinney et al. 2001b, Mehrtens & Cuffey 2003). They also become major contributors to hardgrounds in the Middle Ordovician (e.g. Wilson et al. 1992). Extensive bryozoan-dominated or bryozoan-bearing biogenic constructions were identified in the Ordovician sediments around the globe, notably in North America, Europe, North Africa, and China (Webby 2002). At the beginning of the Silurian, reef growth was suppressed, recovered slowly and reached its maximum in the Wenlock (e.g. Copper 2002). Common types of biogenic



constructions in the Silurian were bryozoan-microbial reefs associated with encrusting bryozoans (e.g. Ernst et al. 2015, Li et al. 2018). Otherwise, bryozoans contributed to different types of small patch-reefs known from some areas such as Canada or China (e.g. Hewitt & Cuffey 1985, Qiu 1990). Devonian bryozoan reefs and bioherms are rather scarce (Fagerstrom 1994, Wood 1999, Scholz et al. 2005, Cuffey 2006). They were involved in the early stages of reef formation and were mainly scattered reef-dwellers (e.g. Ernst & Königshof 2008) or minor components of coraldominated reef-mounds (e.g. Fernández et al. 1996, van Loevezijn & Raven 2017). In the Carboniferous, a new type of organic structure became widespread, namely mudmounds, which were produced by baffling activity of fenestrate bryozoans (e.g. Fagerstrom 1994, Cuffey 2006, Wyse Jackson 2006). Due to cilia-generated feeding currents, sediment is moved to the stagnant regions near the sea bottom, enhancing sediment precipitation (McKinney et al. 1987). Other bryozoan groups such as branched trepostomes or encrusting fistuliporines, often in association with other invertebrates and algae, were involved in various reefs and bioherms of the Carboniferous (e.g. Sugimura & Ota 1971, Bonem 1978). The peak of the bryozoan biogenic activity in reefal structures occurred in the Viséan (Webb 2002). Bryozoan-produced buildups in the Permian are similar to those of the Carboniferous and are dominated by arborescent fenestrate bryozoans. A particular case is bryozoanmicrobial reefs that originated on shelves of the Late Permian Zechstein Sea. These reefs reached sizes up to 100 m in height, persisted in high-salinity epeiric sea and covered vast areas in present-day Poland, Germany, and Great Britain (e.g. Kerkmann 1969, Paul 1980, Hollingworth & Tucker 1987, Peryt et al. 2016). The bryozoan fauna included mainly fenestrate taxa, accompanied by few trepostomes and cystoporates (e.g. Korn 1930, Dreyer 1961, Ernst 2001, 2002, 2007). Another example of the Permian reefs with involved bryozoan fauna is represented by the Late Permian Capitan reef, which is mainly produced by frondose bryozoans and sponges (Wood et al. 1996). It is characteristic for bryozoans in the Late Paleozoic that they largely occur in biogenic constructions of deeper water, in contrast to organisms with aragonitic and high-Mg calcite skeletons such as calcareous algae and sponges (Wahlman 2002). In the post-Palaeozoic time, bryozoans were rarely involved in organic buildups on such a scale as in the Paleozoic. Bryozoans are important components in some Triassic reefs (e.g. Senowbari-Daryan et al. 1993, Schäfer & Fois 1986), but largely occurred in non-reefal sediments. In the Jurassic and Cretaceous, bryozoan-dominated biogenic constructions were also rare.

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Bryozoan-rich sediments in this interval, especially in the Cenozoic, are mainly cool-water carbonates produced in higher latitudes (Taylor & Allison 1998). Warm-water tropical carbonates experience early lithification, whereas cool- or cold-water carbonates remain long time uncemented (e.g. Flügel 2010). As a result, organic particles are often strongly abraded (e.g. Smith & Nelson 1994, 1996, Taylor & James 2013) and do not contribute significantly to reefal structures. However, bryozoans are often the principal contributor of the carbonate production outside of the tropics in the Cenozoic (e.g. Nelson et al. 1988, Spjeldnaes & Moissette 1997), being a part of the bryozoan-molluscan (BRYOMOL) dominated facies (Fig. 2.14 F, G). Especially the Danish Basin in Europe represents a huge area of carbonate precipitation with significant contribution of bryozoans as carbonate particles (e.g. Cheetham 1971, Surlyk 1997, Bernecker & Weidlich 2005, Bjerager & Surlyk 2007; Fig. 2.14 E). The total thickness of carbonates produced here in the Late Cretaceous and Early Cenozoic exceeds 2,000 m. Extensive bryozoan mounds containing abundant branched cyclostomes and cheilostomes were developed in relatively deep water below the photic zone. Impressive bryozoan mounds are known from the middle Eocene of the Great Australian Bight (Sharples et al. 2014). These mounds were deposited parallel to the shelf margin for more than 500 km containing individual reef mound complexes up to 60–150 km long and 15 km wide and as thick as 200 m. Despite the clear dominance of coral reefs, bryozoan-dominated biogenic constructions account for ca 5% of buildups in the Cenozoic (Perrin 2002). Bryozoan-dominated buildups are characterized by high diversity, mainly associated with heterotrophic organisms (e.g. Herbig 1986, Pisera 1996, Martin et al. 1997). Bryozoans are often dominant in deep-water mud mounds, often in association with corals and sponges (e.g. James & von der Borsch 1991). Recent bryozoan-stromatolite reefs are known in shallow waters of southwestern Netherlands (Bijma & Boekschoten 1985).

2.6 Evolutionary patterns in bryozoans Bryozoans are an excellent group for studying evolution. Their morphological complexity, often well represented in the skeleton, and their abundant fossil record represent outstanding conditions for evolutionary studies. Moreover, bryozoans are exclusively colonial, “modular” organisms, a key innovation that constitutes an additional level for the impact of evolutionary processes (Hageman 2003).

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The modular construction of bryozoans is an immensely important aspect in evolutionary studies of this group. Coloniality itself represents a huge advantage but also bears some risks. Advantages include task sharing (polymorphism), higher levels of regeneration, higher fertility and life duration, better feeding, as well as ecological plasticity and efficient utilization of space for living (e.g. Beklemishev 1969, Jackson & Winston 1981, Hughes 1983, Coates & Jackson 1985, Jackson & McKinney 1990, Marfenin 1997, Borges 2009, Simpson et al. 2017). However, modularity results in low rates of evolution, lower system diversity, and higher susceptibility for infections due to asexual reproduction (e.g. McKey et al. 2010). The costs of modularity seem to be negligible, however, because of the clear evolutionary success of modular organisms. Colonial integration (degree of interdependence and cooperation among modules) leads to an increase of colonial functions and to the development of non-feeding heterozooids when required (e.g. Boardman & Cheetham 1973, McKinney & Jackson 1989). Especially cheilostomes show an immense increase of integration in their evolution. In highly polymorphic bryozoans, phenotypic differences of zooids in highly integrated colonies are accompanied by partial or complete functional loss of many organ systems, including feeding and sexual competency (Ryland 1970, Carter et al. 2010, Ostrovsky 2013). The influences of the physical environment, including such essential characteristics as water energy, substrate availability, but also interactions with biotic agents (competition on substrate, predation, food) are important factors of bryozoan evolution. In the following, important evolutionary processes and patterns observed in bryozoans are discussed.

2.6.1 Macroevolutionary trends in bryozoans Macroevolutionary trends affect the fundamental aspects of evolution of organisms, their most vital functions that determine their evolutionary success. Evolutionary trend means a long-termed change in a given direction (Stanley 1979). In bryozoans, the following aspects appear to be fundamental: 1) growth, 2) feeding, 3) reproduction, and 4) defense.

2.6.1.1 Growth and growth forms Calcified bryozoans are known from the Early Ordovician, but they certainly had long foregoing evolution.

Unmineralized bryozoans (phylactolaemates and ctenostomes) show a relatively simple organization and are regarded as ancestral to the calcified clades (e.g. Taylor & Larwood 1990, Gorjunova 1996, Todd 2000, Taylor & Waeschenbach 2015). Acquisition of a calcified skeleton had crucial consequences because rigid walls do not allow deflection of the body wall required for polypide eversion (e.g. Taylor 1981b, Larwood & Taylor 1979, see chapter 3). Therefore, special adaptations are required in order to achieve the hydrostatic pressure inside of a rigid construction (e.g. Taylor & Larwood 1990). Another effect of skeletal mineralization is the necessity of additional communication between zooids. Biomineralization evolved at least twice in Bryozoa. In the Early Palaeozoic (apparently Cambrian), calcification of the outer body of a soft-bodied, Arachnidiumlike bryozoan apparently gave rise to stenolaemates (Larwood & Taylor 1979, Gorjunova 1996). Again, in the Jurassic, cheilostomes arose from ctenostomes (Banta 1976, Taylor 1986). Calcification has an immense effect for an organism because it increases its constructional ability. Soft-bodied organisms have limited possibilities to produce different growth forms. The majority of ctenostomes and phylactolaemates develop fewer, and often simple, encrusting growth forms (e.g. Ryland 1970, Larwood & Taylor 1979, Taylor & Larwood 1990). Rigid erect colonies with calcified walls and extrazooidal skeleton allow to escape the competition for space at the substrate, provide better access to food in the water column, and increase colony surface area (e.g. McKinney & Jackson 1989). An erect mode of life brings some problems, too. They must have sufficient mechanical stability, especially on soft, unstable substrates. Therefore, bryozoans develop various kinds of supporting structures and structural heterozooids summarized under the name kenozooids (Levinsen 1902, 1909). Mesozooecia and exilazooecia of trepostomes and their kind in other groups can be regarded as being kenozooids (compare to the contrasting opinion by Boardman & Buttler 2005, who consider these structures rather as products of extrazooidal tissue and therefore call them mesopores and exilapores). Such tubular or prismatic structures have a constructional (space-filling) role for colonies that expand in volume (e.g. Key et al. 2001). Important trends in the growth of bryozoan colonies (Cheilostomata) are represented by zooidal (Lidgard 1986) and frontal budding (Banta 1972). Both strategies represent a significant advantage against the more primitive interzooidal budding because of faster growth on the substrate and, thus, occupation of space (McKinney & Jackson 1989).



2.6.1.2 Feeding Bryozoans show a universal mode of feeding by extracting their food from self-generated water currents. Water currents are produced by orchestrated beating of cilia on tentacles (e.g. Cook 1977; Winston 1977, 1978, see chapter 6). The size of the lophophore and the number of tentacles influence the feeding capacity of bryozoans giving them obvious competitive advantages (e.g. McKinney 1992, 1993a, Lidgard et al. 1993). However, the size and morphology of lophophores alone do not maintain the success of feeding. Bryozoans also face problems of utilization of filtered water, so they exhibit various patterns of coordinated activity of autozooids often supported by colonial phenomena (McKinney & Jackson 1989, McKinney 1990). Adaptations to expel filtered water include maculae that consist of skeletal material or heterozooids (Banta et al. 1974, Taylor 1975, 1979) or sharp edges of branches lacking zooecia (McKinney & Jackson 1989). Colony forms themselves are also important instruments for regulation and enhancement of feeding currents (Cowen & Rider 1972, McKinney 1977, 1981b, McKinney & Jackson 1989). Reticulate colonies of fenestrates were apparently efficient for enhanced feeding, which determined the success of this group in the Late Palaeozoic (Cowen & Rider 1972, Taylor 1999). Complex growth forms like a spiralled system of fans developed in Palaeozoic genera Archimedes, Ikelarchimedes, and Kazarchimedes, as well as in the Recent cheilostome Bugula turrita, provided up to 30 times increase of filtering surface per substrate surface taken by the colony (McKinney 1980, 1981b). Especially the genus Archimedes, with more than 60 species, achieved wide distribution and abundance in the Carboniferous of North America (McKinney 1993b; Fig. 2.12 B).

2.6.1.3 Reproduction Bryozoan colonies grow asexually by zooidal budding. However, new colonies are formed via sexually developed larvae (see chapters 4 and 5), although asexual propagation by fragmentation is known (McKinney 1983, O’Dea et al. 2004). Generally, two types of larvae, feeding, longlived, and non-feeding, short-lived, are found in Bryozoa. The first type is regarded being the ancestral condition and usually does not need brooding care (e.g. Zimmer & Woollacott 1977, Taylor 1988, see chapters 4 and 5). The transition from a long-lived feeding larva to a short-lived non-feeding one should result in the isolation of distant populations, thus accelerating speciation rates (e.g. Jackson 1986, Taylor 1988, Jablonski 1986, 2005,

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Jablonski & Lutz 1983). Such a transition can be stated if brooding structures (brooding care) are developed. In the Palaeostomata, brooding care was especially distributed in the Order Fenestrata, whereas indications for brooding in other orders are rather sparse (see section 2.3.1.2). In the Cheilostomata, no brooding structures are known before the Albian (Taylor 1988), and the appearance of the nonfeeding (coronate) larval type had apparently triggered the cheilostome radiation in the Mesozoic (Lidgard et al. 1993). Another evolutionary aspect of brooding concerns polyembryony, a phenomenon known from Stenolaemata (e.g. Borg 1926, Ström 1977; see chapter 4). Polyembryony, or embryonic fission, has developed in tubuliporine cyclostomes and was apparently also characteristic for the Palaeostomata (e.g. McKinney 1981a, Pachut & Fisherkeller 2010, Ostrovsky 2013). In polyembryony, one (primary) embryo forms many (up to 100) secondary embryos, which are genetically identical (clones). Secondary embryos are brooded in one or few large gonozooids within a colony. In cheilostomes, a single embryo is usually brooded within an ovicell (cheilostome-typical brood chamber) of which many can be developed in a single colony (see overviews in Ström 1977, Ostrovsky 2013; chapter 4). All the embryos are genetically different. Both strategies (large gonozooids with many identical embryos vs. many smaller ovicells with one genetically unique embryo per ovicell) have different evolutionary importance. However, the cheilostome brooding strategy bears some clear benefits; in first line, higher genetic variability, which brings faster adaptation and higher evolutionary rates, as well as higher insusceptibility for infections and lower energetic costs for brooding (e.g. Taylor 1988, Schäfer 1991, Lidgard et al. 1993). Moreover, in case of partial destruction of a colony, gonozooid(-s) may be lost, resulting in complete interruption of reproduction. In cheilostomes, ovicells in survived parts of partially destroyed colonies can secure reproductive success.

2.6.1.4 Defense Bryozoans are eaten by various animals (e.g. McKinney et al. 2003, Lidgard 2008). The targets of predators are individual zooids or whole colonies. As sessile animals without possibility of escape, they need effective defensive structures and defensive-relevant strategies. Therefore, predation represents an immensely important driving factor in the evolution of bryozoans (e.g. Dyrynda 1986). Modern bryozoans are known to react on predation by the development of spines (e.g. Yoshioka 1982, Harvell

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1984). Besides mechanical methods of defense, various arsenals of chemical substances are used by bryozoans to repel attackers (e.g. Figuerola et al. 2014). Indications for the evolutionary response to predation are known from fossil bryozoans, too. Anstey (1990) supposed that an increase of zooecial depth was a distinct trend in Palaeozoic stenolaemates, which he interpreted as a response to increasing predation. Trepostome bryozoans developed deeper zooecia starting from the Late Silurian, which coincided with the occurrence of the midPaleozoic predator revolution (e.g. Signor & Brett 1984, Brett & Walker 2002). Another trend that can be interpreted as being defensive is the development of various types of styles and inhomogeneities in the walls of Palaeostomata (Ernst 2013). Such adaptations apparently served the strengthening of colonies as protection against grazers. Also fenestrates, one of the most successful clades of the Palaeostomata, developed various protective structures in the form of dashed keels or protecting nets produced by fused keel nodes. Remarkably, taxa with such structures appeared in the time of increased predation from Silurian to the Late Devonian (McKinney et al. 2003, Ernst 2013), and their majority disappeared by the beginning of the Carboniferous. It appears obvious that the production of defensive structures is associated with high energetic costs for bryozoans, and taxa with defensive structures are rather disadvantaged in conditions of lower predation. In the post-Palaeozoic clades, a variety of evolutionary trends related to defense are known. The majority of post-Palaeozoic cyclostomes possess calcified frontal walls, and the number and grade of calcification of their frontal walls increased in the interval from the Early Cretaceous to the Holocene (McKinney et al. 2003). A parallel trend is observed in cheilostomes from the Late Jurassic until the present. Increasing calcification of frontal shields in Cheilostomata is regarded as strengthening character of branched colonies and may be interpreted as adaptation against grazing (e.g. Cheetham & Thomsen 1981, Cheetham 1986, Anstey 1990). Other obvious adaptations against predation of cheilostomes include opercula, spines, and various heteromorphs, especially avicularia (e.g. Winston 1984, Carter et al. 2010, 2011). Similar structures are known in fossil bryozoan groups, for example, aviculomorphs in Fenestrata (McKinney 1998) or eleozooids in melicerititid cyclostomes (e.g. Taylor 1985b, Jablonski et al. 1997), which are also of supposed protective function. The development and distribution of erect growth forms can be regarded as influenced by predator stress (e.g. Anstey 1987, McKinney & Jackson 1989, McKinney et al. 2003). In contrast to encrusting colonies, erect ones are more vulnerable to grazing. Therefore, the decreased

number of erect growth forms from Early Cretaceous to Holocene is explained by increased predation. Moreover, the shift of the erect growth forms from the shallow areas into deeper and more cryptic habitats is supposed to be a reaction to the diversification of various predators during the Late Mesozoic and Early Cenozoic.

2.6.2 Homeomorphy and convergence Bryozoans demonstrate various examples of homeomorphy (e.g. Voigt & Flor 1970, Hinds 1975, Blake 1980, McKinney et al. 1993, Taylor & Badve 1995, Taylor 1987, Taylor & McKinney 1996, Ernst et al. 2012c). Homeomorphy results from convergent evolution of traits (homoplasy), often through heterochrony (Anstey 1987). Important developments like the evolution of free-walled stenolaemates from fixed-walled ancestors are explained by heterochrony (Larwood & Taylor 1979, Taylor 2000a). Both paedomorphosis and peromorphosis are known among bryozoans at almost equal rates (Anstey 1987). As modular organisms, bryozoans demonstrate homeomorphy both on colonial and zooidal level. For example, lyra-shaped colonies evolved in Mississippian fenestrates and in Eocene cyclostomes (McKinney et al. 1993) or helical colonies in the Mississippian fenestrate Archimedes and the Eocene cyclostome Crisidmonea (Taylor & McKinney 1996). Numerous examples of homeomorphy are also known on the zooidal level. One of the best examples in stenolaemate bryozoans is demonstrated by the development of four-sided autozooecial chambers in trepostome bryozoans (Boardman & McKinney 1976), which developed independently several times during the Palaeozoic (e.g. Rhombotrypa, Rhombotrypella, Tetratoechus, and Eodyscritella). Lunaria are a characteristic feature of cystoporate bryozoans, but lunaria-like structures are also known from cryptostomes and cyclostomes (Voigt 1993, Ernst et al. 2012c). Also polymorphs show homeomorphy. Aviculomorphs of Palaeozoic fenestrates and eleozooids of melicerititid cyclostomes are analogous formations to avicularia of cheilostomes (e.g. Taylor 1985b, McKinney 1998). Certainly, the wide extent of homeomorphy significantly compromises stenolaemate bryozoan taxonomy. Indeed, molecular sequence data obtained from Recent cyclostomes has shown numerous skeletal morphological characters (e.g. maculae, diaphragms or hemiphragms, brooding structures) to be homoplaseous (Waeschenbach et al. 2009). Therefore, caution is required in interpreting morphological characters of bryozoans, especially in taxa distantly positioned in age.

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Wissenschaftlichen Erforschungen des Balatonsees Vol. 2(1). Hölzel, Wien: 1–22. Vine, G.R. (1884): Fourth report of the committee consisting of Dr. H.R. Sorby and Mr. G.R. Vine, appointed for the purpose of reporting on fossil Polyzoa. In: Reports of the 53rd Meeting of the British Association for the Advancement in Sciences. John Murray, London: 161–209. Vinogradov, A.V. (1996): New fossil freshwater bryozoans from the Asiatic part of Russia and Kazakhstan. Paleontol J 30: 284–292. Viskova, L.A. (1992): Marine post-Palaeozoic Bryozoa [Russian]. Trudy Paleontolog Inst Akad Nauk SSSR 250: 1–187. Viskova, L.A. & Morozova, I.P. (1988): A revision of the system of higher taxa of the phylum Bryozoa [Russian]. Paleontol Zhur 1988(1): 10–21. Viskova, L.A. & Pakhnevich, A.V. (2010): A new boring bryozoan from the Middle Jurassic of the Moscow region and its micro-CT research. Paleontol J 44(2): 157–167. Voigt, E. (1977): Arachnidium jurassicum n. sp. (Bryoz. Ctenostomata) aus dem mittleren Dogger von Goslar am Harz. Neues Jahrb Geol Palaontol Abh 153: 170–179. Voigt, E. (1985): The Bryozoa of the Cretaceous-Tertiary boundary. In: Nielsen, C. & Larwood, G.P. (eds.). Bryozoa: Ordovician to Recent. Olsen & Olsen, Fredensborg: 329–342. Voigt, E. (1993): Zwei neue Bryozoen-Genera (Cyclostomata) aus dem westfalischen Cenoman. Zitteliana 20: 361–368. Voigt, E. & Flor, F.D. (1970): Homöomorphien bei fossilien cyclostomen Bryozoen, dargestellt am Beispiel der Gattung Spiropora Lamouroux, 1821. Mitt Geol Staatsinst Hamb 39: 7–96. Walter, B. (1993): Une nouvelle faune de bryozoaires de l’Hauterivien inferieur du Jura. Geobios 26(5): 555–574. Walliser, O.H. (1996): Global events in the Devonian and Carboniferous. In: Walliser, O.H. (ed.). Global Events and Events Stratigraphy in the Phanerozoic. Springer, Berlin, Heidelberg: 225–250. Waeschenbach, A., Cox, C.C., Littlewood, D.T.J., Porter, J.S. & Taylor, P.D. (2009): First molecular estimate of cyclostome bryozoan phylogeny confirms extensive homoplasy among skeletal characters used in traditional taxonomy. Mol Phylogenet Evol 52(1): 241–251. Waeschenbach, A., Taylor, P.D. & Littlewood, D.T.J. (2012): A molecular phylogeny of bryozoans. Mol Phylogenet Evol 62: 718–735. Wahlman, G.P. (2002): Upper Carboniferous-Lower Permian (Bashkirian-Kungarian) mounds and reefs. In: Kiessling, W., Flügel, E. & Golonka, J. (eds.). Phanerozoic reef patterns. SEPM Special Publication 72: 271–338. Walter, B. (1969): Les Bryozoaires jurassiques en France. Étude systématique. Rapports avec la stratigraphie et la paléoécologie. Doc Lab Géol Fac Sci Lyon 35: 1–328. Wang, X.-D., Wang, X.-J., Zhang, H. (2001): Diversity patterns of Carboniferous and Permian rugose corals in South China. Geol J 41: 329–343. Wang, X.D., Wang, X.J., Zhang, F. & Zhang, H. (2006): Diversity patterns of Carboniferous and Permian rugose corals in South China. Geol J 41: 329–343. Webb, B. (2002): Latest Devonian and Early Carboniferous reefs: depressed reef building after the Middle Paleozoic collapse. In: Kiessling, W., Flügel, E. & Golonka, J. (eds.). Phanerozoic reef patterns. SEPM Spec Publ 72: 239–270.

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Webby, B. (2002): Patterns of Ordovician reef development. In: Kiessling, W., Flügel, E. & Golonka, J. (eds.). Phanerozoic reef patterns. SEPM Special Publication 72: 129–179. White, C.A. (1878): Descriptions of new species of invertebrate fossils from the Carboniferous and Upper Silurian rocks of Illinois and Indiana. Proc Acad Natl Sci Phila 30: 29–37. Wignall, P.B. & Hallam, A. (1992): Anoxia as a cause of the Permian/ Triassic mass extinction: facies evidence from northern Italy and the Western United States. Palaeogeogr Palaeoclimat Palaeoecol 93: 21–46. Wilson, M.A. & Taylor, P.D. (2014): The morphology and affinities of Allonema and Ascodictyon, two abundant Palaeozoic encrusters commonly misattributed to the ctenostome bryozoans. In: Rosso, A., Wyse Jackson, P.N. & Porter, J.S. (eds.). Bryozoan Studies 2013. Museo delle Scienze, Trento: 259–266. Wilson, M.A., Bosch, S. & Taylor, P.D. (2015): Middle Jurassic (Callovian) cyclostome bryozoans from the Tethyan tropics (Matmor Formation, southern Israel). Bull Geosci 90: 51–63. Wilson, M.A., Palmer, T.J., Guensburg, T.E., Finton, C.D. & Kaufman, L.E. (1992): The development of an Early Ordovician hardground community in response to rapid sea-floor calcite precipitation. Lethaia 25: 19–34. Winston, J.E. (1977): Feeding in marine bryozoans. In: Woollacott, R.M., Zimmer, R.L. (eds.). Biology of Bryozoans. Academic Press, New York: 233–271. Winston, J.E. (1978): Polypide morphology and feeding behaviour in marine ectoprocts. Bull Mar Sci 28: 1–31. Winston, J.E. (1984): Why bryozoans have avicularia – a review of the evidence. Am Mus Novit 2789: 1–26. Wood, A. (1948): “Sphaerocodium,” a misinterpreted fossil from the Wenlock Limestone. Proc Geol Assoc 59: 9-IN5. Wood, R. (1998): The ecological evolution of reefs. Annu Rev Ecol Syst 29: 179–206.

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Wood, R. (1999): Reef Evolution. Oxford University Press, Oxford. Wood, R., Dickson, J.A.D. & Kirkland-George, B. (1996): New observations on the ecology of the Permian Capitan Reef, Guadalupe Mountains, Texas and New Mexico. Palaeontology 39: 733–762. Wöss, E.R. (2005): Biologie der Süßwassermoostiere (Bryozoa). Denisia 16: 21–48. Wyse Jackson, P.N. (2006): Bryozoa from Waulsortian buildups and their lateral facies (Mississippian, Carboniferous) in Belgium and Ireland. Courier Forschungsinstitut Senckenberg 257: 149–160. Wyse Jackson, P.N. & Bancroft, A.J. (1994): Possible opercular structures in the fenestrate bryozoan Thamniscus from the Upper Carboniferous of northern England. In: Hayward, P.J., Ryland, J.S. & Taylor, P.D. (eds.). Biology and Palaeobiology of Bryozoans. Olsen & Olsen, Fredensborg: 215–218. Wyse Jackson, P.N. & Bancroft, A.J. (1995): Generic revision of the cryptostome bryozoan Rhabdomeson Young & Young, 1874, with description of two species from the Lower Carboniferous of the British Isles. J Paleontol 69(1): 28–45. Xia, F. (2002): Fenestrate Bryozoa with avicularia-like structures from the Middle Jurassic of north Tibet and the origin of cheilostome bryozoans. Acta Micropalaeontol Sin 19(3): 237–255. Xia, F., Zhang, S. & Wang, Z.-Z. (2007): The oldest bryozoans: new evidence from the Late Tremadocian (Early Ordovician) of east Yangtze Gorges in China. J Paleontol 81(6): 1308–1326. Yoshioka, P. (1982): Predator-induced polymorphism in the bryozoan Membranipora mebranacea (L.). J Exp Mar Biol Ecol 61: 233–242. Zimmer, R.L. & Woollacott, R.M. (1977): Structure and classification of gymnolaemate larvae. In: Woollacott, R.M. & Zimmer, R.L. (eds.). Biology of Bryozoans. Academic Press, New York: 57–89.

Thomas Schwaha

3 Morphology of bryozoans 3.1 A short glimpse into morphological and anatomical research of bryozoans Research on the morphology or anatomy of bryozoans dates back way into the 19th century. Researchers as early as 1837 (Farre 1837) and others (e.g. Hancock 1850, Allman 1856) were successful in studying some general body plan features such as lophophore or digestive tract structure, or even thick retractor muscle bundles. More detailed observations were conducted in the late 19th century (Nitsche 1868, 1869, 1871, Reichert 1870, Kraepelin 1887, 1892, Braem 1890), which also started to use sectioning methods for histological analyses. Despite their age, some of these observations proved to be extraordinarily correct and had marvelous precision in their descriptions (e.g. Braem 1890). In the end of the 19th and beginning of the 20th century, several contributions on the morphology and development were published, which for example dealt with the development of cyclostomes (Harmer 1893, 1896, 1898), morphology of gymnolaemates (Calvet 1900, Harmer 1902), or the large work of Borg (1926) on the anatomy of cyclostome bryozoans. To date, the latter remains one of the most detailed studies on cyclostome anatomy. Toward the mid-20th century, several studies form Marcus were among the most important achievements in morphological research (Marcus 1926a, b, 1934, 1937, 1938, 1939, 1941, 1942). His research covered morphological studies on phylactolaemate, ctenostome, and cheilostome bryozoans. Another important researcher advancing the field of morphological research was Silén, who published important contributions on various aspects of mostly gymnolaemate bryozoans in the course of almost five decades (e.g. Silén 1938, 1942, 1944a, b, c, 1945, 1947). In the second half of the 20th century, a number of various important contributions were conducted. A few examples are studies on the nervous system (Lutaud 1969, 1973a, b, 1974, 1976, 1977, 1979a, 1981, 1984, 1993), lophophoral base (Gordon 1974), gut structure (Bobin & Prenant 1952, Bullivant & Bils 1968, Gordon 1975a, b, c, Schäfer 1986, Markham & Ryland 1987), cyclostome morphology and development (Nielsen 1970, Nielsen & Pedersen 1979, Nielsen & Riisgard 1998), ctenostome pore plates (Bobin 1958a, b, 1962, 1964, 1971), or cheilostome body walls (Banta 1968a, b, 1969, 1970, 1971, 1972, 1973, 1977). https://doi.org/10.1515/9783110586312-003

3.2 Outline of this chapter The chapter gives a general introduction into the comparative morphology of bryozoans. It will focus primarily on zooidal (autozooidal level) starting with axes orientations and terminology, general zooidal organization including structure and function of different organ systems. Skeletal or cystid characters will not be dealt with here. Instead, soft tissue morphology will be comparatively presented. Specific taxon-specific structures are only briefly mentioned, with detailed explanations following in the chapters of the respective clade. Gonads are omitted as they will be treated in more detail in chapter 4 of this volume.

3.3 General body organization and features Bryozoans are essentially all colonial with individuals, zooids, forming a colony. Colony shape drastically varies among the clades, genera, and species. The general morphology of normal zooids, or in case of polymorphic taxa, the autozooid, is more or less identical within a colony. Each zooid is comprised of a cystid, which is the cellular body wall (endocyst) and its cuticle (ectocyst), and the polypide, which comprises the soft tissues such as the lophophore, digestive tract, and associated tissues. This division is purely artificial as both of these structures are interconnected, inseparable tissues (Mukai et al. 1997). However, the terms are well established and have been used ever since their proposition in the 19th century (see Allman 1856). Especially in the dominant groups Stenolaemata and Cheilostomata, cystid characters are often synonym for (calcified) skeletal structures. Uncalcified forms such as the Phylactolaemata and ctenostome Gymnolaemata lack a high diversity of cuticular structures. Although in use for ~150 years, a clear line is generally not drawn on assessing especially transitional structures such as the vestibular wall toward the regular body wall, i.e. whether they are part of the cystid or polypide. The best way to clearly separate zooidal parts as cystid and polypide is indicated whether the body parts experience distinct dislocations or movements during polypide retraction. Consequently, the cystid encompasses the body wall and its cuticle, interzooidal septa and pores when present,

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Fig. 3.1: General scheme of bryozoan zooids showing the zooid with the polypide protruded on the left and retracted on the right. The corresponding structures are highlighted in the same colors to facilitate understanding of the different structures in protruded and retracted zooids. Axis orientation refers to a single zooid. Abbreviations: a – anus, at – atrium, bw – body wall, cae – caecum, cg – cerebral ganglion, db – duplicature band, f – funiculus, int – intestine, l – lophophore, lb – lophophoral base, o – orifice, rm – retractor muscle, ts – tentacle sheath, v – vestibulum, vm – vestibular muscles, vw – vestibular wall.

muscles and neurites associated with the body wall, and other tissues such as gonads attached to the body wall. All the remaining parts starting from the distal tip with the vestibular wall, tentacle sheath, lophophore and digestive tract, and any attached muscles or peritoneal strands are part of the polypide (Fig. 3.1). Colonies and zooids always follow a certain growth direction. Most commonly, a larva settles to metamorphose and form the ancestrula, the founding zooid (Reed 1991). Depending on the specific taxon, different ways of asexual propagation or colony development, astogeny, are possible. In any case, a certain growth direction always persists, which is evident in the structure of the colony. Distinct colony margins mark the area of asexual bud proliferation and the formation of new zooids. On zooidal level, there is thus always a direction of their origin and direction toward the colony margin. These directions are always termed proximal – toward the origin – and distal – toward the newly formed zone or colony margin (e.g. Cheetham & Cook 1983, Mukai et al. 1997). Additional budding sites are orthogonal to the proximo-distal axis such as lateral buds on each side. Zooidal shapes vary from box shaped, to tubular, to sigmoidal (e.g. Jebram 1986). Zooids, including their

orificial/apertural area, are thus commonly bent toward one side. In myolaemates, this side is termed the frontal side. Phylactolaemate colonies show heterogeneous arrangements of zooids that range from tubular to densely aggregated as in the gelatinous forms. Consequently, a frontal side is traditionally not specified and the term is not used. The opposite side of the frontal side of myolaemates is referred to as the basal side (e.g. Boardman 1983, Cheetham & Cook 1983) and thus represents the side facing the substrate (in encrusters). In erect forms, the latter is often termed vertical wall, i.e. orthogonal to the substrate (e.g. Boardman 1983, 1998, Taylor et al. 2015). Some bryozoans can form buds on the frontal side (e.g. Lidgard 1990). The cystid terminates distally in each zooid at the orificial or apertural area. The term orifice is used for the opening bordered by soft tissues and is, particularly in phylactolaemates and ctenostomes, synonymously used with the term aperture. In cyclostomes and cheilostomes, the term aperture is more commonly used for the skeletal aperture, which does not necessarily coincide with the orifice. Past papers frequently used both terms in a random fashion. It should be emphasized that these terms should be used more cautiously. The outer opening in living zooids

3.4 Body cavity 



is always the orifice. In general, this part of the zooid can be addressed as apertural or orificial area. The orifice is always bordered by the vestibular wall (cf. Schwaha et al. 2011). In retracted zooids, it extends proximally into the inverted tentacle sheath. The diaphragmatic sphincter is always located at the border of these structures (see also below in muscle systems). The tentacle sheath itself is a thin and delicate structure lacking any distinct cuticle, whereas the vestibular wall is thicker and is often lined by a cuticle similar to the remaining body wall (e.g. Mukai et al. 1997). In protruded zooids, the tentacle sheath is always extended and shifts the lophophore into a higher position for suspension feeding. The shape and extent of the vestibular wall can show high variation among bryozoans in protruded zooids. In general, three different forms can be recognized: 1) the vestibular wall stays mostly invaginated and its epithelial lining is arranged as in retracted zooids; 2) the vestibular wall only slightly everts and forms only a shallow indentation at the orificial area; and 3) the vestibular wall, often of quite extensive length, is everted to its full extent similar to the tentacle sheath. In the latter case, the protruded zooid can extend far from the orifice/aperture or even the peristome. The latter is often the case in many ctenostomes (see chapter 10 of this volume). The collar of gymnolaemates is particularly indicative of the extent of vestibular wall protrusion, because it is located at the border of the vestibular wall toward the tentacle sheath. The tentacle sheath has previously been termed introvert in previously literature (e.g. Tamberg & Shunatova 2016, cf. Ryland 2005). The term introvert is functionally descriptive and is applicable as mentioned above sometimes also to the vestibular wall. In addition, the term introvert is used in many other phyla (e.g. many cycloneuralians). This strongly advocates the use of the term tentacle sheath as it is more precise and restricted to bryozoans. The epithelia of the tentacle sheath continue into the lophophoral base wherefrom the tentacles emerge. It is characterized by a distinctly thicker epithelium and an extensive extracellular matrix (ECM). Within the lophophoral base lies the mouth opening that continues into the u-shaped digestive tract, which terminates with the anus outside of the tentacle crown. The u-shaped digestive tract allows to address an anal vs. an oral side of each zooid. The side with the oral gut shank consequently is the oral side, whereas the side with the hindgut and anus represents the anal side. Some past articles sometimes referred to these sides as “dorsal” and “ventral” (e.g. Mukai et al. 1997), which, however, is not useful since it does neither reflect an ontogenetical dorso-ventral

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side nor a physiological one. The terms oral and anal are solely references for the polypide and not the entire zooid. Depending on the taxonomic clade and group, the anal and oral side of each polypide may be synonymous with the fronto-basal or proximo-distal side of each zooid.

3.4 Body cavity In all bryozoans, body cavities are essential for the hydrostatic polypide protrusion-retraction process. Retraction is always effectuated via the prominent retractor muscles (Figs. 3.2–3.4), whereas protrusion is effectuated by increasing the pressure in the zooidal body cavity via different mechanisms (cf. Ryland 1970, Taylor 1981). Bryozoans are in general coelomate animals, which means that a secondary body cavity is lined by a complete epithelium that has its apical side facing the voluminous cavity (cf. Bartolomaeus 2001). The coelomic epithelium or peritoneum is in its original state adjacent to the epidermis of the body wall, with both basal membranes facing each other. This situation as such is present only in the Phylactolaemata (Gruhl et al. 2009; see also Fig. 3.1). Phylactolaemates protrude their polypides via contraction of regular body wall musculature; i.e. the entire zooid is compressed, which forces the polypide outward (cf. Schwaha & Wanninger 2012, Gawin et al. 2017). Cyclostomes (Stenolaemata) have calcified cystids (either internal or external; see chapter 8) that surround most of the polypides and leave no compressible space for the hydrostatic protrusion mechanism (e.g. Taylor 1981). Consequently, cyclostomes evolved a separate hydrostatic mechanism in order to enable polypide protrusion: the peritoneal layer of the body wall detached from the epidermis to form the membranous sac (Borg 1926, Nielsen & Pedersen 1979). Medially toward the polypide, the peritoneum remains directly adjacent to the lining of the respective organs (e.g. gut or tentacle sheath). The membranous sac forms a flexible sac that surrounds the polypide. The separation of the peritoneum from the original body wall divides the body cavity into two distinct cavities: the one enclosed within the sac, which corresponds to the original coelomic cavity, called endosaccal cavity, and the cavity located between the basement membranes of the epidermis and the peritoneal lining, the exosaccal cavity (Fig. 3.3). The membranous sac has only few connections to the epidermis of the body wall: at the insertion of the retractor muscle and muscular funiculus and in the proximal apertural area when an attachment organ (ligamentous attachment of the proximal apertural area

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Fig. 3.2: Schematic drawing of the general organization of a phylactolaemate bryozoan showing the polypide in protruded (left) and retracted condition (right). Scheme on the left is modified and redrawn from Schwaha (2019). Abbreviations: a – anus, at – atrium, bc – body cavity, bw – body wall, ca – cardia, cae – caecum, cg – cerebral ganglion, db – duplicature band, ep – epistome, f – funiculus, fg – foregut, int – intestine, la – lophophoral arm, mo – mouth opening, o – orifice, pya – pyloric area, rm – retractor muscles, ts – tentacle sheath, v – vestibulum, vd – vestibular dilatator, vw – vestibular wall.

to the lateral skeletal wall) is present. Circular muscles, termed annular muscles, in the lining of the membranous sac enable polypide protrusion (Nielsen & Pedersen 1979, Taylor 1981, Schwaha et al. 2018). In gymnolaemates, a distinct coelomic cavity completely lined by a peritoneum is reduced in the body wall and only remains on the lining of internal organs (e.g. gut) (Mukai et al. 1997, Schwaha et al. 2020). Ontogenetically, both layers of the body wall are quite distinct in buds. After the complete formation of the zooid, however, the outer body wall consists mainly of a continuous epidermis and only a patchy inner peritoneal layer. This condition is already quite evident in light microscopical sections, and transmission electron microscopical examinations have recently confirmed this (Shunatova & Tamberg 2019). The reason for the reduction of the peritoneum is not known. Distinct gymnolaemate specific parietal muscles traverse the body cavity on each lateral side of the polypide and act in its compression in order to enable polypide protrusion (Fig. 3.4). Peritoneally derived tissues associated with the body wall are present as cells aggregating at interzooidal pores or as funicular strands of (mostly) cheilostomes (cf. Bobin 1977, Lutaud 1983). Gonads and their development

are generally also associated with mesodermal tissue associated with the body wall (see Ostrovsky 2013, and also chapter 4).

3.5 Coelomic partitions or canals In older literature, the coelomic cavity of bryozoans was considered trimeric, with three separate coelomic cavities arranged from proximal to distal. This hypothetical condition was thought to be present in phylactolaemates, with an anterior cavity in the epistome, a second in the lophophore, and a third below the latter. With the lack of an epistome, one of the three coelomic cavities was stated as lost in myolaemates (e.g. Hyman 1959). Newer observations have shown that there is no indication of a trimeric organization of bryozoans (cf. Gruhl et al. 2009, Schwaha et al. 2011, Schwaha 2018) and that previous assumptions were biased by an outdated and non-valid view that lophophorates, including bryozoans, are trimeric and closely related to deuterostomes such as ambulacrarians, which possess a trimeric body organization (cf. Schwaha et al. 2020).



3.5 Coelomic partitions or canals 

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Fig. 3.3: Schematic drawing of the general organization of a cyclostome bryozoan showing the polypide in protruded (right image) and retracted condition (left image). Redrawn and modified from Ryland (1970). Abbreviations: a – anus, anm – annular muscles of the membranous sac, atl – attachment ligament, cae – caecum, cg – cerebral ganglion, dex – distal exosaccal cavity, ds – diaphragmatic sphincter, ens – endosaccal cavity, exo – exosaccal cavity, f – funiculus, fg – foregut, int – intestine, lb – lophophoral base, o – orifice, py – pylorus, rm – retractor muscles, ts – tentacle sheath, v – vestibulum, vw – vestibular wall.

3.5.1 Phylactolaemata Phylactolaemate zooids show only little differentiation of their body cavity. The oral and anal rows of tentacles show separate canals: the orally located ring canal and the anally situated forked canal (Braem 1890; Fig. 3.5 A, B). Both canals form via the peritoneal layer of the polypide bud (outer budding layer) protruding medially against the epidermal layer. These protrusions later fuse during ontogeny to form a continuous canal (Schwaha et al. 2011). The ring canal supplies only a few tentacles on the oral side, ranging from four to eight. On its proximal side, muscles are located within the ECM of the two

peritoneal layers (Gawin et al. 2017, Schwaha 2020). Likewise, neurite bundles pass through this area from the pharynx and proceed into the tentacle sheath (Ambros et al. 2018). The forked canal commonly supplies only four to six tentacles located within the lophophoral concavity on the anal side. Its epithelium is thicker than the remaining peritoneum and also shows strong ciliation (Verworn 1887, Braem 1890, Cori 1890, 1893, Gruhl et al. 2009, Schwaha et al. 2011, Schwaha 2018, 2020). The forked canal has often been interpreted as vestigial metanephridium. Sperm and excretory coelomocytes/phagocytes were encountered in the lumen of the forked canal. Despite its unclear homology to possibly other nephridial structures,

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Fig. 3.4: Schematic drawing of the general organization of a gymnolaemate bryozoan showing its main body axes and the polypide in protruded (upper image) and retracted condition (lower image). Redrawn and modified from Cori (1941). Abbreviations: a – anus, ca – cardia, cae – caecum, cg – cerebral ganglion, co – collar, fg – foregut, fm – frontal membrane, int – intestine, lb – lophophoral base, ori – orifice, pm – parietal muscles, pp – pore plate, py – pylorus, rm – retractor muscles, ts – tentacle sheath, vw – vestibular wall.

it appears to carry out similar functions (cf. Schwaha et al. 2020). In Cristatella mucedo, a separate, supposed excretory, bladder can form at the median junction of the forked canal. An additional canal present in all phylactolaemates is the epistomial canal (Fig. 3.5 A, B). This emerges between the oral and anal shanks of the gut, extends distally and directly adjacent to the lining of the cerebral ganglion, and bends orally into the tongue-like protrusion over the mouth opening – the phylactolaemate specific epistome (cf. Gruhl et al. 2009, Schwaha et al. 2011, Schwaha 2018). The epistomial cavity strongly correlates with the size and extent of the epistome itself (see chapter 7).

3.5.2 Myolaemata The coelomic partitioning is only recently investigated in more detail (Shunatova & Tamberg 2019). In both cyclostomes and gymnolaemates, a ring canal is present at the lophophoral base. In gymnolaemates it has paired openings to the remaining body cavity on the anal side of the zooid, close to the cerebral ganglion (e.g. Mukai et al. 1997,

Schwaha et al. 2011; Fig. 3.5 C, D). In cyclostomes it was reported to be closed, lacking any pores to the remaining endosaccal cavity (Shunatova & Tamberg 2019). The ring canal of cyclostomes and gymnolaemates is generally crossed by the buccal dilatators (Schwaha & Wanninger 2018, Schwaha et al. 2018).

3.6 Body walls The body wall or endocyst (as cellular part of the cystid) can have up to two distinct epithelial layers, the outer epidermis that sheds the cuticle and/or the mineralized skeleton and the inner peritoneum or coelomic epithelium (cf. Mukai et al. 1997, Gruhl et al. 2009). These differ distinctly in each clade.

3.6.1 Phylactolaemata Since zooids are the largest encountered among all Bryozoa, the body wall is also the most prominent and thickest in size. The phylactolaemate epidermis is characterized by

3.6 Body walls 



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Fig. 3.5: Coelomic organization of bryozoans. (A, B) Phylactolaemata. A, Oblique view from the oral side of a horseshoe-shaped lophophore showing the ring canal on the oral and the forked canal on the anal side. The latter arches over the tongue-like epistome canal that extends in the epistome above the mouth opening (not displayed). General shape of the lophophore redrawn from Marcus (1934). B, Longitudinal median section of the lophophoral base showing the arrangement of the coelomic canals. Redrawn and modified from Schwaha (2018). (C, D) Gymnolaemate coelomic system. C, Oblique view as in (A). D, Section similar to B. Only a ring canal is present at the lophophoral base. Abbreviations: cg – cerebral ganglion, con – circum-oral nerve ring, ep – epistome, epc – epistome coelom, fc – forked canal, gah – ganglionic horns, gl – ganglion lumen, itm – intertentacular membrane, la – lophophoral arms, loc – lophophoral concavity, mo – mouth opening, rc – ring canal.

a high abundance of glandular cells of different sizes and shapes (Fig. 3.6). Glandular composition often varies also depending on the location, i.e. basal attached vs. lateral free wall. The content of the glandular vesicles has not been properly identified, but histochemical stains and slight indications on the basic chemical composition of glandular vesicles has been conducted and also applied for systematic purposes (Mukai & Oda 1980). One of the main functions of the glands is probably the secretion of either adhesive substances that agglutinate detritus or any particle to their body wall or secretion of the ectocyst itself. In gelatinous forms, glandular cells are also abundant in areas where no ectocyst is formed. Some of the gelatinous species also possess so-called white spots, which consist of concentrated glandular patches (see Gruhl 2013, see also chapter 7). The peritoneum has only few cellular inclusions and remains inconspicuous. It is ciliated and ciliary

bundles are regularly distributed over the entire range of the body wall. These are responsible for creating ciliary currents within the body cavity (e.g. Wood 2014). This circulation of body fluids runs proximally on the oral side and distally on the anal side (Meyer 1927). These circulatory actions are probably a consequence of the density of ciliary patches and beating direction. The peritoneum covering the digestive tract shows none to little ciliation on the oral side of the gut, but a densely ciliated anal side (cf. Ambros et al. 2018). Ciliary bundles acting in circulation are also present in the inner lining of the lophophoral arms.

3.6.2 Cyclostomata Due to the detachment of the peritoneum to form the membranous sac (see section 3.4, Fig. 3.3) only a thin

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Fig. 3.6: Body wall organization and diversity in phylactolaemate bryozoans. (A, B) Hyalinella punctata. (C, D) Cristatella mucedo. (E) Lophopus crystallinus. (F) Plumatella sp. A, General overview showing epidermal glands and a detached ectocyst. Note also peritoneal ciliation. B, Body wall in the apertural area showing prominent glands and the ectocyst interspersed with small, deeply stained inclusions. C, Body wall of the creeping sole showing distinct folds of the epidermis. D, Lateral body wall showing large glandular inclusions. E, Highly glandular apertural area. F, Body wall of a species not producing gelatinous ectocyst (not shown). Abbreviations: at – atrium, bwm – body wall musculature, crp – creeping sole, ec – ectocyst, egl – epidermal glands, ep – epidermis, o – orifice, pe – peritoneum, pec – peritoneal cilia, rm – retractor muscle, ts – tentacle sheath, vw – vestibular wall.



epidermis remains as body wall. The little available ultrastructural data, but also general histological appearance, show that the cyclostome epidermis is rather thin (Fig. 3.7 A–D). Apically, it secretes a thin cuticle (sometimes called periostracum, Nielsen & Pedersen 1979, Tavener-Smith & Williams 1972). Distinct variations are present within the different taxonomic groups depending whether zooids are fixed- or free-walled forms (see chapter 8). Fixed-walled cyclostomes such as Crisia (Nielsen & Pedersen 1979) have external calcification with a thick mineralized layer secreted from the epidermis. In free-walled forms, the mineralized skeleton is externally covered by additional membranous areas; i.e. the outermost layer of the body wall is an unmineralized cuticle followed by the epidermis (Brood 1976, Ross 1977, Boardman 1983; see also chapter 8). Pseudopores are small circular pores in the external mineralized skeleton. These appear as regular pores on the calcified layer of the body wall and can be easily recognized in many taxa. The epidermis forms distinct, large plugs that fill these holes. Externally, a thickened plug of cuticle fills the pseudopore (Ross 1977, Nielsen & Pedersen 1979; Fig. 3.7 D). Pseudopores are generally considered to function in gas exchange.

3.6.3 Gymnolaemata In general, the epidermis of gymnolaemates is similar in size to that of cyclostomes. The peritoneum is patchy and does not form a continuous epithelium (Fig. 3.7 E–H). Its few cells are elongate to spindle shaped and are adjacent to the epidermis (Mukai et al. 1997). The latter secretes a cuticle that remains unmineralized in ctenostomes and mineralizes in cheilostomes (Cheetham & Cook 1983). The chemical composition of the cuticle is unknown but often considered chitinous. Cuticle thickness can vary and basally is often rather thin compared to the frontal and lateral sides. In several ctenostomes, the cuticle is thick and composed of multiple layers that often histologically show different staining properties, which implies different components and properties of the cuticle. Particularly some ctenostomes often attach particles onto their ectocyst wall (Fig. 3.7 F). Since parietal muscles insert directly via the epidermis and attach to the cuticle, special “tendon” cells are present in the epidermis that are reinforced with tonofilaments.

3.7 Communication areas and pores As true colonial animals, the tissues of individual zooids comprising the colony are interconnected. The degree of

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this interzooidal connection varies among the different bryozoan clades. Phylactolaemates generally have widely open interconnections of the coelomic cavity. Thus, individual zooids are the least separated of all bryozoans. The degree of individual separation also varies within phylactolaemates. The large gelatinous species share a large coelomic cavity and polypides basically bathe in the same fluid, whereas the tubular branching forms such as the plumatellids of fredericellids have smaller interzooidal communication areas of the body cavity. These are at each branching point of the colony (Wood 2014). Several species of plumatellids also form interzooidal, cuticular septa that constrict, but not close, this interzooidal connection (Mukai et al. 1997, Wood & Okamura 2005). Cyclostomes are little investigated concerning interzooidal communication and the general morphological scheme of crisiid or tubuliporid cyclostomes depicted in all major text books (e.g. Ryland 1970, Mukai et al. 1997) show skeletal constrictions at the proximal side of zooids or in the vertical walls with open connections. These would allow fluid exchange from the exosaccal cavity of one zooid to another. There is, however, increasing data that show interzooidal pores to be plugged with cells (Schwaha, pers. observation, Fig. 3.7 B). Possibly both forms of open and plugged interconnections can co-occur but require more detailed analyses. Gymnolaemates always have cuticles (unmineralized in ctenostomes, and mineralized in cheilostomes) that form septa between neighboring zooids. These are pierced by thin pores that are plugged with a typical arrangement of several cells: special cells, cincture cells, and limiting cells (Bobin 1977). Special cells are dumbbell-shaped and plug the interzooidal pore via the thin midpiece of the dumbbell. Beyond the pore, the cells broaden into the bulbous parts. Only the bulbous part of the special cell in the proximal zooid has a nucleus, whereas the distal portion lacks any (Bobin 1977, Mukai et al. 1997). Cincture cells directly line the pore and contain muscle fibers (Gordon 1975d). Limiting cells mainly wrap around the special cells on each side of the pore. Pore plates, i.e. cuticular pores and associated cells, vary in their morphology from being large cellular aggregations to very small complexes (Fig. 3.8; see also Gordon 1975d). With few exceptions, pores are single on each wall in most ctenostome gymnolaemates, whereas there are multiple pores in the pore plates of cheilostomes (Mukai et al. 1997, Schwaha et al. 2020). It is not clear whether multiporous pore plates have multiple or a single origin, which could indicate whether cheilostomes might have a last common ancestor with one of the multiporous ctenostome clades.

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Fig. 3.7: Body wall structure in cyclostome and gymnolaemate bryozoans. (A–D) Cyclostomata. (E–H) Gymnolaemata. A, Cross-section of a decalcified colony branch of Cinctipora elegans showing little structure after mineralized substances have been removed. Only a thin cuticle surrounds the zooids. B, Details of the cystid wall of C. elegans close to the pharynx showing the cystid wall of an interzooidal wall. Note the few fibers in the decalcified cystid wall and cellular plugs in the interzooidal pores. C, Longitudinal section of a cystid wall of Hornera sp. showing little differentiation in the decalcified matrix. D, Pseudopores filled with prominent epidermal cells in Crisia sp. E, Section of the body wall of the ctenostome Hislopia malayensis showing a distinct epidermis and patchy, dispersed peritoneal cells. F, Cross-section of a branch of the ctenostome Anguinella palmata showing dense incrustations on top of the ectocyst. G, Ctenostome Paludicella articulata showing a thin epidermis below the cuticle and the attachment site of parietal muscles. H, Longitudinal section of the cheilostome Securiflustra securifrons. Few vesicular cells are visible close to the body wall. Abbreviations: bc – body cavity, cw – calcified wall, dg – digestive tract, ec – ectocyst, emb? – possible embryo embedded in cuticular incrustations, ep – epidermis, fpo – functional polypide, inc – incrustations of the ectocyst, izp – interzooidal pore, izw – interzooidal wall, lb – lophophoral base, pc – peritoneal cells, ph – pharynx, pm – parietal muscles, psu – pseudopores filled with cells, vec – vesicular cells.

3.8 Lophophore 

3.8 Lophophore 3.8.1 General structure Two lophophore forms can be distinguished: the horseshoe-shaped lophophore of the Phylactolaemata (Fig. 3.2) and the circular lophophore of the Myolaemata (Figs. 3.3 and 3.4). Phylactolaemate zooids are distinctly larger than myolaemate ones. Two lateral extensions, the lophophoral arms, extend into the anal direction to form the horseshoe-shape typical for this group. Notably, the only phylactolaemate showing a secondarily reduced circular lophophore as adults are fredericellids, which also show a lower number of tentacles (cf. Gruhl et al. 2009, Shunkina et al. 2015). Large lophophores of the gelatinous phylactolaemates have about 70 or more tentacles (Wood & Okamura 2005). A distinct feature of phylactolaemates is the presence of an intertentacular membrane at the lophophoral base (see chapter 7). The circular lophophore of myolaemates is in comparison to the phylactolaemate one smaller and carries fewer tentacles. In many ctenostomes and cyclostomes, only eight tentacles are present, whereas the upper range is limited to about ~30 tentacles. A range of 14–16 tentacles is quite common among myolaemates (Winston 1978). It never shows any indication of a horseshoe-shaped arrangement. A few ctenostomes possess an oral groove that apparently acts as rejection tract and gives the lophophore a distinct bilateral shape (Prouho 1892, Atkins 1932, Braem 1939). Gymnolaemates show distinct intertentacular pits (Gordon 1974, Schwaha et al. 2011, Weber et al. 2014, Schwaha et al. 2020). These are located on the lophophoral base between each pair of tentacles and form narrow and ciliated grooves (Gordon 1974). These pits are supposed to have a sensory function. A similar structure, so-called intertentacular bases, was recently found in the cyclostome Cinctipora elegans. These differ, however, to the intertentacular pits by lacking any distinguishable lumen and thus also ciliation (Schwaha et al. 2018).

3.8.2 Tentacles and ciliation Tentacles are generally triangular in bryozoans, with the pointed tip being directed toward the mouth opening (or the inner side of the lophophoral arm in case of corresponding tentacles of phylactolaemates). This side is

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traditionally referred to as frontal (not to confuse with the frontal zooidal side of myolaemates). The broader side of the tentacle is hence called abfrontal and situated opposite of the main feeding currents (Mukai et al. 1997; Fig. 3.9). In myolaemate bryozoans, a distinct arrangement of nine cells is present in tentacles in respect to their cross-section. Three narrow and elongated cells form the frontal side, two flattened cells the abfrontal, and four the lateral area (see Smith 1973, Lutaud 1973a, Nielsen & Riisgard 1998; Fig. 3.9 C, E). Phylactolaemates show a similar arrangement of epidermal cells of the tentacles. Their number, however, always exceeds nine, which probably corresponds to a general larger size of the lophophore and individual tentacles (Mukai et al. 1997, Gruhl & Schwaha 2015; Fig. 3.9 A). Underneath the epidermal cells lies a conspicuous ECM that separates these cells from the peritoneal lining of the tentacle coelom. The ECM in gymnolaemates has distinct abfronto-lateral extensions (Smith 1973, Lutaud 1977, Mukai et al. 1997; Fig. 3.9 E). Whether similar structures are present in cyclostomes remains unknown, since only the genus Crisia has been studied on ultrastructural level and details on the shape of the ECM were not noted (Nielsen & Riisgard 1998, Temereva & Kosevich 2018). The ECM also contains the major tentacle neurite bundles. The inner peritoneal lining is composed of few cells that on the frontal and abfrontal side contain the myofibrils of the longitudinal tentacle muscles (Mukai et al. 1997; Fig. 3.9 A, C, E). On the lateral side, a pair of subperitoneal cells is present in all bryozoans (Smith 1973, Lutaud 1977, Gruhl et al. 2009, Shunatova & Tamberg 2019; Fig. 3.9 A, E). These were sometimes also referred to as peritoneal neurite bundles due to the presence of microtubules in these cells (e.g. Gordon 1974, but see also Weber et al. 2014). It should also be noted that a distinct tentacle coelom was not detected in tentacle sections of the cyclostome Crisia (Nielsen & Riisgard 1998). Instead, a central cell was found to be located medially of each tentacle. Likewise, a distinct cavity of the ring canal and tentacles could not be detected (on light microscopical level) in Cinctipora elegans (Schwaha et al. 2018). Most recent data on cyclostome tentacle ultrastructure, however, revealed a tentacle coelom similar to other bryozoans and concluded that the lumen is difficult to detect in retracted specimens (Shunatova & Tamberg 2019). Bryozoan tentacles are in contrast to brachiopods and phoronid tentacles multiciliated, which could be a consequence of miniaturization (Schwaha et al. 2020). The tentacles of the lophophore show a rather uniform ciliation with four different categories of cilia: sparse abfrontal

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cilia of sensory nature (Shunatova & Nielsen 2002), lateral cilia that create the main feeding current, frontal cilia that transport particles within the lophophore toward the mouth opening, and stiff latero-frontal cilia that have seem to have mainly sensory and sieving function (cf. Nielsen & Riisgard 1998, Riisgard & Larsen 2010, Riisgard et al. 2010, Gruhl & Schwaha 2015; Fig. 3.9). In gymnolaemates, the frontal cilia emerge from the median frontal cell and the laterofrontal ones from its adjacent cells. The lateral cilia emerge from the two lateral cells. In phylactolaemates, the arrangement is similar, but with the frontal cilia emerging from several cells. This general construction is present in phylactolaemates and gymnolaemates. Cyclostomes, however, lack frontal cilia. In at least three different genera belonging to different clades (Crisia: Articulata (Nielsen & Riisgard 1998), Cinctipora: Cinctiporidae (Schwaha et al. 2018), Hornera: Cancellata (Batson pers. communication)) the frontal surface of the tentacles produces mucus-like spheres that are released toward the frontal side of the lophophore and apparently are consumed by the bryozoans (Fig. 3.9 D). Thus, it appears that cyclostomes use some kind of mucus entrapment of food particles for feeding. An alternate strategy, in regard to the loss of the otherwise important frontal cilia, is that the long laterofrontal cilia act as sieve and continuous tentacle flicks effectuated by their longitudinal musculature would act in transport of substances toward the mouth opening (Nielsen & Riisgard 1998).

3.9 Digestive system 3.9.1 General features of the digestive tract The digestive system in bryozoans is u-shaped in all bryozoans and very similar among the different clades. It consists of a fore-, mid-, and hindgut (Fig. 3.10). The foregut starts at the lophophoral base with a mouth opening that continues into the pharynx followed by the

3.9 Digestive system 

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esophagus. The pharynx and esophagus are generally differentiated by their ciliation: the pharynx epithelium carries apical cilia that project into the lumen, whereas the esophagus lacks these (Silén 1944b, Mukai et al. 1997). This distinction is somewhat clearer in phylactolaemate bryozoans but is more difficult in cyclostomes and gymnolaemates due to the lack of cilia over most parts of the pharynx. Cyclostomes and gymnolaemates possess a triradiate, myoepithelial pharynx. The lumen is quite narrow in its relaxed condition and is lined by a distinct cuticle (Marcus 1939, Gordon 1975a, Nielsen 2013, Schwaha et al. 2020). Distinct cilia are sparse in this cuticle-lined part of the pharynx, and dense ciliation is only present in the area of the mouth opening. The lower part of the foregut, the esophagus, varies in its length in myolaemates. In cyclostomes, it is generally rather short, whereas gymnolaemates often have a very long esophagus (cf. Schwaha et al. 2011). All bryozoans possess a distinct cardiac valve at the transition of the foregut into the midgut. This valve hinders reflux of food particles during the retraction process (Silén 1944b). The tubular cardia is rather short in phylactolaemates and cyclostomes but can be elongated in gymnolaemates. Notably, ctenostome gymnolaemates often possess a distinct proventriculus or gizzard in the cardiac region. This ranges from a simple cardiac constrictor in e.g. victorellids (e.g. Braem 1951) or nolellid ctenostomes (Schwaha & Wanninger 2018), to a bulbous proventriculus lined by a thick cuticle as in the genus Hislopia (Annandale 1916, Wiebach 1967), or to gizzards with distinct teeth-like structures (e.g. Bowerbankia/ Amathia; Fig. 3.11). Similar gizzard-like modifications of the cardiac region are also sometimes encountered in cyclostome and cheilostome bryozoans (see Markham & Ryland 1987). The caecum or main part of the stomach and midgut represents the largest region of the digestive tract in bryozoans. It is the main area of food digestion and ingested particles stay longest in this part of the digestive tract. The caecum is commonly elongated or bulbous and thus has a large surface area for digestion.

◂ Fig. 3.8: Interzooidal pore complexes of gymnolaemate bryozoans. (A–C) Ctenostomes. (D–F) Cheilostomes. A, Section of a pore plate of Hislopia malayensis showing multiple, large cells forming the cellular plug. B, Section of a young bud connected to the mother zooid (bottom) of Paludicella articulata. C, Section of a lateral wall of Flustrellidra hispida. Note the presence of two pores in the pore plate. D, Section of a small pore of Caberea boryi. F, Section of a small pore of Lanceopora sp. F, Confocal laser scanning microscopy of Cellaria fistulosa. F-actin labeled in the cincture cells (red) and serotonin located in the special cells. Abbreviations: bc – body cavity, cc – cincture cell, cmg – cuticular margin of the pore plate, dg – digestive tract, izs – interzooidal septum, lc – limiting cell, mtm – median transverse muscle, pp – pore plate, sc – special cell, yb – young bud.

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A pylorus or pyloric region joins the caecum on the anal side of the zooid and represents the last part of the midgut. In phylactolaemates, this area is inconspicuous and not recognizable, but in myolaemates, it is distinctly ciliated and important for the mode of digestion (Silén 1944b, Mukai et al. 1997, Schwaha et al. 2020). The hindgut follows the pylorus and consists of a simple intestine that terminates via the anus in the tentacle sheath outside of the tentacle crown. The position of the anus on the tentacle sheath varies. In phylactolaemates and cyclostomes, it generally is located close to the lophophoral base and thus always close to the lophophore in protruded polypides. In gymnolaemates, the anus is often located in the distal area of the tentacle sheath, close to the vestibular wall (see e.g. Alcyonidium d’Hondt, 1983, Hypophorella expansa Ehlers, 1876). The functional significance is not analyzed, but it could be correlated with the separation of fecal pellets from feeding currents, which otherwise is facilitated in different pathways (see McKinney 1997). Concerning particle uptake, particles are transported via tentacle cilia toward the mouth opening where particles or substances are ingested by peristaltic movements of the foregut musculature in phylactolaemates or a rapid expansion by the special pharynx of myolaemates (Silén 1944b). Transport is generally fast in the foregut and food particles are transferred quickly via the cardiac valve toward the first part of the stomach, the cardia. The modes of digestion vary distinctly between the Phylactolaemata and Myolaemata. Phylactolaemates knead food particles up and down the caecum lumen via peristaltic contractions of dense circular muscles, whereas myolaemates use ciliary action of the pylorus to create rotating food masses in the stomach that are only occasionally shifted from the caecum to the cardia during the digestion process (Silén 1944b, Schwaha et al. 2020). Ingested, potential food particles are in direct contact to the stomach epithelium in the kneaders, in contrast to the rotators that barely have any contact with food particles. In addition, the gut musculature (see section 3.12.5) also reflects the divergent feeding modes of kneaders and rotators.

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3.9.2 Cytological features of the digestive epithelium The mouth opening is ciliated in bryozoans and multiple cilia protrude medially into the mouth opening and aid in food transport. Some putative sensory cells are located within the digestive epithelium surrounding the mouth in some bryozoans (Gerwerzhagen 1913, Nielsen 2013, Schwaha pers. observation). These are probably responsible for food particle testing, which can result in food rejection when these are unpalatable (Bullivant 1968). The pharyngeal epithelium is characterized by elongated, narrow cells. In phylactolaemates and gymnolaemates, there are always large vacuoles that fill large parts or even almost the entire epithelial cell. These can be located in the basal part of the cells in phylactolaemates (Fig. 3.12 A–C) and in the apical part in gymnolaemates (Fig. 3.13 C–E). In cyclostomes, these are present, similar to gymnolaemates, at least in Cinctipora elegans in the apical part of the cell. Others cyclostomes seem to lack these vacuoles (Nielsen 2013; Fig. 3.13 A, B), but a larger survey of gut structure is required to confirm this. The vacuoles do not have any secretory product or duct and apparently are turgescent, stabilizing the foregut epithelium. Only in the phylactolaemate Stephanella hina the pharynx sometimes shows numerous small vesicles in the apical portion (Schwaha, pers. observation; Fig. 3.12 C). As myoepithelium, steno- and gymnolaemates show striated sarcomeres at the lateral cell membranes on the epithelium, enabling the rapid contraction used for suction feeding. An interesting, but unexplored, aspect is whether the presence of large vacuoles yields a more efficient suction pump in gymnolaemates compared to cyclostomes that lack these vacuoles. The esophagus is in respect to its cellular composition inconspicuous, probably since it is merely a transporting tube. In the phylactolaemate esophagus, large vacuoles are present that are in contrast to the pharynx located in the apical part of the cells. The cyclostome esophagus is similar to the pharynx and the gymnolaemate one is

◂ Fig. 3.9: Tentacle structure in bryozoans. (A, C, E) Schematic cross-sections of Phylactolaemata (A), Cyclostomata (C), and Gymnolaemata (E). A is redrawn and modified from Gruhl and Schwaha (2015), C is based on data from Nielsen and Riisgard (1998) and Temereva and Kosevich (2018), and E is modified and redrawn from Lutaud (1977). The frontal side shows up and the abfrontal down on each scheme. Muscles in the peritoneal lining of the tentacles are indicated in red. (B, D, F) Light microscopical cross-sections of tentacles of the phylactolaemate Cristatella mucedo (B), the cyclostome Cinctipora elegans, (D) and the gymnolaemate Paludicella articulata (F). Abbreviations: af – abfrontal neurite bundle, afc – abfrontal cell, afs – abfrontal sensillum, fc – frontal cilia, ftc – frontal cells, lac – lateral cell, laf – lateroabfrontal neurite bundle, lc – lateral cilia, lfc – laterofrontal cilium, lfn – laterofrontal neurite bundle, lfr – laterofrontal cell, mfn – mediofrontal neurite bundle, muc – mucus secretions, spc – subperitoneal cell, tc – tentacle coelom, tpe – tentacle peritoneum, ts – tentacle sheath.

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Fig. 3.10: Schematic drawing of the gut of bryozoans. Redrawn and modified from Silén (1944b). (A) Phylactolaemata. (B) Cyclostomata. (C) Some ctenostome Gymnolaemata. (D) Gymnolaemata. Abbreviations: a – anus, ca – cardia, cae – caecum, es – esophagus, int – intestine, mo – mouth opening, ph – pharynx, py – pylorus.

generally characterized by its lack of striated sarcomeres and also lack of distinct vacuoles. Little information is available for the cellular diversity of the stomach of bryozoans. Most information is based on old histological analyses and some feeding experiments, which showed that digestion is intra- and extracellular (summarized in Mukai et al. 1997). Ultrastructural details of the stomach are restricted to two phylactolaemates (Malchow 1978, Mukai et al. 1997) and one cheilostome (Gordon 1975a). The stomach is characterized by two principal cell types, the acidophil and basophil type. These differ in their particular histological, staining properties and their cellular contents. Basophil cells appear darker in histological semithin sections. Especially caecal cells often include various cellular inclusions that range from lipid droplets, zymogen-like vesicles and other, variable inclusions (cf. Mukai et al. 1997). Figure 3.14 gives a glimpse in the diversity of caecal structure, but detailed studies focusing on the digestion process including

cellular dynamics of uptake and/or release of substance are urgently required.

3.10 Funicular system In its simplest form, the funicular system comprises a simple peritoneal strand that extends from the proximal tip of the caecum to the body wall. Such a situation is found in phylactolaemate bryozoans (Wood 2014; Figs. 3.1 and 3.2). The funiculus in the latter can vary distinctly in length and always has longitudinal muscles of unknown function in its lining (Schwaha & Wanninger 2012). This simple cord is important in phylactolaemates as it is the site of statoblast (dormant stage) formation (Fig. 3.15 A and B) and also the area where the testes develop in most species (Wood 2014). The funiculus is simple and short in cyclostomes. Topologically and structurally, it is similar to the

▸ Fig. 3.11: Cardiac modifications in ctenostome Gymnolaemata. (A–D) Semithin sections. (E–F) Confocal laser scanning microscopy. A, Proventriculus of Hislopia malayensis showing a prominent cuticular inner lining and massive muscles surrounding the cardia. B, Amathia semiconvoluta showing the gizzard with hard teeth. C, Cardiac constrictor of Victorella pavida. D, Gizzard of Amathia verticillata showing numerous teeth. E, Muscles of the gizzard of Amathia verticillata. F, Cardiac constrictor of Mimosella verticillata. Abbreviations: bc – body cavity, ca – cardia, cae – caecum, cds – cardiac constrictor, cw – cystid wall, fgm – foregut musculature, gml – muscular layer of the gizzard, gzt – gizzard teeth, lb – lophophoral base, pcu – cuticular lining of the proventriculus, pml – muscular layer of the proventriculus, pro – proventriculus, rm – retractor muscles.



3.10 Funicular system 

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Fig. 3.12: Digestive tract of bryozoans. (A) Cross-section of the esophagus and intestine of the phylactolaemate Plumatella sp. (B) Longitudinal section of the foregut of the phylactolaemate Hyalinella punctata. (C) Longitudinal section of the foregut of the phylactolaemate Stephanella hina. (D) Section of the midgut of the cyclostome Patinella radiata. (E) Section of the pylorus and intestine of the cheilostome Lanceopora sp. Abbreviations: a – anus, bw – body wall, ca – cardia, cae – caecum, cg – cerebral ganglion, cv – cardiac valve, es – esophagus, int – intestine, ph – pharynx, py – pylorus, rm – retractor muscles, ts – tentacle sheath.



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Fig. 3.13: Digestive tract of bryozoans. (A) Longitudinal section of the gut of the cyclostome Crisia sp. (B) Longitudinal section of the foregut of the cyclostome Patinella radiata. (C) Longitudinal section of the foregut of the cheilostome Securiflustra securifrons. (D) Longitudinal section of the foregut of the ctenostome Arachnidium fibrosum. (E) Longitudinal section of the gut of the cheilostome Lanceopora sp. Abbreviations: ca – cardia, cae – caecum, cg – cerebral ganglion, cv – cardiac valve, ec – ectocyst, es – esophagus, int – intestine, l – lophophore, lb – lophophoral base, mo – mouth opening, ph – pharynx, py – pylorus, rm – retractor muscles, t – tentacles, te – testes.

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Fig. 3.14: Structure of the caecum in bryozoans. All section images showing different cellular inclusions in the cells of the caecum. (A) Phylactolaemate Hyalinella punctata. (B) Phylactolaemate Cristatella mucedo. (C) Cyclostome Hornera sp. (D) Cyclostome Crisia sp. (E) Ctenostome Hislopia malayensis. (F) Cheilostome Securiflustra securifrons. (G) Ctenostome Alcyonidium gelatinosum. (H) Cheilostome Cellaria fistulosa. Abbreviations: ec – ectocyst, gl – gut lumen, lb – lophophore base, nml – neuro-muscular layer surrounding the distal tip of the caecum, ppc – pseudopore cell, rm – retractor muscle.



phylactolaemate funiculus and possesses longitudinal muscles (Figs. 3.3 and 3.15 C). As mentioned above, it is one of the few areas where the membranous sac attaches to the cystid wall and thus most likely acts in fastening the polypide to the skeletal wall. Gonads do not appear to be distinctly associated with the funiculus in this clade (Boardman 1998). Gymnolaemate funicular systems appear to be more diverse and complex than previously understood and remain an important area of future research (Schwaha et al. 2020). Ctenostomes show a large variety of funicular systems that range from simple strands as observed in phylactolaemate and cyclostome zooids to strands that interconnect zooids within the colony and act as colonial system of integration (CSI). Superfamilies that have rather flat, dense colonial forms such as the Hislopioidea or Alcyonidioidea seem to have only simple funicular cords that resemble the phylactolaemate or cyclostome condition. In this case, it is a simple, often muscular, strand that extends from the caecum to the body wall (Fig. 3.15 D, E, G, I). As far as analyzed, many gymnolaemates have a second funicular strand in parallel to the first one – from the pyloric region to the body wall (Fig. 3.15 D, G). It seems that either one or both of these may be present in ctenostomes. The presence of both of these funicular cords has first been demonstrated in Paludicella articulata (Allman 1856), which is a representative of the superfamily Paludicelloidea. The general ctenostome bauplan, including its funicular system that is depicted in most textbooks, generally refers to the vesicularioidean ctenostome Bowerbankia/ Amathia, which has a distinct proximal funicular cord from the proximal caecum tip toward the pore plate (e.g. Bobin 1977, Reed 1991). Vesicularioideans are stolonate (with kenozooidal polymorphs, see chapter 10) and derived forms of ctenostome bryozoans (Jebram 1973). This indicates that this particular type of commonly depicted funicular system is probably not the ancestral condition. The other stolonate form of ctenostomes, the Walkerioidea, lack any distinct interconnecting funicular cords (Jebram 1973, Schwaha & Wanninger 2018, Schwaha et al. 2020). In other ctenostomes, the funicular system is less described: in arachnidioideans, the funicular system was generally described as absent (Jebram 1973, 1986). However, in few nolellid species, an interconnecting cord has been described (Calvet 1900). Many interpretations of interconnecting tissue strands are also based on artificial shrinkage of the epidermis away from the ectocyst, which gives the impression of thin funicular cords located within (Schwaha, pers. observation). This

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is not surprising given the thin size of the epidermis and lack of a peritoneal lining (see section 3.6 and chapter 10). Descriptions of many other species have not been properly reinvestigated, which calls for the necessity of its study for the future. While there is significant lack of knowledge on this organ system, the most important thing to notice is that not all ctenostomes share interconnecting cords. In the latter case, as present in vesicularioideans, distinct transport of nutrients could be demonstrated (Bobin 1977). It is possible that a complex CSI integrating zooids evolved multiple times in ctenostomes (Schwaha et al. 2020). The funicular system of cheilostomes is generally depicted and described as complicated network of interconnecting strands that connect the polypide with each communication pore (Fig. 3.15 F). It has been shown in some autoradiographic studies to be employed in substance exchange between zooids (Best & Thorpe 1985). The complex branching CSI of cheilostomes was considered a possible apomorphy of this clade enabling its large success and extensive forms of polymorphism (Schwaha et al. 2020). The latter indicates that an elaborate system of polymorph-nourishing system has to be present. Likewise, the funicular system plays a vital role in placental cheilostomes, which evolved multiple times (Ostrovsky 2013). However, it should be stressed that the funicular system of probably less than 10 of the 5.000+ cheilostomes species was actually studied in detail.

3.11 Nervous system The nervous system has recently been summarized by Gruhl and Schwaha (2015). Past this review, additional important contributions have been provided in this field (Shunkina et al. 2015, Temereva & Kosevich 2016, Ambros et al. 2018, Schwaha et al. 2018, Worsaae et al. 2020). Historical aspects of the study of the bryozoan nervous system are found in Gruhl and Schwaha (2015) or Shunkina et al. (2015). A cerebral ganglion (or brain) is the center of the nervous system in all bryozoans (Fig. 3.16). It lies adjacent to the pharyngeal wall and during the budding process develops via an invagination of the inner budding layer in the prospective pharyngeal area (cf. Nielsen 1971, Schwaha et al. 2011). As an invagination of an epithelial layer, the ganglion originally contains a small lumen. The latter is retained in all phylactolaemates (Fig. 3.16 A) but lost in most myolaemates. Only few ctenostome

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species possessing a small lumen within the cerebral ganglion were encountered (Paludicella articulata Weber et al., 2014, Amathia gracilis Temereva & Kosevich, 2016; see also Fig. 3.16 D). Recent data on the cyclostome Crisia eburnea also indicate that a lumen is present (Temereva & Kosevich 2018). The presence of the lumen in a cyclostome and some ctenostomes indicate that the lumen was present in the ancestral myolaemate bryozoan and independently lost among the different clades. Still, ontogenetic analyses are required to assess whether the adult lumen forms de novo or indeed, as in phylactolaemates, represents the lumen of the initial vesicle anlage of the ganglion during budding. Additional ganglionic structures were described in form of lateral ganglia on each lateral side of the cerebral one in the cyclostome Cinctipora elegans (Schwaha et al. 2018) and an unpaired visceral ganglion proximally adjacent to the cerebral one in malacostegine cheilostomes (Lutaud 1977). In association with the horseshoe-shaped lophophore, phylactolaemates possess two extensions of the ganglion into disto-anal direction along the lophophoral arms (Fig. 3.5 A). These are termed ganglionic horns, contain a small lumen confluent with the main ganglionic lumen, and innervate the tentacles of the lophophoral arms (Shunkina et al. 2015, Gruhl & Schwaha 2015). In all bryozoans, a circum-oral nerve ring (CON) emanates from both lateral sides of the ganglion (Figs. 3.17 A, C, E and 3.18 C, E–G, I). It extends from the ganglion around the pharynx/mouth region and forms a complete ring on the oral side in almost all species studies. Next to the ganglion itself, the CON is the main origin of the tentacular neurite bundles (Figs. 3.17 and 3.18). In addition to the tentacular neurite bundles, two distinct neurite bundles extend from the cerebral ganglion in all bryozoans: the tentacle sheath neurite bundles, which also extend into the apertural area and body wall, and the visceral neurite bundles (Figs. 3.17 A, C, E and 3.18).

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In cyclostomes and gymnolaemates, a second ring nerve is located proximally of the circum-oral nerve ring (Temereva & Kosevich 2016, 2018). This so-called outer nerve ring is sometimes incomplete and the complete traverse on the oral side was not found in all species (cf. Schwaha et al. 2018; Figs. 3.17 A, C, E and 3.18). In some gymnolaemates, only the root of this nerve ring from the proximal side of the ganglion is present. In this case, it forms the so-called trifid nerve (Lutaud 1977, Schwaha et al. 2018; Fig. 3.17 C).

3.11.1 Tentacle innervation In each tentacle, four to six epidermal and sometimes two additional peritoneal neurite bundles have been described (Gruhl & Schwaha 2015). In particular, the subperitoneal neurite bundles remain enigmatic because they have been addressed as nerves (Gordon 1974), subperitoneal cell, or enclosed peritoneal cells (see section 3.8.2 and also Fig. 3.9). Only few studies employing immunocytochemical methods or electron microscopy were able to visualize or see such inner peritoneal neurite bundles (Temereva & Kosevich 2016, 2018, Schwaha et al. 2018).

3.11.1.1 Phylactolaemata Phylactolaemates have a set of three frontal neurite bundles, a mediofrontal and two laterofrontal bundles. The abfrontal side shows an identical situation with a medioabfrontal and two lateroabfrontal bundles (Shunkina et al. 2015, Ambros et al. 2018; Fig. 3.9). All tentacular neurite bundles have an intertentacular origin from so-called radial nerves in phylactolaemates (Fig. 3.17 D). These emerge directly from the cerebral ganglion, the CON, or the ganglionic horns. In their distal traverse, the radial nerves branch off several neurite bundles of different sizes in their proximo-distal traverse. Most proximally

◂ Fig. 3.15: Funicular system of bryozoans. (A, B) Phylactolaemata. (C) Cyclostomata. (D, E) Ctenostome Gymnolaemata. (F–I) Cheilostomata. A, Whole mount of Hyalinella punctata showing a thin funicular cord from the proximal caecum end. B, Section of a young statoblast anlage in the funicular cord of Hyalinella punctata. C, Elongated funiculus of Patinella radiata. D, F-actin staining of a retracted zooid of Mimosella verticillata showing the two funicular muscles at the caecum. E, Funicular muscle at the proximal side of the caecum in Alcyonidium gelatinosum. F, Schematic drawing of the funicular network of the cheilostome Electra pilosa. Redrawn from Lutaud (1983). G, Whole mount of Electra pilosa, stained for cell nuclei, showing the two funicular muscles at the caecum. H, Longitudinal section of Securiflustra securifrons showing the interzooidal pore plate and interconnecting funicular strands. I, F-actin staining of a zooid of Beania mirabilis showing the funicular muscle and some strands in connection with the multiporous pore-plate. Abbreviations: cae – caecum, cw – cystid wall, db – duplicature band, ds – diaphragmatic sphincter, f – funiculus, fm – funicular muscle, fun – funicular network, int – intestine, l – lophophore, lb – lophophoral base, o – orifice, ooc – operculum occlusor, op – operculum, p – polypide, pdm – parieto-diaphragmatic muscles, ph – pharynx, pm – parietal muscles, pp – pore-plate, pvm – parieto-vestibular muscles, py – pylorus, rel – retracted lophophore, rm – retractor muscle, sba – statoblast anlage, tm – tentacle muscles, ts – tentacle sheath, v – vestibulum, vw – vestibular wall.

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Fig. 3.16: Histological sections of the cerebral ganglion at the lophophoral base. (A) Longitudinal section of the cerebral ganglion of the phylactolaemate Lophopus crystallinus. Note the typical croissant-shape of the ganglionic cells and the orally directed ganglionic lumen. (B) Longitudinal section of the foregut and adjacent small ganglion of the cyclostome Patinella radiata. (C) Cross-section of the lophophoral base of the ctenostome Hislopia malayensis. (D) Cross-section of the cerebral ganglion of the ctenostome Paludicella articulata. Abbreviations: a – anus, bc – body cavity, bd – buccal dilatator, ca – cardia, cae – caecum, cg – cerebral ganglion, cgc – central ganglion cell, con – circum-oral nerve ring, cv – cardiac valve, ec – ectocyst, ep – epistome, es – esophagus, gl – ganglion lumen, int – intestine, itp – intertentacular pits, lb – lophophoral base, mo – mouth opening, ph – pharynx, rc – ring canal, rm – retractor muscle, ts – tentacle sheath.



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Fig. 3.17: Schematic overview of the main elements of the nervous system of bryozoans. A–C are schemes of the lophophoral base viewed from the anal side. D–F are lateral views of the main branches of tentacle innervation. (A, D) Phylactolaemata. (B, E) Cyclostomata. (C, F) Gymnolaemata. Based on the few descriptions, two different origins of the mediofrontal neurite bundle are displayed in the cyclostome scheme in E. The median and left have two rootlets and the right shows a single root emerging from the circum-oral nerve ring. Abbreviations: afn – abfrontal tentacle nerve, arr – abfrontal nerve roots, cg – cerebral ganglion, con – circum-oral nerve ring, cts – compound tentacle sheath neurite bundle, fn – mediofrontal nerve, fnr – frontal nerve roots, gah – ganglionic horn, gl – ganglion lumen, itm – intertentacular membrane, itp – intertentacular pit, laf – lateroabfrontral nerve, lfn – laterofrontal nerve, onr – outer nerve ring, rn – radial nerve, trf – trifid nerve, tsn – tentacle sheath neurite bundles, tsp – tentacle sheath plexus, vcn – visceral neurite bundles, vp – visceral plexus.

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Fig. 3.18: Nervous system of the lophophoral base, anti-acetylated alpha tubulin stainings. (A) General overview of an extended lophophore of the phylactolaemate Hyalinella punctata. (B) Detail of the cerebral ganglion and main emanating neurite bundles of Hyalinella punctata. (C) Retracted tentacle crown of the cyclostome Cinctipora elegans. (D, E) Ctenostome Paludicella articulata. D, View of the anal side showing the cerebral ganglion. E, View from the oral side showing the oral connection of the circum-oral nerve ring. (F) Lophophoral base of the cyclostome Cinctipora elegans. (G) Lophophoral base of the ctenostome Mimosella verticillata. Asterisks mark a branching of the tentacle innervating neurite bundle emanating from the cerebral ganglion. (H) Cerebral ganglion of the cheilostome Electra posidoniae. (I) Lateral view of the lophophoral base of the cheilostome Margaretta cereoides. Abbreviations: a – anus, api – apertural innervation, cg – cerebral ganglion, con – circum-oral nerve ring, gah – ganglionic horn, ilc – intestinal cilia, l – lophophore, lag – lateral ganglion, o – orifice, onr – outer nerve ring, ots – oral tentacle sheath neurite bundle, pan – parietal neurite bundle, ph – pharynx, php – pharyngeal plexus, pmn – pharyngeal main neurite bundles, rn – radial nerve, tni – tentacle innervation, trf – trifid nerve, tsn – tentacle sheath neurite bundle, tsp – tentacle sheath plexus, vcn – visceral neurite bundles.



a series of thin bundles branch toward the frontal side of a tentacle. Medially, they fuse with the opposite branches from the neighboring radial nerves to form the mediofrontal neurite bundle. On the terminal, distal end, the radial nerve splits into four branches, two diverging frontally and two abfrontally. The frontally emanating branches become the laterofrontal neurite bundles, whereas the abfrontal ones represent the roots of the latero- and medio-abfrontal neurite bundles. Shortly after their emergence from the radial nerve, a distinct neurite bundle branches off medially from these roots to form the medio­ abfrontal neurite bundle. The remaining bundle of the roots form the lateroabfrontal neurite bundle (Fig. 3.17 D). Two additional bundles emerge from the radial nerves that innervate the lophophoral base and intertentacular membrane, the “additional radial nerve” and the “basal radial nerve” (Shunkina et al. 2015). In the traverse of the basal radial nerve, distinct intertentacular perikarya are present between each pair of tentacles (Ambros et al. 2018; see also Fig. 3.19 H).

3.11.1.2 Myolaemata Cyclostomata Only Crisia and Cinctipora were so far studied concerning the innervation pattern of the tentacles. Tentacle neurite bundles range from four to six, with the latter appearing to be the more accurate innervation pattern similar to phylactolaemates. It consists of three frontal and three abfrontal neurite bundles. The laterofrontal and both abfrontal neurite bundles emerge from an intertentacular radial nerve (also termed intertentacular fork) that emerges from the cerebral ganglion or the CON. At the distal tip of the radial nerve, two pairs of thin neurite bundles emanate laterally to adjacent tentacles – one pair frontally and the second abfrontally. As in phylactolaemates, the frontal branch forms the laterofrontal neurite bundle and the abfrontal one is the root for the lateroabfrontal and medioabfrontal neurite bundle. The origin of the mediofrontal neurite bundle differs and originates either directly from the CON or ganglion or via two roots emerging from the latter two structures (Schwaha et al. 2018, Temereva & Kosevich 2018; see also Fig. 3.17 E). Basal intertentacular perikarya were described in Cinctipora elegans (Schwaha et al. 2018). These strongly resemble the perikarya also found in phylactolaemates. Gymnolaemata Gymnolaemates have similar tentacle neurite bundles as other bryozoans but lack the lateroabfrontal neurite bundles. Thus, only four are present in each tentacle

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(Fig. 3.9). The branching of the tentacle neurite bundles is similar to cyclostomes with the laterofrontal neurite bundle emerging from the intertentacular radial nerve (intertentacular fork) to split into adjacent tentacles again. The mediofrontal neurite bundles emerge directly from the ganglion or CON in most analyzed species (Schwaha et al. 2011, Weber et al. 2014; see also Fig. 3.17 F). Some species also have two very short rootlets that fuse in the median tentacle plane (Pröts et al. 2019), similar as in Crisia (Temereva & Kosevich 2018). The abfrontal neurite bundle in ctenostomes emerges as in all other bryozoans from the intertentacular fork and in cheilostomes has generally been described as directly emerging from the ganglion or CON in the tentacle plane (Lutaud 1977, 1993). Newer data on the cheilostome nervous system indicate that the abfrontal bundles also emerge from an intertentacular position (Prömer et al., 2019 Fig. 3.18 I). However, there seems to be the tendency of reducing one of the roots of the abfrontal neurite bundle to a single, asymmetric one. Some ctenostomes also tend to have an asymmetric arrangement of such nerves. Superficially, this resembles a single neurite bundle emerging as previously interpreted (cf. Lutaud 1977). As in the other two clades, intertentacular perikarya are present in gymnolaemates, too. These are located in the intertentacular pits (Schwaha & Wanninger 2015; see also Fig. 3.19 F, G). Probably these intertentacular perikarya are homologous among the different clades, perhaps even synapomorphic for bryozoans.

3.11.2 Peripheral innervation 3.11.2.1 Tentacle sheath, apertural, and body wall innervation Phylactolaemata The tentacle sheath of phylactolaemates is innervated by a diffuse plexus that orginates from two distinct concentrated neurite bundles that emanate from the cerebral ganglion (Figs. 3.17 A and 3.18 A). The following description is for retraced zooids, i.e. when the tentacle sheath is wrapped around the retracted lophophore. Note that distal and proximal directions vary according to retracted vs. protruded state. One of the neurite bundles forming the plexus emerges directly above the ganglion and extends into distal direction as thick neurite bundles. In their traverse, they branch off thinner bundles that form part of the plexus. A second, prominent neurite bundle emerges on each lateral side of the ganglion and passes circumorally around the foregut and into the basal, proximal wall of the orally situated ring canal (Fig. 3.18 B). From the latter,

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Fig. 3.19: Selected aspects of the nervous system of bryozoans. (A) General overview of the peripheral innervation of the cheilostome Electra pilosa. Redrawn and modified from Lutaud (1977). The neurite bundles displayed in blue and dashed are sensory and originate from the direct nerve of the ganglion. The red are motor nerves emanating from a branch of the trifid nerve. (B) Overview of the cheilostome Electra posidoniae. (C) Detail of the apertural area of E. posidoniae. (D) Apertural innervation of the ctenostome Paludicella articulata. (E) Sensory cells in the pharyngeal epithelium of the ctenostome Paludicella articulata. (F–H) Intertentacular perikarya at the lophophoral base: F, Ctenostome Paludicella articulata; G, Cheilostome Margaretta cereoides; H, Phylactolaemate Stephanella hina. Abbreviations: con – circum-oral nerve ring, ctn – compound tentacle sheath neurite bundle, dbn – duplicature band neurite bundles, dn – direct nerve, ds – diaphragmatic sphincter, dsi – diaphragmatic sphincter innervation, ip – intertentacular perikarya, itp – intertentacular pits, lb – lophophoral base, op – operculum, pan – parietal innervation, phc – pharyngeal cilia, pp – peripharyngeal plexus, rm – retractor muscles, rn – radial nerves, rtl – retracted lophophore, sc – sensory cells, tf1 – one of the major branches of the trifid nerve, tni – tentacle innervation, tsn – tentacle sheath neurite bundle, vcn – visceral neurite bundles.

the neurite bundles proceed distally into the tentacle sheath. Along the oral traverse of these bundles, several thin neurite bundles separate from the thick bundle and proceed into the tentacle sheath to form part of the plexus (Ambros et al. 2018).

Distally toward the orificial or apertural area, neurite bundles of the tentacle sheath plexus branch off into each duplicature band and further into the body wall. In addition, neurite bundles of the tentacle sheath proceed into the vestibular wall. In the latter, predominantly

3.11 Nervous system 



longitudinal neurite bundles are present that also enter the body wall. The body or cystid wall in phylactolaemates has a diffuse nerve plexus (Ambros et al. 2018).

Myolaemata A diffuse plexus innervating the tentacle sheath as found in phylactolaemates is not present in myolaemates. Instead, a pair of anally situated prominent neurite bundles innervates the tentacle sheath and proceeds distally toward the apertural area (Figs. 3.17 B, C and 3.18 C, D, H). In the analyzed cyclostomes, C. elegans and Crisia eburnea, these paired bundles originate directly from the distal area of the cerebral ganglion (Schwaha et al. 2018, Temereva & Kosevich 2018). In gymnolaemates, a second neurite bundle adjoins this direct neurite to form the “greater tentacle sheath nerve” as designated by Lutaud (1973b) or as termed here as “compound tentacle sheath neurite bundle” (see also Mukai et al. 1997; Fig. 3.19 A). This second branch originates from the root of the trifid nerve, which emerges on the lateral sides of the ganglion and splits into three branches. One of these three branches bends distally (again, in retracted zooids) toward the tentacle sheath and after a short distance joins the direct tentacle sheath neurite bundle to form the compound tentacle sheath neurite bundle (Fig. 3.19 A). In cyclostomes, one or two bifurcations may be present in the traverse of the tentacle sheath neurite bundle toward the apertural area. In gymnolaemates, distinct bifurcations are not described, but since each of the four duplicature bands is innervated by one neurite bundle, an additional branching has to take place in the direction of the apertural area. The innervation of the apertural area and body wall is only little investigated in cyclostomes (cf. Schwaha et al. 2018, Temereva & Kosevich 2018, Worsaae et al. 2020). In Cinctipora elegans, neurite bundles are radially arranged in the attachment organ and the vestibular wall. No distinct neurite bundles were recognized beyond the apertural innervation and thus not in the cystid wall. The lining of the original peritoneum, the membranous sac, shows several fine, circular neurite bundles. In Crisia, the apertural area mainly shows a strong neurite bundle in the diaphragmatic sphincter and vestibular wall. The body wall shows thick longitudinal neurite bundles (Temereva & Kosevich 2018). These have no distinct comparable structure in phylactolaemates or gymnolaemates. It seems that the structure of the body wall, free-walled as in Cinctipora vs. fixed-walled as in Crisia, might be reflected in the innervation of the body wall. A diffuse body wall plexus as

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described for Crisia (Lutaud 1979b) has not been found by newer methods. In gymnolaemates, the apertural area is innervated by extensions of the compound tentacle sheath neurite bundle (Fig. 3.19 A–D). These extend along the anal side of the tentacle sheath into the diaphragmatic sphincter and further into the vestibular wall. Distinct neurite bundles also extend from the tentacle sheath innervation into each duplicature band. The innervation of the aperture is only investigated in the cheilostome Electra in more detail. The innervation pattern strongly correlates to muscular structures such as the diaphragmatic sphincter (Lutaud 1977; Fig. 3.19 B, C). Thin neurite bundles pass along each duplicature band from the tentacle sheath toward the body wall. Parietal neurite bundles extend from the frontally inserted duplicature bands and run proximally toward the frontal attachment sites of the parietal muscles (Fig. 3.19 A, B, D). The bundles meander around the circular to elliptical muscle insertion areas and proceed toward the most proximal-most muscle bundle (Lutaud 1977, Gruhl & Schwaha 2015, Schwaha & Wanninger 2015). Additional innervation such as the general Hiller’s plexus (see Hiller 1939, Lutaud 1969) was not recently found by immunocytochemical methods.

3.11.2.2 Visceral innervation Phylactolaemata The foregut is innervated by a regular and dense plexus that emerges from the cerebral ganglion and also from the CON. On the anal side of the foregut, a pair of thicker neurite bundles are often present (Figs. 3.17 A and 3.18 B). The foregut plexus extends over the pharynx and esophagus with almost all bundles terminating in the cardiac valve. Several neurites or neurite bundles branch medially from the plexus to innervate special sensory cells wedged into the epithelium of the foregut (Ambros et al. 2018). Only few, mostly longitudinal neurite bundles extend further into the cardia and partially into the distal area of the caecum. The digestive epithelium of the caecum is bordered by a distinct neuromuscular layer that shows prominent circular musculature associated with a diffuse layer of neuronal fibers (Fig. 3.14 A). A connection of this layer to the remaining nervous system has so far not been observed. The intestine shows only little innervation consisting of few circular neurite bundles adjacent to the circular musculature of the intestine and few sensory branches that extend into the intestinal epithelium similar as observed in the foregut. These probably represent anal density receptors that register whether the intestine is

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filled with a fecal pellet, which will require subsequent removal (Ambros et al. 2018). Myolaemata The visceral innervation in this clade is mainly present as few longitudinal neurite bundles in the foregut. In cyclostomes, there are anally situated visceral neurite bundles and lateral ones, both emerging from the cerebral ganglion (and the lateral ganglia where present). These terminate in the area of the cardia. The remaining digestive tract shows no distinct neurite bundles (at least with immunocytochemical methods, see Schwaha et al. 2018, Temereva & Kosevich 2018, Worsaae et al. 2020). In gymnolaemates, the visceral innervation is more constricted to the anal side of the foregut and does not exceed as laterally as in cyclostomes. Up to three types of visceral neurite bundle types are differentiated: mediovisceral, mediolateral, and lateral neurite bundles (e.g. Weber et al. 2014, Temereva & Kosevich 2016; see also Figs. 3.17 B, C and 3.18 D, H). In both cyclostomes and gymnolaemates, few bundles also branch from the main visceral neurite bundles into sensory cells embedded in the epithelium of the foregut similar as in phylactolaemates (Nielsen 2013; Fig. 3.19 E). Likewise, a peripharyngeal plexus can also be found in the distal area of the pharynx, close to the mouth opening, in both clades (Weber et al. 2014, Temereva & Kosevich 2018).

3.12 Muscular system The following data are a summary from previous studies of Schwaha et al. (2011, 2018), Schwaha and Wanninger (2012, 2018), Gawin et al. (2017), and unpublished observations. Bryozoan muscle systems can be subdivided into six different functional areas or systems (Schwaha & Wanninger 2018): 1) muscles associated with the body wall, which are also essential for the protrusion process of bryozoans; 2) apertural muscles; 3) lophophoral muscles; 4) tentacle sheath muscles; 5) digestive tract muscles; and 6) retractor muscles.

3.12.1 Muscles associated with the body wall Only phylactolaemate bryozoans show regular body wall musculature. In most analyzed phylactolaemates, it is an orthogonal grid of longitudinal and circular muscles embedded in the basal ECM of the epidermis and peritoneum, with the circular muscles being associated with the epidermis and the longitudinal one to the peritoneum. In

the phylactolaemates Lophopus crystallinus and Pectinatella magnifica, additional diagonal muscles can occur in certain body parts as a third layer (Gawin et al. 2017). In the Myolaemata, the regular body wall musculature is modified and never occurs as regular body wall musculature as in phylactolaemates. In cyclostomes (and most likely all Stenolaemata), the body wall muscles are reduced to circular muscles associated with the peritoneum (Fig. 3.3). These are remains of the original body wall muscles and are called annular muscles that extend from the proximal tip of the membranous sac toward its distal tip slightly below the apertural area. Some diagonal fibers have been detected, too (Schwaha et al. 2018). Gymnolaemates reduced the original body wall musculature to a series of transverse muscle bundles that directly cross the body cavity from the basal or lateral side toward the frontal side. These so-called parietal muscles are characteristic for all gymnolaemates and occur either as a continuous series of thin bundles over a large area of the zooid or concentrated to distinct, serial bundles (Jebram 1986, Schwaha & Wanninger 2018; Figs. 3.4 and 3.20 B, C). Particularly the formation of cryptocystal shields in anascan cheilostomes can distinctly modify the arrangement of these muscles, too (see also Cheetham & Cook 1983, Perez & Banta 1996, Banta et al. 1997, and chapter 11). Compression of these different muscles enable the protrusion process of zooids. While the arrangement of these muscles varies among the different clades, the mechanism to increase pressure in the body cavity and thus exert force on the retracted polypide remains the same.

3.12.2 Apertural muscles of bryozoans These are associated with the apertural or orificial area and in association with the polypide protrusion and retraction are important for opening and closing this area. Generally, bryozoans possess two different muscular systems associated with this area: the first are muscles at the base of the epithelial lining of the vestibular wall followed by a diaphragmatic sphincter that in retracted zooids separates the distal cavity, the vestibulum, from the proximal cavity formed by the inverted tentacle sheath, the atrium. The second group constitutes muscles that traverse the body cavity laterally of the apertural area and extend between the body wall and either the vestibular wall, diaphragm, or tentacle sheath (Schwaha et al. 2011; Fig. 3.21). The diaphragmatic sphincter is present in all bryozoans and homologous in all groups – in cyclostomes, it is generally referred to as atrial sphincter (Borg 1926,



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Fig. 3.20: Selected aspects of the muscular system of bryozoans. (A) Two retracted zooids of the cyclostome Cinctipora elegans showing the sparse digestive tract musculature. (B) Retracted zooid of the cheilostome Fenestrulina malusii. (C) Retracted zooid of the ctenostome Hislopia malayensis. (D) Retracted zooid of the cyclostome Crisia sp. (E) Caecum of the phylactolaemate Cristatella mucedo showing the dense arrangement of circular muscles. (F) Retractor muscle of the ctenostome Triticella flava showing distinct striations. (G) Retractor muscle of the ctenostome Cryptopolyzoon wilsoni showing smooth muscle fibers. Abbreviations: anm – annular muscles, apm – apertural muscles, cae – caecum, db – duplicature bands, ds – diaphragmatic sphincter, fm – funicular muscle, int – intestine, lb – lophophoral base, ooc – operculum occlusor, pdm – parieto-diaphragmatic muscle, ph – pharynx, pm – parietal muscles, pp – pore plate, pv – proventriculus, rm – retractor muscles, tm – tentacle muscles, vm – vestibular muscles.

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Fig. 3.21: Schematic drawing of the organization of apertural muscles in bryozoans. Modified from Schwaha et al. (2011). (A, B) Phylactolaemata. (C, D) Cyclostomata. (E–H) Ctenostome Gymnolaemata. (I) Cheilostomata. Abbreviations: a – atrium, ao – attachment organ with ligaments, db – duplicature bands, ds – diaphragmatic sphincter, o – orifice, ocl – operculum occlusor, op – operculum, pdi – parieto-diaphragmatic muscles, pve – parieto-vestibular muscles, ts – tentacle sheath, v – vestibulum, vd – vestibular dilatators, vm – vestibular muscles.

Nielsen & Pedersen 1979, Schwaha et al. 2018). Muscles in the vestibular wall are most prominent in phylactolaemates (Schwaha & Wanninger 2012, Gawin et al. 2017), very few in cyclostomes (Schwaha et al. 2018), and present in some gymnolaemates (predominantly ctenostomes) (Schwaha et al. 2011, Schwaha & Wanninger 2018, Schwaha, pers. observation). In phylactolaemates, the regular mesh of body wall muscles continues directly into the musculature of the vestibular wall at the distal tip of the cystid. Consequently, it also consists of a regular mesh of circular and longitudinal muscle fibers with the latter being more prominent. At the transition from the body wall into the vestibular wall, the circular muscles are

densely arranged and form the orificial sphincter. In ctenostome gymnolaemates, the musculature in the vestibular wall sometimes shows a similar arrangement. However, an orifical sphincter is lacking in most ctenostomes and is present only in few species (Schwaha & Wanninger 2018). Some ctenostomes also possess diagonal muscles in the vestibular wall (Schwaha et al. 2011). The second group of apertural muscles can be differentiated into two distinct sets: a) Duplicature bands (= parietovaginal bands of gymnolaemates, cf. Schwaha et al. 2011) are peritoneally lined, thin bands that in phylactolaemates extend from the tentacle sheath (rarely from the diaphragm itself) toward

▸ Fig. 3.22: Apertural muscles of bryozoans. (A–E) Semithin sections. (F) Confocal laser scanning microscopy. A, Section of the phylactolaemate Plumatella sp. showing a duplicature band connecting the tentacle sheath with the body wall. B, Phylactolaemate Hyalinella punctata showing duplicature band including muscle fibers and separate vestibular dilatators. C, Ctenostome Alcyonidium gelatinosum showing elongated duplicature band. D, Longitudinal section of the cyclostome Cinctipora elegans showing attachment organ and ligament as well as the prominent diaphragmatic sphincter. E, Ctenostome Arachnidium fibrosum showing prominent vestibular muscles. F, Apertural muscles of the cheilostome Fenestrulina malusii. Note also the rim of the operculum. Abbreviations: ao – attachment organ, atl – attachment ligament, bw – body wall, ec – ectocyst, ep – epidermis, co – collar, db – duplicature band, dia – diaphragm, ds – diaphragmatic sphincter, ooc – operculum occlusor, op – operculum, pdm – parietodiaphragmatic muscle, pec – peritoneal cilia, rtl – retracted lophophore, ts – tentacle sheath, vd – vestibular dilatator, vm – vestibular muscles, vw – vestibular wall.



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the lateral body wall (Figs. 3.21 and 3.22 A–C, F). Distinct smooth longitudinal muscle fibers traverse each band. In retracted state, these bands are stretched and extend distally. Like the entire apertural area, these bands are radially arranged in phylactolaemates. Four of these bands are generally present in gymnolaemates. For unknown reasons, several ctenostomes have reduced these bands (cf. Schwaha et al. 2011, Schwaha & Wanninger 2018). As far as known, all cheilostomes possess the general set of four bands (Schwaha et al. 2011); however, some genera show six (Schwaha pers. observation) or eight (Schwaha et al. 2011). The term parietovaginal bands is the most frequently used term for these peritoneal bands in gymnolaemates. Since these are clearly homologous to the duplicature bands of phylactolaemates (Schwaha et al. 2011), it is recommended to use the term duplicature bands for all bryozoans. b) Distally of the duplicature bands, a series of muscle bundles originates from the lateral body wall and attaches to the vestibular wall. In phylactolaemates, these are thin, but numerous, muscle fibers, the vestibulum dilatators, that are found radially over the entire length of the vestibular wall (Figs. 3.21 A, B and 3.22 B). In gymnolaemates, these are few and thick bundles (Fig. 3.22 E, F) that in accordance with the duplicature bands are present as four bundles in most taxa. Only few ctenostomes sometimes have only two functional sets, which is mostly in association with lip-like closing structures of the aperture or orifice (Schwaha & Wanninger 2018). Likewise, cheilostomes possess only two bundles on each lateral side of the apertural area (Mukai et al. 1997, Schwaha et al. 2011). A proximal, smaller portion of these muscles attaches to the diaphragm (parietodiaphragmatic muscle). In ctenostomes, the more distally located parieto-vestibular muscles show a lot of variation in their extent and insertion areas at the vestibular wall. The parietovestibular muscles are homologous to the operculum occlusors – the main closing muscles of the operculum of cheilostomes (Schwaha et al. 2011; Fig. 3.21). The cyclostome condition is only little analyzed and understood. Some attempts have been made to homologize the apertural muscles of phylactolaemates and gymnolaemates to cyclostomes (Schwaha et al. 2011). The attachment ligaments of the attachment organ are topologically similar to the duplicature bands of other bryozoans, but there is a high variation in the presence, absence, and symmetry of the entire structure (see Boardman 1998). Likewise, separate muscle fibers, vestibular

muscles, in the distal zooidal area are only present in few analyzed genera, such as Crisia (Nielsen & Pedersen 1979, Worsaae et al. 2020; termed longitudinal ectodermal muscles by the authors, Fig. 3.20 D) or Tubulipora. Based on their position, these are reminiscent of the vestibular muscles of phylactolaemates and gymnolaemates. However, a profound survey and ground pattern reconstruction of the entire apertural area is necessary to draw any conclusions on the evolution of these structures among cyclostomes.

3.12.2.1 Functional aspects of the apertural muscles Apertural muscles apparently evolved in connection with the polypide retraction process in order to close, but also open, the apertural area properly. Clearly, muscles of the vestibular wall and the diaphragmatic sphincter facilitate closure of the apertural area when zooids are retracted. Hence, opercular structures closing the orifice render distinct vestibular wall muscles unnecessary and explain their absence among cheilostomes. The vestibular muscles have two functions: on the one side, they widen the apertural area for protruding polypides, and on the other hand, they minimize the space between vestibular wall and body wall and prevent fluid of the body cavity from being pressed into this area during the protrusion process. Likewise, the modification of these muscles into the operculum occlusor of cheilostomes enables operculum closure. The function of the duplicature bands is not entirely understood. It appears that these bands fasten the position of the retracted polypide in zooids and perhaps enable a more coordinated protrusion of the polypide. Particularly their absence among several ctenostomes remains enigmatic.

3.12.3 Lophophoral muscles The musculature of the lophophore can be differentiated into muscles in each individual tentacle and those located at the lophophoral base (Gawin et al. 2017, Schwaha & Wanninger 2018). The latter are in strong association with the mouth opening and the musculature of that area.

3.12.3.1 Tentacle muscles Tentacle muscles are very similar among bryozoans. In all studied species, they consist of a frontal and an abfrontal



longitudinal muscle band. These muscles are located at the basal side of the peritoneal lining of each tentacle. In phylactolaemates, the tentacle muscles are continuous with their corresponding lophophoral base muscles, whereas myolaemates show a gap of variable size separating the tentacle muscles from the lophophoral base musculature (Schwaha et al. 2011, 2018, Schwaha & Wanninger 2018). Tentacle muscles can be smooth or as in most analyzed bryozoans composed of striated sarcomeres. Few examples are known where both can occur in a single species. Distinct spot-like or sphincter muscles can occur at the distal tips of tentacles (Schwaha & Wanninger 2018).

3.12.3.2 Lophophoral base muscles Phylactolaemata The lophophoral base muscles in phylactolaemates are distributed over the entire lophophore at the proximal insertion of each tentacle and can be divided into abfrontal and frontal lophophoral base muscles (Fig. 3.23). The former is a series of oblique muscle bands that extend toward the tentacles with a series of muscles shaped in a stack of inverted V’s. The frontal tentacle muscles extend as single or double basal rootlets at the lophophoral base (or the lophophoral arms). In larger, gelatinous species, the frontal roots on the lophophoral arms are connected to thin muscles extending along the ganglionic horns. The frontal roots of the oral tentacles are connected with the circular musculature of the pharynx (Gawin et al. 2017; Fig. 3.24). Due to the horseshoe-shaped lophophore with the lophophoral arms, additional muscle bundles are present at the lophophoral base. These comprise lophophoral arms muscles that are situated on the proximal side of the lophophoral arms and effectuate movement of the entire lophophoral arms (Schwaha & Wanninger 2012). In larger species such as the gelatinous Cristatella mucedo or Pectinatella magnifica, these are more prominent than the smaller plumatellids (Gawin et al. 2017). Two additional muscle systems are distinguishable at the lophophoral base of phylactolaemates: ring canal musculature and muscles of the epistome. As mentioned in section 3.5.1, the ring canal of phylactolaemates is restricted to the short oral row of tentacles. On its proximal side, several muscles extend radially from the foregut into the lophophoral base and adjacent tentacle sheath. The epistome of phylactolaemates has two possible muscle patterns, one with smooth fibers located in its epithelial

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lining or separate muscle fibers traversing its internal cavity (Gawin et al. 2017, see also chapter 7).

Myolaemates In myolaemates, the tentacle base musculature is present as three to four sets that are distinctive and not in direct contact with the tentacle muscles. These can be categorized into a) longitudinal muscles of the lophophoral base, b) circular lophophoral base muscles of gymnolaemates, c) buccal dilatators, and d) proximal lophophoral base muscle of cyclostomes. a) The longitudinal fibers are present as abfrontal or frontal longitudinal muscles. Frontal muscles were only detected in cyclostomes (Schwaha et al. 2018; Figs. 3.23 C and 3.24 D), whereas the abfrontal muscles are present in cyclostomes and gymnolaemates. In cyclostomes, these consist of a simple short branch of longitudinal muscles at the lophophoral base (Schwaha et al. 2018; Fig. 3.23 C). In gymnolaemates, the abfrontal lophophoral base muscle is similar in cteno- and cheilostomes, with some variation of their size and extent. In addition, so-called v-shaped muscles are located slightly distally and are partially overlapping to the longitudinal lophophoral base muscles. These can show considerable variation in their presence and appearance in ctenostomes (Schwaha & Wanninger 2018; Figs. 3.23 D–G and 3.24 E). b) The circular lophophoral base muscles of gymnolaemates are either present as a complete muscular ring on the frontal side of the lophophoral base, slightly above the mouth opening, or as short intertentacular, separate muscle fibers of similar orientation, but broader in proximo-distal axis. The latter were originally termed basal transversal muscle in the cheilostome Cryptosula pallasiana (Gordon 1974). They are present in all cheilostomes, whereas ctenostomes show different variations ranging from a complete ring of circular fibers, an almost complete ring or even just a few fibers as in cheilostomes (Schwaha & Wanninger 2018; see also Fig. 3.24 E). c) Buccal dilatators are present in all myolaemates and are (mostly striated) radial muscle fibers that extend from the medial side of the lophophoral base, pass the lophophoral ring canal and insert on the outer wall of the inner peritoneal lining below each tentacle (Gordon 1974, Schwaha et al. 2011, 2018, Schwaha & Wanninger 2018; Fig. 3.23 C, D, G). The buccal dilatators aid the prominent myoepithelial pharynx of

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myolaemates in extending the pharyngeal volume and thus are a distinct apomorphy together with the myoepithelial pharynx (Schwaha et al. 2020). d) Proximal lophophoral base muscles are only present in cyclostomes and are located proximally of the longitudinal abfrontal lophophoral base muscles. They consist of three triangularly arranged short fiber bundles of different fiber orientation (Schwaha et al. 2018; Figs. 3.23 C and 3.24 D).

3.12.4 Tentacle sheath muscles In phylactolaemates, the musculature of the tentacle sheath consists either of an orthogonal grid of circular and longitudinal muscles or solely longitudinal muscles (Schwaha & Wanninger 2012, Gawin et al. 2017; Fig. 3.24 A). The orthogonal arrangement is considered as ancestral condition since it reflects the situation of the body wall musculature. In myolaemate bryozoans, only longitudinal muscles are present (Fig. 3.23 F). The only exceptions are victorellid and walkerioidean ctenostomes, which possess a basket of diagonal muscles in the tentacle sheath (Schwaha & Wanninger 2018, Pröts et al. 2019). The muscle fibers of the duplicature bands originating from the tentacle sheath (see apertural muscles above) are continuations of the tentacle sheath fibers.

3.12.5 Digestive tract muscles These muscles differ in accordance to the general digestion modes of kneading phylactolaemates and rotating myolaemates (see section 3.9.1). In phylactolaemates, the entire digestive tract is covered with a dense array of circular muscles that allow peristaltic movements of food particles within the gut (Figs. 3.20 E and 3.24 A). All circular muscles are striated except in the intestine, where they are smooth fibers (Schwaha & Wanninger 2012, Gawin et al. 2017). Few thin longitudinal fibers were detected in the

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foregut of the phylactolaemate Asajirella gelatinosa (Mukai et al. 1997). Myolaemates have as apomorphic character the myoepithelial pharynx. Circular striated muscles surrounding the foregut are present as found in phylactolaemates, but the pharynx epithelium is thickened in three areas, which diminishes the pharynx lumen to a triradiate shape. The pharyngeal epithelium is a myoepithelium with distinct striated myofibrils inserted at the lateral cell membranes of the adjacent cells of the epithelium. The pharyngeal lumen is very narrow, and contraction of the muscles in the epithelium results in a rapid expansion of this lumen leading to suction (Schwaha et al. 2020). This process is enhanced by the buccal dilatators, whereas the outer pharyngeal ring muscles compress the pharyngeal volume. The adjoining esophagus is short in cyclostomes and often long in gymnolaemates and has a similar, but less dense, muscular lining as the pharynx and more interspersed longitudinal fibers (Schwaha et al. 2011). The cardia and remaining gut musculature of myolaemates are mostly composed of smooth, sparse fibers. The midgut (cardia, caecum, pylorus) consists mainly of few circular fibers with few longitudinal ones, whereas the intestine always possesses only longitudinal fibers (Figs. 3.15 I and 3.24 B). In ctenostomes with a distinct proventriculus or gizzard, the musculature is very prominent and consists of smooth, dense circular muscles (Figs. 3.11 and 3.20 C). Several ctenostomes show a distinct concentration of circular muscles as cardiac constrictor, which often is diagnostic for species discrimination (Braem 1951, Schwaha & Wanninger 2018).

3.12.6 Retractor muscles The retractors are the most prominent and evident pair of muscles in every zooid. Ontogenetically, they develop from two lateral anlagen (Jebram 1986), which is also

◂ Fig. 3.23: Muscular system of the lophophoral base of bryozoans. All f-actin stainings and confocal laser scanning microscopy. (A) View from the anal side of the phylactolaemate Stephanella hina. (B) Lateral view of a retracted zooid of the phylactolaemate Fredericella sultana. Note the asterisk marks distinctly striated retractor muscle fibers whereas the remaining ones mostly appear smooth. (C) Cyclostome Tubulipora sp. (D) Ctenostome Mimosella verticillata. (E) Cheilostome Calpensea nobilis. (F) Cheilostome Myriapora truncata. (G) Ctenostome Paludicella articulata. Abbreviations: afb – abfrontal base muscle of phylactolaemates, afm – abfrontal base muscle of myolaemates, anm – annular muscles of the membranous sac, bd – buccal dilatator, bwm – body wall musculature, ebm – epistomial base muscles, fbm – frontal base muscles of phylactolaemates, fm – frontal base muscles of cyclostomes, int – intestine, lbm – circular lophophoral base muscles, mo – mouth opening, pom – proximal base muscle of cyclostomes, pxm – pharynx musculature, rm – retractor muscles, tm – tentacle muscles, tsm – tentacle sheath muscles, vem – vestibular muscles of cyclostomes, vm – v-shaped muscles of gymnolaemates.

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Fig. 3.24: Schematic overview of muscular systems in bryozoans. (A) Lateral view of a phylactolaemate zooid. Modified from Gawin et al. (2017). (B) Overview of the gut musculature of a gymnolaemate zooid. Modified from Schwaha and Wanninger (2018). (C) Top view of the lophophoral base musculature of a phylactolaemate bryozoan showing the arrangement of the frontal base muscles. Modified from Gawin et al. (2017). (D) Lateral view of the lophophoral base of the cyclostome Cinctipora elegans showing the four different sets of muscles. Modified from Schwaha et al. (2018). (E) Lateral view of the lophophoral base muscles of gymnolaemate bryozoans. Modified from Schwaha and Wanninger (2018). Abbreviations: a – anus, afm – abfrontal lophophoral base muscle, at – atrium, bd – buccal dilatator, bw – body wall, ca – cardia, cae – caecum, cv – cardiac valve, db – duplicature band, dg – digestive tract, ds – diaphragmatic sphincter, ep – epistome, es – esophagus, f – funiculus, fm – frontal lophophoral base muscle, int – intestine, la – lophophoral arm, lam – lophophoral arm muscle, lb – lophophoral base, lbm – circular lophophoral base muscle/basal transversal muscle, lco – lophophoral concavity, mo – mouth opening, o – orifice, pdi – parieto-diaphragmatic muscle, ph – pharynx, pom – proximal lophophoral base muscle, pve – parieto-vestibular muscle, pyl – pylorus, rm – retractor muscle, rnc – ring canal, ts – tentacle sheath, v – vestibulum, vd – vestibulum dilatator, vm – v-shaped lophophoral base muscle.



reflected in most adults possessing two distinct portions. The muscle bundles are located laterally of each zooid (Figs.  3.1–3.4, 3.20, and 3.23). In phylactolaemates, the retractor muscles of adults emerge from the basal or lateral body wall and insert on the lophophoral base, parts of the tentacle sheath, and various parts of the oral shank of the digestive tract (foregut, cardia, and parts of the caecum) (Gawin et al. 2017). The myolaemate retractor muscles similarly originate from the proximal or lateral body wall and insert at the lophophoral base. Several cyclostomes have an additional bundle that inserts at the cardia (e.g. Nielsen & Pedersen 1979). In gymnolaemates, additional bundles were only recently described in the ctenostome Aethozooides uraniae. In the latter, the retractors attach over the entire range of the oral side from the tentacle sheath until the caecum (Schwaha et al. 2019). In the cheilostome Calpensea nobilis the retractors attach, in addition to the lophophoral base, to distal areas of the tentacle sheath (Schwaha, pers. observation). In general, it seems that the retractor muscles of gymnolaemates are similar, but few exceptions are present. The functional importance of these variations remains unknown, particularly when thick muscle bundles attach to thin tissues like the tentacle sheath. Retractors are generally composed of smooth fibers (Fig. 3.20 G). In phylactolaemates, parts of the fibers show a striated appearance at their attachment site at the lophophoral base (Schwaha & Wanninger 2012; Fig. 3.23 B). Striated retractor muscle fibers are evident in several myolaemates (Worsaae et al. 2020, Schwaha & Wanninger 2018; Fig. 3.20 F). It appears that the amount of retractor fibers correlates with the type of muscles. Species with many, thin fibers tend to have striated fibers and those with fewer but thick fibers smooth ones (Schwaha & Wanninger 2018). However, transmission electron microscopic analyses are still required to assess the different striation or non-striation patterns.

3.13 Excretory system Little is known about excretory systems in bryozoans. Generally, bryozoans are considered to lack specific excretory organs, which in fact means that distinct nephridia are not present. Naturally, excretion of metabolic wastes also has to occur in bryozoans. Excretion is probably effectuated over parts of the lophophore as previous vital dye experiments have shown (Marcus 1926b). In addition, at least

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phylactolaemates form distinct vacuoles on the abfrontal side of the tentacles (e.g. Braem 1890), probably removing substances from the zooid. The abfrontal side of the tentacles carries only few sensory elements, and disposal of undesirable substances in this area does not interfere with ciliary currents. Special coelomocytes termed phagocytes are regarded as excretory mechanisms in bryozoans. Coelomocytes in general are large single cells in the body cavities. These originate from cells detaching from the peritoneal layer into the coelomic cavity – or in case of the patchy peritoneal layer of gymnolaemates into the vestigial coelom. In cyclostomes, single cells can be found in the exosaccal cavity. The phylactolaemate forked canal was previously considered to be a vestigial nephridium. The massive ciliation of this canal accumulates coelomocytes and sperm toward its lumen. Cristatella mucedo even forms special excretory bladders at the median junction of this canal (Schwaha et al. 2020).

3.14 Coelomocytes Free-floating cells in the body cavity have been predominantly described in gymnolaemate and phylactolaemate bryozoans (Mukai et al. 1997; Fig. 3.25). The latest and most actual analysis of these important cells was in 1964 by Mano on the phylactolaemate Lophopodella carteri. Nine distinct forms of coelomocytes variously termed lymphocytes, leucocytes, amoebocytes, phagocytes, granulocytes, or vacuolated cells were categorized in the latter species. Leukocytes of different morphology were also recognized in the other clades. Very little is known on the detailed structure or function of these cells. They appear to play a vital part in phagocytosis (i.e. phagocytes) and were also experimentally shown to accumulate injected vital dye (Harmer 1891). Coelomocytes play a crucial role in the immune system in invertebrates (e.g. Matranga et al. 2005), and this may also be part of their function in bryozoans. Owing to their generally more transparent cystids, they were most frequently observed in phylactolaemates. The abundance of coelomocytes appears to correlate with external stress to the colony or zooid (Schwaha, pers. observation). Coelomocytes also play an important role as guiding structures during zooidal budding (observed in phylactolaemates; Handschuh & Schwaha unpublished observations).

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Fig. 3.25: Bryozoan coelomocytes. (A–C) Phylactolaemata. (D, E) Ctenostome Gymnolaemata. A, Coelomocyte close to the body wall of Hyalinella punctata showing numerous inclusions in the cells. B, Coelomocytes of Hyalinella punctata that appear to be multinuclear. C, Vesicular coelomocyte of Cristatella mucedo partially showing thin appendages. D, Vesicular coelomocytes of Arachnidium fibrosum. E, Coelomocyte of Alcyonidium gelatinosum in association with the lateral body wall. Abbreviations: bc – body cavity, cc – coelomocyte, ec – ectocyst, ep – epidermis, gep – glandular epidermis, izs – interzooidal septum, pe – peritoneum, pp – pore plate.

Literature Allman, G.J. (1856): A Monograph of the Fresh-Water Polyzoa. Ray Society Publications 28. Ray Society, London: 119 pp. Ambros, M., Wanninger, A. & Schwaha, T. (2018): Neuroanatomy of the plumatellid bryozoan Hyalinella punctata reveals a common pattern in a small group of freshwater bryozoans. J Morphol 279: 242–258. Annandale, N. (1916): Zoological results of a tour in the far east. Polyzoa, Entoprocta, and Ctenostomata. Mem Asiat Soc Bengal 6: 13–37. Atkins, D. (1932): The ciliary feeding mechanism of the entoproct Polyzoa, and a comparison with that of the ectoproct Polyzoa. Q J Microsc Sci 75: 393–423.

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Andrew N. Ostrovsky

4 Sexual reproduction in Bryozoa 4.1 Generalities of bryozoan reproduction Bryozoan colonies are hermaphrodites consisting of sterile and sexual zooids. Gonads have mesothelial origin. Sperm is released to the water column, but fertilization is internal, followed by either release of zygotes to the water column in spawners or their retention in incubating species. Accordingly, embryogenesis results in long-living planktotrophic larvae in the former and short-living lecithotrophic larvae in the latter. Larval settlement and metamorphosis result in the first zooid – the ancestrula, which begins asexual budding to form a colony. Incubation can be preparitive (viviparity) or postparitive (brooding) and often accompanied by matrotrophy (extraembryonic nutrition) (reviewed in Ström 1977, Zimmer & Woollacott 1977, Reed 1991, Woollacott 1999, Ostrovsky 2008a, 2013a, b, Ostrovsky et al. 2008, Schwaha et al. 2020).

4.2 Reproductive patterns Bryozoa is an extraordinary group in respect of the diversity of reproductive patterns which evolved. Of six described patterns, five have been found in Gymnolaemata (Ostrovsky 2013a). Pattern I (known in the most ancient cheilostome clade, Membraniporina and in several ctenostomes) includes the production of many to several (less than 10) small oligolecithal oocytes in the fertile zooid, which are fertilized during or shortly after ovulation and spawned in cohorts. Except two ctenostomes, bryozoans with this pattern release zygotes via the intertentacular organ: a two-chambered ciliary tube placed between the two anal tentacles above the cerebral ganglion (Ostrovsky & Porter 2011). Incubation is absent, and embryos develop into planktotrophic cyphonautes larvae. In contrast, brooding and lecithotrophy are characteristic for pattern II (found in most Gymnolaemata). Production of many/several to few meso- or macrolecithal oocytes of small, medium, or large size is accompanied by intraovarian (precocious in cheilostomes) or post-ovulatory fertilization and near-simultaneous or

https://doi.org/10.1515/9783110586312-004

successive oocyte maturation. In most Cheilostomata with this pattern, the oocyte develops with a nurse cell. Sequential or simultaneous brooding of one to several embryos occurs at the surface of the maternal zooid, in the tentacle sheath of the polypide, or in a specialized brood chamber. Only few known brooders release zygotes via the intertentacular or a similar tube-like organ, while in the vast majority, oviposition occurs via a supraneural coelomopore (Ostrovsky & Porter 2011). Noteworthy, the cheilostome Tendra zostericola shows a number of transitional traits between zygote-spawners with planktotrophic larvae and brooders with lecithotrophic larvae, i.e. combining the features of patterns I and II. While having a membraniporine zooidal morphology, this species is a brooder producing lecithotrophic larvae with a rudimentary gut. Several small mesolecithal oocytes are developed in the ovary without nurse cells and are fertilized during or shortly after ovulation. They are further transferred as a group from the fertile zooid to the brood chamber via the intertentacular organ (Repiachoff 1875, 1878, Paltschikova-Ostroumowa 1926, Braiko 1967). Pattern III (found in some species of both gymnolaemate orders and in Phylactolaemata) is characterized by matrotrophic brooding. It includes the production of numerous small oligolecithal oocytes (but only one develops to a larva) in phylactolaemates and several or one oligo- or mesolecithal oocytes in gymnolaemates (whose development is accompanied by a nurse cell in Cheilostomata). Fertilization is intraovarian (precocious in cheilostomes); embryos are brooded in groups (in some ctenostomes) or one at a time in the tentacle sheath of the polypide or in brood chambers (in some ctenostomes and cheilostomes and in all phylactolaemates). Brooding is accompanied by extraembryonic nutrition resulting in non-feeding larvae. Importantly, the reproductive pattern of phylactolaemates combines ancestral and advanced characters – numerous small oocytes, matrotrophy, and putative nurse cells (in Lophopus) (Marcus 1934, Brien & Mordant 1956, Wood 1983). Pattern IV (found in certain cheilostomes and suggested for one ctenostome) combines the features of pattern II (macrolecithal oocytes) and of pattern III (extraembryonic nutrition). It is characterized by sequential (in cheilostomes) production of several

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macrolecithal oocytes (of small, medium or large size), which are transferred into a specialized brood chamber where they develop into lecithotrophic larvae. Brooding is matrotrophic. In cheilostomes with this pattern, the oocytic development is accompanied by a nurse cell, and fertilization is intraovarian and precocious (Ostrovsky 1998; Moosbrugger et al. 2012; Nekliudova et al. 2019a). The ctenostome Flustrellidra hispida supposedly has reproductive pattern IV, too. The ovary produces several relatively small macrolecithal oocytes that are further transferred to the modified tentacle sheath. Simultaneous brooding is accompanied by embryonic growth suggesting matrotrophy (Pace 1906, Kvach et al. 2019). Pattern V (known only in the cheilostome family Epistomiidae) involves the production of a single, small oligolecithal oocyte developing into an embryo in the coelomic cavity of the enlarged fertile zooid (presumably, inside the ovary). Embryonic development in this viviparous group is accompanied by extraembryonic nutrition (Dyrynda & King 1982). Similarly, viviparity and extraembryonic nutrition are characteristic of the sexual reproduction of cyclostome bryozoans. One to two small oligolecithal oocytes are developed in the ovary where cleavage starts. Further development is characterized by polyembryony supported by matrotrophy. Development of numerous larvae occurs in the enlarged polymorphic gonozooids (Harmer 1893, 1896, 1898, Borg 1926). This variant is considered as reproductive pattern VI. Gonozooids are absent in the family Cinctiporidae, which suggests that larval development might occur inside very large autozooids (Schwaha et al. 2018).

4.3 Colony sexual structure and sexual polymorphism Bryozoans are colonial hermaphrodites, with spermatogenic tissue and ovaries developing either within the same zooid (zooidal hermaphroditism) or in different zooids (zooidal gonochorism). Experiments with the cyclostome Filicrisia geniculata suggest the existence of colonies of separate sexes (Jenkins et al. 2015), but this is probably an exception. Mature bryozoan colonies consist of sterile and sexual zooids that can be gonochoristic (male and/ or female) or hermaphroditic or both. Depending on the time of gamete maturation in colonies with zooidal hermaphroditism, autozooids (and thus colonies too)

may be protandrous, protogynous, or simultaneous hermaphrodites. A similar pattern is observed in colonies with zooidal gonochorism, with male and female zooids sometimes exhibiting sexual dimorphism. Morphological differences between male and female zooids are correlated with gamete release and embryonic incubation and may involve the polypide, the cystid, or both (Reed 1991, Ostrovsky 2013a). In general, a bryozoan colony is a dynamic system in which the gonads form, mature, and function at different times in different zooidal generations (Ostrovsky 1998). For instance, protandrous hermaphrodite colonies of the cheilostome Cribrilina annulata consist of sterile, male, and hermaphrodite zooids. Correspondingly, they are initially sterile, then male and hermaphrodite. After degeneration of the spermatogenic tissue in male and hermaphrodite zooids, the colony becomes female. In overwintering colonies of C. annulata, ovaries degenerate and form again at the beginning of the next reproductive season in younger peripheral zooids. Thus, colonies in this species change from female to winter-sterile and further to protandrous-hermaphrodite again in spring (Nekliudova et al. 2019b). Zooids may change sex or acquire it. For instance, in the overwintered colonies of the cheilostome Chartella papyracea, the former female zooids that lost their ovary last autumn developed spermatogenic tissue in spring (Dyrynda & Ryland 1982). In the cheilostomes Celleporella hyalina, Antarctothoa bougainvillei, and A. tongima, some autozooids may become males after 1–2 months of normal functioning (Rogick 1956, Powell 1967, Cancino & Hughes 1988). In the latter species, some female zooids may become male.

4.4 Gonado- and gametogenesis Primordial germ cells differentiate from somatic cells in the cystid wall. Spermatogonia develop from them either within the cystid mesothelium of the main body cavity or in association with the funiculus or within both, while oogonia either appear in the mesothelial layer of the polypide bud, or, in some taxa, within the cystid mesothelium (Reed 1991, Ostrovsky 2013a). Germ cells cannot be detected by light microscopy (Hageman 1983), while their divisions result in the formation of either spermatogonia or oogonia. The male gonad (spermatogenic tissue) lacks any epithelial walls, ducts and accessory glands in bryozoans. It can be compact (then sometimes termed testis), continuous, or consist of diffuse cell clusters (Figs.  4.1, 4.2, and



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Fig. 4.1: Spermatogenesis in phylactolaemate and cyclostome Bryozoa. (A, B) Plumatella fungosa (stained whole mounts). Early compact (A) and fully formed (B) testes with ripe sperm. (C, D) Hyalinella punctata (histological sections). Lobate testis showing early (C) and more advanced (D) stages of spermiogenesis. (E, F) Crisia sp. (histological sections). Spermatogenic tissue filling the proximal part of cystid; sperm is seen among spermatocytes in F to the left from the caecum and in the insert. Abbreviations: f – funiculus, p – polypide.

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4.3  A). Mitotic divisions of the spermatogonia result in morulae consisting of spermatocytes united around the central cytophore (Fig.  4.1 D). Each spermatocyte undergoes meiotic division with four spermatids to emerge. Further on, spermatogenesis results in the formation of spermatozoids clustered in spermatozeugmata (at least in some Gymnolaemata). Mature spermatogenic tissue is a cell complex consisting of fully formed male gametes and their progenitors at different developmental stages (Figs. 4.1 B, D, F and 4.2 C‒F) (reviewed by Franzén 1977, Reed 1991). The fate of oogonia is different, and whereas some of them grow and differentiate to form the oocytes, others divide to support the oogonial population that exists in the ovary for some time. In most cheilostomes, oogonia divide to form oocytic doublets (Ostrovsky 1998, 2013a). Ovarian structure has been mainly studied using light microscopy in bryozoans, and ultrastructural studies are rare. The growing female cells (oogonia and early oocytes) are recognizable when they become larger than the somatic cells, and while they grow in size, they become surrounded by a single-cell layer of mesothelial cells that become the prospective ovarian wall (Ostrovsky 2013a). Further development of the female gonad includes multiplication and growth of both somatic (ovarian) and sexual cells (Figs.  4.3‒4.6). In cheilostomes, the ovary is always approached by funicular strands. Depending on the reproductive pattern, the ovarian structure and oogenesis differ. Pattern I. In malacostegan cheilostomes, the ovary develops on the lateral wall of an autozooid with functional polypide, at the site where several funicular strands fuse (Hageman 1983, Ostrovsky 2013a) (Figs. 4.2 A, B and 4.3 A‒D). The parietal peritoneum transforms to a follicular epithelium around oogonia and oocytes (Fig. 4.3 E, F). According to Hageman (1983), the follicular cells regulate vitellogenesis, synchronize differentiation of oocytes, and transport low-molecular metabolites toward them. Follicle cells are also able to phagocytose abortive oocytes. The mature ovary consists of three zones: peripheral germinal, central growth zone, and centro-apical ovulatory. The germinal zone includes oogonia and early previtellogenic oocytes, whereas the central growth zone contains developing oocytes at various stages of vitellogenesis. Oocytes are entirely surrounded by the follicle cells forming from one to several layers. Both the follicle cells and oocytes possess a developed synthetic machinery (including rough endoplasmatic reticulum). The intercellular spaces between follicle cells are filled with a proteinaceous substance (secreted by these cells)

that is endocytosed by growing oocytes. Thus, ultrastructural observations indicate both auto- and heterosynthetic sources of yolk (Hageman 1983, Shevchenko et al. in preparation). The ovarian structure of the cheilostome brooder Tendra zostericola is similar to the one described above for the broadcasting species (Shevchenko et al. in preparation). Before ovulation, mature oligolecithal oocytes move into the centro-apical ovulatory zone. Ovulation follows the breakdown of the nuclear envelope. Flattened oocytes accumulate in the body cavity of the maternal zooid in groups of 20 to 30 before spawning. The number of mature oocytes produced by one zooid ranges from 4–5 to 40–50 in different zygote-spawning cheilostomes (e.g. Fig.  4.3 D‒F) and from 6–10 to 60 in ctenostomes with I pattern (Fig. 4.4 A, B) (Ostrovsky 2013a). Pattern II. In most brooding cheilostomes, the ovary develops in association with the polypide bud later relocating to the basal zooidal wall (Ostrovsky 2013a). Growing oogonia are normally solitary, round or oval. The division of the oogonium results in an oocyte doublet of two sibling cells, interconnected by a cytoplasmic bridge (Fig. 4.5 A). These cells will later differentiate into a vitellogenic oocyte and its nurse cell (Fig. 4.5 B‒D) (Dyrynda & King 1983, Ostrovsky 1998, 2013a). The destiny of each sibling is presumably determined by fertilization: the fertilized cell becomes the vitellogenic oocyte. Syngamy obviously occurs immediately after the division of the oogonium into the oocyte doublet (Ostrovsky 2008a, 2013a). In contrast, sperm fuses with a mature oocyte in the ctenostome Bowerbankia/ Amathia gracilis and is developed without a nurse cell (Reed 1988). The number of oocytes in the ovary strongly varies in both gymnolaemate orders, ranging from 1 to 25 oocytic doublets in Cheilostomata and from 1 to 80‒100 solitary oocytes in Ctenostomata (Ostrovsky 2013a) (Fig.  4.4 C). In the former, vitellogenesis usually occurs in only one doublet in the ovary at the same time, whereas one to many developing oocytes can simultaneously maturate in ctenostomes. At the same time, the ovary normally contains one to few growing and late (premitotic) oogonia. In cheilostomes with pattern II the leading doublet in the ovary occupies the central (often apical) position surrounded by follicle cells (Fig. 4.5C). As synthesis and transport in the ovary are enhanced, the follicle cells are progressively enlarged. In the ovary with a vitellogenic oocyte, the follicle cells covering the animal pole of the oocyte flatten and become “squamous,” while those limiting the oocyte laterally and basally from the sides



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Fig. 4.2: Spermatogenesis in gymnolaemate Bryozoa. (A‒D) Electra pilosa (stained whole mounts). Hermaphrodite zooids with spermatogenic tissue formed along the zooidal walls and early ovary (A, B). Close views showing proximal part of zooidal cavity filled with spermatogenic tissue (C, D) and ovary (C). (E) Calyptotheca hastingsae (histological section). Spermatogenic tissue showing various stages of spermatogenesis. (F) Triticella flava (histological section). Hermaphrodite zooid with ovary surrounded by spermatogenic tissue. Abbreviations: ov – ovary, p – polypide, rm – retractor muscles, st – spermatogenic tissue, tw – transversal wall.

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Fig. 4.3: Oogenesis in Cheilostomata (pattern I). (A‒F) Electra pilosa (stained whole mounts). A, Part of the colony showing hermaphrodite zooids with functional polypides, early ovaries and spermatogenic tissue near zooidal walls. B‒D, Early ovaries showing various stages of growth. E, F, Close-up of hermaphrodite zooids showing spermatogenic tissue and part of the ovary consisting of the peripheral follicle cells and oocytes of various ages. Abbreviations: fo – follicle cells, ov – ovary, p – polypide, pv – previtellogenic oocyte, rm – retractor muscles, st – spermatogenic tissue, vo – vitellogenic oocyte.



Fig. 4.4: Oogenesis in Ctenostomata (patterns I and II). (A, B) Hislopia malayensis (histological sections). Early ovary on zooidal wall (A). Mature ovary with microlecithal oocytes on various stages of development (attachment zone to zooidal wall is out of section plane) (B). (C) Hypophorella expansa (histological section). Zooid with mature ovary containing two macrolecithal oocytes.

become elongated, cubic or prismatic (Fig. 4.5 C, D). Synchronous growth of the cells of the oocyte doublet continues throughout the previtellogenic period (Fig. 4.5 A, B). In early stages of the vitellogenic phase, the oocyte is normally larger than the nurse cell (Fig. 4.5 D). Both cells contain yolk granules in their cytoplasm. Later during vitellogenesis, the oocyte growth rate greatly exceeds that of the nurse cell, which finally stops growing. The nucleus of the mature nurse cell occupies most of the cell volume, indicating intense activity. During this period, the development of all other (younger) oocyte doublets in the ovary is suppressed (Fig.  4.5 D). In the course of ovulation, the nurse cell is separated from the oocyte and degenerates in the zooidal cavity or in the ovary. Ovulation of the leading doublet triggers vitellogenesis in the succeeding oocyte doublet (Ostrovsky 1998, 2013a). Ultrastructural studies showed that the follicular epithelium consists of inner squamous and outer columnar cells during vitellogenesis in the cheilostome Chartella papyracea (Dyrynda & King 1983). The columnar follicle

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cells show signs of synthesis and transport activity being filled with rough endoplasmic reticulum, whereas both the vitellogenic oocyte and its nurse cell form microvilli and pinocytotic vacuoles in this species. The nurse cell also produces and transports ribosomes to its sibling via the cytoplasmic bridge. Size increase and development of the synthetic apparatus in the follicle cells are characteristic of vitellogenesis in the ctenostome Bowerbankia/Amathia gracilis. Else, these cells develop extensive basolateral interdigitations and gap junctions. Ultrastructural observations indicate both auto- and heterosynthetic sources of yolk in this species (Reed 1988, 1991). As in cheilostomes, oocytes are macrolecithal in ctenostomes with this pattern (Fig. 4.4 C). Pattern III. In contrast to non-matrotrophic species, placental bryozoans have ovaries of fewer cells that produce small oligolecithal oocytes. For example, the follicle of Bugulina flabellata is represented by a continuous layer of flattened cells with a small cone of columnar cells (Dyrynda & King 1983). The same structure was described in Bugula neritina (Ostrovsky 2013a, Mathew et al. 2018). In B. flabellata, the ovary contains one to three oocytic doublets, although most of the studied species have no more than two. Similar to non-placental species, the onset of vitellogenesis is accompanied by the formation of microvilli on the oolemma and on the plasma membrane of the nurse cell. The ovary is developed in association with a polypide bud. Only histological data are available in respect to the oogenesis in Phylactolaemata. The ovary forms on the body wall (Fig.  4.6 A, B, D‒F), and both germinal and ovarian cells have a mesothelial origin. Up to 50 oocytes were recorded in the female gonad, but only one small oligolecithal oocyte maturates, being further moved to the brood chamber (Brien & Mordant 1956, Wood 1983). Marcus (1934) reported the presence of nurse cells in the ovary of Lophopus crystallinus, but this aspect should be further checked. Pattern IV. Depending on the oocyte size and the role of placentation, the ovary structure and oogenesis in cheilostomes with this pattern correspond to that associated with patterns II or III (Ostrovsky 2013a, b). For example, ultrastructural studies showed that in Bicellariella ciliata, a vitellogenic doublet of a relatively small macrolecithal oocyte and its nurse cell is enveloped by squamous follicle cells as in species with pattern III (Moosbrugger et al. 2012). Their cytoplasm contains numerous mitochondria, multivesicular-like bodies, cisternae of rough endoplasmic reticulum, and yolk-like inclusions. Small vesicles fusing with the cell

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Fig. 4.5: Oogenesis in Cheilostomata (pattern II). (A‒C) Caberea boryi (histological sections). Zooids with functional polypide and ovaries on various stages of development, containing previtellogenic (A), early vitellogenic (B), and mature macrolecithal oocyte doublets (C). (D) Cribrilina annulata. Histological section of the ovary, containing previtellogenic and early vitellogenic oocyte doublets. Abbreviations: fo – follicle cells, nc – nurse cell, ov – ovary, p – polypide, pv – previtellogenic oocyte, rm – retractor muscles, sz - subovarian zone of translucent follicle cells, vo – vitellogenic oocyte.

membrane were detected and indicate exocytosis. The onset of vitellogenesis is detected by the appearance of yolk platelets in the cytoplasm and formation of microvilli in both the oocyte and its nurse cell. Some autosynthetic activity is present in both these cells, too, since multivesicular-like bodies and cisternae of the rough endoplasmic reticulum were found in them. Similar aspects of oogenesis and ovarian structure were recently described in another cheilostome Celleporella hyalina (Nekliudova et al. 2019a). Patterns V and VI. Data on the ovarian structure and oogenesis in the cheilostomes of the family Epistomiidae and in Cyclostomata are very scant. In contrast to all other bryozoans, Epistomia bursaria was described to have a “follicle” of “nurse” cells surrounding the oocyte, which is connected to it by cytoplasmic bridges

(Dyrynda & King 1982). Published illustration shows the oocyte being suspended in the central part of the spherical “follicle,” whose cells are distinctly different from the small oligolecithal oocyte. These cells were suggested to originate from the “germ cell” lineage, but this needs confirmation, because they could be of somatic origin as well. The presence of a nurse sibling was not reported. One (sometimes two) small oligolecithal oocytes develop inside the ovary in Cyclostomata, which is associated with a forming polypide bud. The ovary is small and consists of follicle cells enveloping the oocyte (Borg 1926, Reed 1991). Ultrastructural data are missing. Cinctipora elegans possesses a unique mode of oogenesis with oocytes appearing between the pharyngeal cells (Schwaha et al. 2018).



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Fig. 4.6: Oogenesis and brooding in Phylactolaemata (pattern III). (A, B, D, E) Plumatella fungosa (stained whole mounts). Ovary and brooding sac developing close to each other on the body wall. Young ovary in A, D, E and mature ovary in B. (C, F) Hyalinella punctata (histological sections). C, Two microlecithal oocytes (basal part of the ovary is out of the section plane). F, Oocyte in ovary, brooding sac with embryo, and polypide bud developing close to each other on the body wall. Abbreviations: bs – brooding sac, em – embryo, fc – follicle cells, o – oocyte, ov – ovary, p – polypide, pb – polypide bud, vo – vitellogenic oocyte.

4.5 Gamete release and fertilization Gonads lack gonoducts and gametes are released through coelomopores. Cross-fertilization dominates, although self-fertilization has been recorded in experiments. Sperm is released through terminal pores of the tentacles (all or on the two on the anal side) and enter fertile zooids via a supraneural coelomopore. Fertilization is intraovarian (sometimes precocious) or near-ovulatory (Temkin 1994,

1996, Ostrovsky 2008a). Up to 15 sperm were found in a single ovary, not counting those in previtellogenic or vitellogenic oocytes, in some brooding cheilostomes (Ostrovsky 2008a, 2013a). In the phylactolaemate Lophopus crystallinus, up to 150 spermatozoa may be contained in the ovary and up to 18 oocytes may be simultaneously fertilized (Marcus 1934). In Gymnolaemata, zygotes are released via a ciliated intertentacular organ or a similar tube-like organ called

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ovipositor (in spawners and some brooders) or supraneural coelomopore (in most brooders and some spawners). The presence of this pore needs yet to be confirmed in Cyclostomata (Stenolaemata). Phylactolaemata possess a vestibular pore participating in statoblast liberation, but whether it could be used as a conduit for sperm is unknown (Ostrovsky & Porter 2011). Karyogamy is always postponed until zygote release from the maternal zooid.

4.6 Embryonic incubation The vast majority of Bryozoa incubate their young being either brooders or viviparous. In the first case, the fertilized egg is usually deposited to a kind of marsupial chamber, whose cavity communicates with the external medium. Thus, this is a postparitive incubation. Marsupial structures are absent in some brooders, however. In the viviparous forms incubation is intracoelomic, i.e. preparitive. The studied phylactolaemates brood a single embryo inside an internal brood sac, which is an invagination of the body wall (Fig. 4.6 A, B, D‒F). The method of oviposition is unknown, however. The brood chamber develops near the ovary, and it is possible that the zygote moves from the gonad to the brood sac without being first released (Brien 1953, Reed 1991). Embryonic development is supported by matrotrophy. In contrast, cyclostomes are viviparous, which indicate an independent evolution of this mode of parental care. They incubate numerous embryos inside the voluminous polymorphic zooids (gonozooids) (Fig. 4.7), which are absent in the family Cinctiporidae, however. The primary embryo develops from a fertilized egg and further continuously buds off numerous secondary embryos developing to larvae (Fig.  4.7 E). Both embryonic multiplication and growth are accompanied by placentation. In Cinctipora elegans embryo budding (if present), growth and development might occur inside very large autozooids after polypide degeneration (Schwaha et al. 2018). Both viviparous and brooding forms are known among Gymnolaemata. Viviparity was only recorded in the cheilostome family Epistomiidae. The single larva develops inside an enlarged female polymorphic zooid in this case. Brooding has evolved several times independently in Ctenostomata. This is confirmed by the differences in the structure and position of the brood structures as well as distribution of the brooding forms on the ctenostome phylogenetic tree (Todd 2000). The simplest mode is external brooding of few to many embryos enveloped in

a fertilization envelope that is attached to the fertile zooid (e.g. Triticella flava, Panolicella nutans) (Fig.  4.8 E). In some species, embryos can be temporarily withdrawn to the vestibule during polypide retraction (e.g. Alcyonidium duplex, Bulbella abscondita). Another mode is the immersion of embryos into the vestibular (Fig.  4.8 C) or cystid (Fig.  4.8 D) wall (e.g. Tanganella muelleri, Sundanella sibogae, Nolella cf. papuensis). Two more modes of incubation in ctenostomes are brooding of one or several inside the modified tentacle sheath (Alcyonidium spp., Bowerbankia gracilis, Amathia verticillata, Bantariella cookae, Flustrellidra hispida) (Fig. 4.8 A, B, F) and inside a special outer embryo sac that is known among boring forms (Penetrantia) (Braem 1939, 1951, Ström 1977, Ostrovsky 2013a, Schwaha et al. 2019, Kvach et al. 2019). The most diverse brood chambers are met among Cheilostomata (Ostrovsky & Taylor 2005, Ostrovsky 2013a) (Figs. 4.9‒4.12). The three main groups are (1) external membranous sacs (Aetea, Eucratea loricata, “Carbasea” indivisa, Leiosalpinx australis); (2) skeletal (calcified) chambers, including all ovicells and brood chambers formed by spines and kenozooids (most cheilostomes); (3) internal brood sacs formed by non-calcified zooidal walls (in at least 26 families) (Cheetham & Cook 1983, Ostrovsky 2004, Ostrovsky 2008b, c, 2013a; Ostrovsky et al. 2008, 2009b, c). Brood chambers have evolved at least seven times in this clade. The nature of the external membranous brood sacs (Fig. 4.9 A, B) is yet to be uncovered. They can either be an outgrowth of the tentacle sheath wall, a cuticular chamber produced by the external cystid wall, or, similar to ctenostomes, a sticky fertilization envelope (Stach 1938, Cook 1977, Ström 1977). Else, they are known from species of four different cheilostome clades, which indicates their independent evolution. Skeletal incubation chambers include the acanthostegal brood chambers of Tendridae (Fig. 4.9 C) and ovicells and ovicell-like structures. The former consist of adjoining zooidal opesial spines, the frontal membrane, and the epistegal space between them (Repiachoff 1875, 1878, Paltschikova-Ostroumowa 1926, Braiko 1967, Ostrovsky & Taylor 2005). This type of skeletal brood chambers clearly evolved independently from the others. Conventional ovicells (Figs.  4.10 and 4.12) consist of a two-walled, totally or partially calcified, protective ooecial fold (ooecium) with an enclosed coelomic lumen, an ooecial vesicle (non-calcified part of the distal wall of the maternal autozooid that plugs the ovicell opening), and the brood cavity between them. The outer ooecial wall is termed ectooecium. It is separated from the entooecium (also often called endooecium), which surrounds the brood cavity, by a slit-like ooecial coelom



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Fig. 4.7: Gonozooids of Cyclostomata. (A) Entalophoroecia sp. (B) Telopora lobata. (C) Hornera sp. (D) Crisia sp. E, histological section of gonozooid of Crisia eburnea containing embryos and larvae embedded into a coenocytic placental analogue. Abbreviations: e – embryo, la – larva, os – ooeciostome.

(Fig. 4.10 D). Ooecium size and shape are subject to strong variation from large and hemispherical (Fig. 4.10 A‒D) to small (vestigial) and cap-like (Fig. 4.10 E, F). The calcified ectooecium can bear small cuticular areas (pseudopores) (Fig. 4.10 C) or larger membranous “windows” (Fig. 4.10

E). In some taxa, it is entirely membranous (Fig. 4.10 B). In many cheilostomes, it is overgrown by a secondary calcification produced by the neighboring zooids. The ooecium can be formed either by the daughter (distal) or maternal (egg-producing) zooid. In the former

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Fig. 4.8: Brooding in Ctenostomata. (A, B) Alcyonidium sp. Embryos incubated inside modified tentacle sheath (B, histological section). (C) Tanganella mülleri. Embryos (arrows) immersed in the vestibular wall. (D) Nolella cf. papuensis. Embryos immersed in the cystid wall. (E) Panolicella nutans. Two embryos (arrow) attached to the surface of cystid. (F) Mimosella sp. Embryo incubated inside modified tentacle sheath. Abbreviations: bb – brown body, p –polypide, ts – tentacle sheath, v – vestibulum.



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Fig. 4.9: Brood chambers in Cheilostomata. (A, B) Aetea sp. External membranous brood sacs (arrows) on the distal part of autozooids. (C) Tendra zostericola. Acanthostegal brood chamber in the distalmost zooid and non-brooding zooids. (D) Thalamoporella sp. Colony area with the ovicell-like bilobate brood chamber (arrow) containing three embryos.

case, the ooecium is an outfold of the daughter zooid frontal wall, and the ooecial coelomic cavity communicates with the zooidal coelom of the daughter zooid via communication pore(s), which are normally plugged by nonspecialized epithelial cells. The ooecium that is budded from the maternal zooid is a kenozooid, and the communication pores are plugged by the pore-cell complexes. In both cases, an ovicell can be considered as

complex colonial organ, formed by at least two zooids constituting a cormidium (reviewed in Ostrovsky 2013a). The ovicell opening is closed either by the zooidal operculum (in the cleithral ovicells), or by the ooecial vesicle (acleithral), or both (cleithral and semicleithral). The ooecial vesicle is a contractile fold, being operated by special muscle bands during larval release. Whereas cleithral ovicells raise their opercula during larval

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Fig. 4.10: Brood chambers in Cheilostomata. General view of conventional ovicells, empty, and containing embryos. (A) Copidozoum tenuirostre. (B) Fenestrulina malusii. (C) Caberea boryi. (D) Caulibugula sp. (ecto- and entooecium are well seen). (E, F) Costaticella sp. (frontal (left) and basal (right) views).

extrusion, it is lowered in some species and then termed subcleithral. In some species, ovicells are permanently open, however (non-cleithral) (Ostrovsky 2008c, 2013a). In the hyperstomial (prominent or raised) ovicells, half or more of the brood cavity is above the colony surface (Fig. 4.10 A‒D). If less than half the brood cavity

is above the colony surface, then the ovicell is termed subimmersed. In the immersed and endozooidal ovicells, the entire or near-entire brood cavity is situated below the colony surface. In endozooidal ovicells, the brood cavity is in the proximal part of the distal zooid, whereas in the immersed ovicells, the brood cavity is in



the distal part of the maternal zooid. In both types, the ooecium (usually vestigial) is slightly above the colony surface or is on the same level. The endotoichal ovicells found in the family Cellariidae are highly modified endozooidal ovicells. The development of a thick secondary calcification often transforms ovicells from hyperstomial/prominent to subimmersed, which often happens among ascophorine cheilostomes. One more ovicell type is the peristomial ovicell, in which the ooecium is incorporated into the zooidal peristome. Internal brood sacs (Fig. 4.11) are characterized by a totally or almost totally reduced protective ooecium and the brood cavity being immersed in the maternal zooid. The cavity is either connected with that of the vestibulum or opens independently of it to the outside (Ostrovsky et al. 2006, 2007, 2009c, Ostrovsky 2013a). In some taxa (e.g. family Adeonidae), internal brooding results in modified cystid shape and enlarged size, thus being a reason for sexual zooidal dimorphism (reviewed in Ostrovsky 2008c, 2013a).

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Whereas most of the brooding Cheilostomata possess so-called conventional ovicells, ovicell-like structures have evolved independently in a number of taxa. For example, Scruparia (Scrupariidae) has a high, helmet-shaped ooecium of two halves (supposed kenozooids), whose coelomic cavities are separated from each other. In Alysidium parasiticum (Alysidiidae), the ooecium is also bipartite. It consists of two hemispherical hollow plates capable of bending outward owing to their separate attachment to the maternal zooid by a cuticular base. These ooecial “valves” are true kenozooids, whose coeloms are separated from the visceral coelom of the maternal zooid by a pore plate each. A very complex brood chamber (synecium) of six flat plate-like kenozooids is formed in the genus Catenicula from the same family, Alysidiidae. Each kenozooid is attached to the maternal autozooid or neighbor kenozooid by an elastic cuticular joint, altogether forming a globular basket-like brood chamber. In Thalamoporella (Thalamoporellidae), the calcified ooecium of the ovicell-like cleithral brood chamber is formed by the fusion of two hollow hemispherical lobes

Fig. 4.11: Internal brooding in Cheilostomata. General view of colonies with zooids containing embryos in internal brood sacs. (A) Cranosina coronata. (B) Smittipora sp. (C) Steginoporella sp. Abbreviations: e – embryo, o – oocyte.

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originating from the frontal wall of the maternal autozooid (Fig. 4.9 D). Although the median suture is retained, a septum separates the lobes only at the ooecial base. Thus, the ooecium is not a kenozooid but a “bilobate” outgrowth of the frontal zooidal wall, whose common coelom communicates directly with the visceral coelom of the maternal autozooid via two large openings (Ström 1977, Ostrovsky 2013a). Unique ovicell-like brood chambers from costae have evolved in Bellulopora (Belluloporidae). Their ooecia consist of kenozooidal costae. These costae fuse with each other but leave pores that allow sea water to enter the cavity freely. The floor of the brood-cavity is uncalcified (Ostrovsky & Taylor 2005). Acanthostegal brood chambers of Tendridae, conventional ovicells, and the ovicell-like brood chambers of Bellulopora all originated independently from opesial spines. In all these cases, the first step was bending of the originally jointed spines to form a brood space, including loss of their articulation (basal joints). In tendrids, spines were eventually closely opposed to each other, whereas they were fused by lateral outgrowths in Bellulopora. In both cases, however, the brood cavity was not isolated from the ambient water. The evolution of conventional ovicells was probably associated with the reduction of ooecial spines to two, accompanied by their flattening and enlargement. Finally, these two flattened lobes were able to finally fuse to the complete ooecium. In cribrilinid cheilostomes, the left and right ooecial halves can originate from the fusion of their spines without previous reduction in number (Ostrovsky & Taylor 2004, 2005, Ostrovsky 2013a). Finally, it is reasonable to assume that all other ovicell-like brood chambers have evolved from kenozooids and various modifications of spines or spine-like protuberances, articulated or not, via their flattening and apposition. Interestingly, in contrast to most cheilostome brooders, those cheilostomes that evolved such chambers independently brood several embryos simultaneously – in the genera Scruparia, Tendra and Thalamoporella as well as in Macropora and Monoporella (the latter two having conventional ovicells). The viviparous Epistomiidae presumably evolved from a brooding ancestor (Ostrovsky 2013a).

4.7 Matrotrophy and evolution of reproductive patterns Bryozoa is an outstanding animal group having extraembryonic nutrition in all major classes (Ostrovsky et al. 2009a, Ostrovsky 2013a). Also, considering the number of

families that completely consist of or include matrotrophic species, this phylum concedes only chordates, plathyhelminthes, and arthropods (Ostrovsky et al. 2016). All studied Phylactolaemata are matrotrophic. Although the nutritive role of spot- and ring-like contact zones (so-called “placentae”) between embryo and the wall of the brood sac is not proven yet, the progeny greatly increases in size during development (Braem 1890, 1897, 1908). Recent ultrastructural analysis showed evidence of endocytosis by the cells of the brood sac presumably absorbing nutrients from the coelomic fluid in this group (Bibermair et al. in prep.) All studied cyclostomes are matrotrophic, too. Growing embryos develop inside gonozooids surrounded by a coenocytic placental analogue (Fig.  4.7 E), which participates in the synthesis and transport of nutrients. This nourishing structure partly originates from the modified membranous sac – the hydrostatic apparatus of the cyclostome autozooid (Borg 1926, Nekliudova et al. in prep.). Noteworthy, placentation could be a necessary prerequisite to trigger the evolution of cyclostome polyembryony, which in turn could stimulate the origin of gonozooids. Also, it is feasible that multiplication of embryos supported by a placenta exists in Cinctiporidae, and the lack of swollen gonozooids was compensated by the gigantic size of their autozooids, which could act as incubation chambers. The small size of the oocytes in Cinctipora elegans compared with the large primary disc of the ancestrula indicates considerable embryo growth during incubation, which could be supported by matrotrophy as in the rest of cyclostomes (Schwaha et al. 2018). Among Ctenostomata, embryo size increase was detected in some species of the genus Nolella that brood their progeny inside invaginations of the body wall (Hincks 1880, Prouho 1892, Harmer 1915). Hypertrophy of the cells in the wall of such brood chamber was recorded in Sundanella sibogae (Braem 1939) and Labiostomella gisleni (Silén 1944, although the systematic position of this species is unclear). Embryos being brooded inside the tentacle sheath also enlarge in four species from four different ctenostome clades (Joliet 1877, Bogoiavlenskii 1905, Pace 1906, Waters 1914, Zirpolo 1933, Banta 1968). Ultrastructural evidence of both exocytosis by the cells of the placental analogue and endocytosis/pinocytosis by embryonic cells has been recently obtained in Amathia verticillata (Schwaha et al. 2019). Among Cheilostomata, the viviparous Epistomiidae are matrotrophic. The source of the placental analogue is unclear (Dyrynda & King 1982) and potentially could be a modified follicle (see also Ostrovsky 2013a). In cheilostome brooders, the placental analogue (Fig. 4.12 B, C, E) is a hypertrophied epithelium of the body wall – either an ooecial vesicle or internal brood sac – associated with



4.7 Matrotrophy and evolution of reproductive patterns 

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funicular cells. In Celleporella hyalina, these cells also greatly multiply, forming a “tissue-like organ” (Nekliudova et al. 2019a) (Fig. 4.12 F). In fact, the main structural principles of the placental analogues are the same among Bryozoa. Incubated embryos are surrounded, totally or partially, by temporarily hypertrophied nourishing cells or a coenocyte (in cyclostomes).

A hypothetical scenario of the main steps of the origin of placentotrophy in gymnolaemate bryozoans suggests that the evolution of new reproductive patterns included transitions from reproductive pattern I to pattern II, from pattern II to pattern IV, and further from pattern IV to pattern III (Ostrovsky et al. 2009a, Ostrovsky 2013a, b). In this line of thought, two shifts in oogenesis ‒ from oligo- to

Fig. 4.12: Matrotrophic brooding in Cheilostomata. (A) Bugulina fulva. Frontal view of the colony branch with ovicells containing growing embryos (arrows) of various sizes. (B‒D) Bicellariella ciliata. Zygote (C) and late embryo (B) in the ovicell. Embryophore (placental analogue) is well seen in B. (D) Lateral view of the colony branch with ovicells containing growing embryos of various sizes. (E) Bugula sp. Histological section of the colony branch with ovicells containing growing embryos of various sizes. Embryophore is shown by arrows. (F) Celleporella hyalina (histological section). Female zooid with the ovicell containing a late embryo. A large embryophore is apposed to the embryo on the left side. Abbreviations: a – ascus, e – embryo, ep – embryophore, fw – frontal wall, oe – ooecium, op – operculum, ov – ovary.

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macrolecithal (from pattern I to II) and back (from IV to III) – would have occurred. The origin of placentotrophy could trigger the latter shift in oogenesis. Finally, the distribution of reproductive patterns across Bryozoa suggests that placentotrophy evolved independently in all three bryozoan classes and within both gymnolaemate orders. Also, considering the differences in the incubation mode and the current position of matrotrophic ctenostomes on the phylogenetic tree (Todd 2000), extraembryonic nutrition could have evolved five times in this order. Among Cheilostomata it has evolved at least 16 times (Ostrovsky et al. 2009a, Lidgard et al. 2012, Ostrovsky 2013a, b).

4.8 Life histories Different bryozoan species have contrasting life-history traits such as colonial sexual structure, number, timing, and duration of the reproduction periods, peaks of larval settlement and tempo of colonial growth, annual number of generations, their longevity and participation in larval production. Several approaches have been developed to classify bryozoan life-histories. Based on year-round observations on Mediterranean bryozoans, Gautier (1962) distinguished: (1) species reproducing in winter, (2) species reproducing in summer, and (3) eurythermic species reproducing year-round (see also Médioni 1972). Gordon (1970) divided New Zealand species into three groups depending on the duration of the reproductive period, defining (1) species with short reproductive period, (2) year-round reproducing species slightly decreasing reproductive activity in winter, and (3) species reproducing from early spring until early winter. Kuznetsov (1941) classified fouling organisms (including bryozoans) on subtidal stones in the Barents Sea based on the number of generations formed during the year. Accordingly, he distinguished species having one, two, or three generations (spring, summer or winter). In fact, this referred to the number (and timing) of reproductive periods in the life of one generation in a year and did not consider coexisting generations. Kuznetsov determined that bryozoan colonies that were established in different seasons may have different lifespans and therefore a different number of reproduction periods. Eggleston (1963, 1972) classified bryozoans in the Irish Sea (near the Isle of Man) using the colony lifespan and the number of generations per year, also considering the terms of their reproduction. Among the species with colonies

living less than 1 year, he distinguished (1) short-lived species whose colonies live no longer than a few months (according to Eggleston, a precise number of generations cannot be distinguished with certainty in populations that release larvae and form young colonies year round); and (2) species with two generations each year (overwintered colonies reproduce in May–June and die in August; colonies of the second generation reproduce in August– September and die in November, while their descendants overwinter). Among the species with colonies living for at least one year, Eggleston first distinguished the annuals and biennials that reproduce once and die afterward or overwinter and take part in larval production next year. So-called perennials are species whose colonies live at least two years, having either seasonal peaks of larval production or reproducing year-round. Besides the above classifications, ecological division of species into r- and K-strategists is often used. These two strategies are merely two extreme variants, with most instances belonging to an r-K continuum (Reznick et al. 2002). Attempts to attribute bryozoan species to these strategists were undertaken by Dyrynda and Ryland (1982), Seed and Hughes (1992), and Harmelin and Aristegui (1988), although some authors regard this theory as not being applicable to modular (colonial) organisms that combine r- and K-selected traits (Jackson & Coates 1986, McKinney & Jackson 1989, Seed & Hughes 1992). In bryozoans, the r-strategy operates with fast-growing colonies consisting of weakly calcified zooids and forming numerous small oocytes that develop into planktotrophic larvae without parental care (Eggleston 1963, Seed & Hughes 1992). Such European species, for example Membranipora membranacea and Electra pilosa, frequently dominate on laminarian fronds. Recently, the life histories of five gymnolaemates with contrast reproductive strategies (broadcasting and brooding, lecithotrophic, and matrotrophic) were described using colony mapping (Nekliudova et al. 2019a, b, Kvach et al. 2019, Shevchenko et al. 2020). Based on pioneering works of Borg (1947) and Dyrynda and Ryland (1982), they showed that bryozoan life-history traits are much more diverse than acknowledged and show much flexibility.

Acknowledgments Thanks to the Austrian Science Fund (grants no. P22696-B17 and P27933-B29) and Russian Science Foundation (grant no. 18-14-00086) for financial support.

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Rogick, M.D. (1956): Studies on marine Bryozoa. VII. Hippothoa. Ohio J Sci 56: 183–191. Schwaha, T., Handschuh, S., Ostrovsky, A.N. & Wanninger, A. (2018): Morphology of the bryozoan Cinctipora elegans (Cyclostomata, Cinctiporidae) with the first data on its sexual reproduction and the cyclostome neuro-muscular system. BMC Evol Biol 18: 92. Schwaha, T., Moosbrugger, M. & Ostrovsky, A.N. (2019): First ultrastructural evidence of placental nutrition in a ctenostome bryozoan: example of Amathia verticillata. Zoomorphology 138: 221–232. Schwaha, T.F., Ostrovsky, A.N. & Wanninger, A. (2020): Key novelties in the evolution of Bryozoa: evidence from the soft-body morphology. Biol Rev 95: 696–729. DOI: 10.1111/brv.12583 Seed, R. & Hughes, R.N. (1992): Reproductive strategies of epialgal bryozoans. Invertebr Reprod Dev 22: 291–300. Shevchenko, E.T., Varfolomeeva, M.A., Nekliudova, U.A., Kotenko, O.N., Usov, N.V., Granovitch, A.I. & Ostrovsky, A.N. (2020): Electra vs Callopora: life-histories of two bryozoans with contrasting reproductive patterns from the White Sea. Invertebr Reprod Dev 64: 137–157. DOI: 10.1080/07924259.2020.1729260. Shevchenko E.T., Matvienko D.A., Nekliudova U.A., Belikova E.V., Kotenko O.N., Ostrovsky A.N. (in preparation): Comparative ultrastructural study of oogenesis in cheilostome bryozoans with contrasting reproductive patterns and its evolutionary implications. Zoology. Silén, L. (1944): The anatomy of Labiostomella gisleni Silén (Bryozoa Protocheilostomata). Kungl Svenska Vetensk Akad Handl Ser 3 21: 1–111.

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Stach, L.W. (1938): Observation on Carbasea indivisa Busk (Bryozoa). Proc Zool Soc Lond B 108: 389–399. Ström, R. (1977): Brooding patterns of bryozoans. In: Woollacott, R.M. & Zimmer, R.L. (eds.). Biology of Bryozoans. Academic Press, New York: 23–56. Temkin, M.H. (1994): Gamete spawning and fertilization in the gymnolaemate bryozoan Membranipora membranacea. Biol Bull 187: 143–155. Temkin, M.H. (1996): Comparative fertilization biology of gymnolaemate bryozoans. Mar Biol 127: 329–339. Todd, J.A. (2000): The central role of ctenostomes in bryozoan phylogeny. In: Herrera Cubilla, A. & Jackson, J.B.C. (eds.). Proceedings of the 11th International Bryozoology Association Conference. Smithsonian Tropical Research Institute, Balboa: 104–135. Waters, A. (1914): The marine fauna of British East Africa and Zanzibar, from collections made by Cyril Crossland, M.A., B.Sc., F.Z.S., in the years 1901–1902. Bryozoa-Cyclostomata, Ctenostomata and Endoprocta. Proc Zool Soc Lond 84: 831–858. Wood, T.S. (1983): General features of the class Phylactolaemata. In: Robinson, R.A. (ed.). Treatise on Invertebrate Palaeontology. Part G: Bryozoa (Revised). Geological Society of America and University of Kansas, Boulder and Lawrence: 287–303. Woollacott, R.M. (1999): Bryozoa. Encyclopedia of Reproduction 1: 439–448. Zimmer, R.L. & Woollacott, R.M. (1977): Structure and classification of gymnolaemate larvae. In Woollacott, R.M. & Zimmer, R.L. (eds.). Biology of Bryozoans. Academic Press, New York: 57–90. Zirpolo, G. (1933): Zoobothryon verticillatum (Delle Chiaje). Mem Accad Nuovi Lincei 17: 109–441.

Alexander Gruhl

5 Larval structure and metamorphosis 5.1 Introduction

5.2 Embryonic development

Sexual reproduction in bryozoans always involves free-swimming larval stages (Fig. 5.1). Fertilization is internal, broadcast sperm are taken up by the recipient zooids, and developing embryos are either brooded in or at the zooid or in specialized colonial brood chambers (often derived zooids). Lecithotrophic larvae either develop from macrolecithal eggs or receive extraembryonic nutrition. After release from the parent colony, they spend only a short period of time in the water column before settling on the substrate. Planktotrophic larvae – the iconic cyphonautes (Fig. 5.1 A) – occur only in a few gymnolaemate species, which, however, are usually distributed across wide geographical ranges due to the larvae’s prolonged pelagic phase (between weeks and months). These species shed numerous fertilized oligolecithal eggs directly into the water column, where the entire embryonic development takes place. Gymnolaemate (cheilostome and ctenostome) bryozoan larvae show considerable variation in terms of both their lifestyles and morphologies, but a common bauplan with a specific set of organs can be identified (Fig. 5.2 A–C). Stenolaemate (cyclostome) larvae also fit into this basic pattern but have a reduced size and morphological complexity, lacking several of the gymnolaemate larval organs (Fig. 5.2 E). Phylactolaemate larvae (Figs. 5.1 C and 5.2 D), however, differ significantly from the previous types. Many authors regard them as having evolved independently in the phylactolaemate lineage and prefer to call them “swimming ancestrulae”. Bryozoan larvae have been subject to extensive research, beginning with early 19th century. Among the first researchers to describe bryozoan larvae were Grant (1827) for marine and Allman (1847) for freshwater bryozoans. Larger monographic accounts include Barrois (1877, 1880), Prouho (1892), Braem (1897, 1908), and Calvet (1900). More recent general reviews and introductory texts on bryozoan reproduction and larval biology are found in Ryland (1976), Woollacott and Zimmer (1977), Zimmer and Woollacott (1977a, b), Mukai (1982), Mukai et al. (1997), and especially Reed (1991).

Bryozoan embryonic development differs tremendously between the classes. It has been followed in most detail in gymnolaemate species (Barrois 1877, Repiachoff 1878a, b, Vigelius 1886, Prouho 1892, Braem 1896, Calvet 1900, Pace 1906, Marcus 1926a, Corrêa 1948, Zimmer 1997, Gruhl 2010a, Vellutini et al. 2017), with most data coming from observations on free-spawners with a cyphonautes larva, but some also from species with macrolecithal eggs and lecithotrophic larvae. While most studies do not extend beyond the 64-cell stage, Gruhl (2010a) provided ultrastructural data from postgastrulation stages and Vellutini et al. (2017) conducted the first highly detailed cell lineage study based on four-dimensional microscopy in Membranipora membranacea. In all examined gymnolaemate species, a consistent pattern, termed biradial cleavage (Fig. 5.3 A), has been described. The first two cleavages are meridional and equal; the third cleavage is equatorial, resulting in larger and more yolk-rich vegetal cells (macromeres). The planes of the fourth cleavage are parallel to that of the first one resulting in a characteristic biradial 16-cell stage with two tiers of two by four cells (Fig. 5.3 A, G). In the vegetal layer, the four central cells are larger than the peripheral ones: two equatorial divisions in the animal octet, an unequal division of the vegetal macromeres, and a further division of the vegetal micromeres lead to a 48-cell embryo. At this stage, a quartet of four large macromeres in the vegetal hemisphere is surrounded by 12 micromeres. In the animal hemisphere, two tiers, each consisting of two by four micromeres, are separated from the vegetal hemisphere by an equatorial ring of 16 micromeres, which later form the corona. This pattern is highly stereotypic in M. membranacea (Vellutini et al. 2017). At later stages, asynchronous divisions with shifted cleavage planes break the biradial symmetry of the embryo. This process is probably initiated by the specification of the D quadrant, which begins at the 28-cell stage by activation of the MAPK pathway. Thus, the embryonal quadrants A and C correspond to the left and right body sides, respectively, and B and D to anterior and posterior, respectively. The embryonal animal-vegetal axis corresponds to the larval dorso-ventral (oral-aboral) axis (Vellutini et al. 2017).

https://doi.org/10.1515/9783110586312-005

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Fig. 5.1: Examples of bryozoan larvae. (A) Cyphonautes larva from plankton sample. (B) Coronate larvae of Schizoporella unicornis (Gymnolaemata, Cheilostomata). (C) Larva of Plumatella repens (Phylactolaemata). (D) Depiction of various cheilostome coronate larvae (from Calvet 1900). (E) Depiction of cyphonautes and shelled lecithotrophic larvae (from Barrois 1877). Scale bars: A–C, 200 µm.

Gastrulation involves the internalization of the central vegetal quartet of macromeres. This happens in part by epiboly (inward directed division of the surrounding micromeres) and in part by delamination (several unequal divisions of the macromeres and resulting 4t­­h micromeres). The cells surrounding the central macromeres form the blastopore rim. Blastopore closure has been reported in some species with lecithotrophic larvae, but in M. membranacea, the blastopore develops into the definitive mouth (Gruhl 2010a) (Fig. 5.3 I). Protostomy is also supported by characteristic expression patterns of endoderm

and foregut marker genes (Vellutini et al. 2017). The origin of the mesoderm is still not totally clarified. The first mesodermal cells become visible at the anterior end of gastrula stages, eventually forming the neuromuscular strand and other parts of the larval musculature. While Gruhl (2010a) documented an ingression of an ectodermal cell, Vellutini et al. (2017) found the anterior mesoderm to originate from 4a–c micromeres produced during gastrulation from the internalized macromeres, thus from endoderm. The origins of further mesodermal tissues, especially the blastemata that underlie internal sac and apical organ, are unclear.

▸ Fig. 5.2: Schematic representations of bryozoan larval morphologies. (A) Generalized gymnolaemate larva. (B) Gymnolaemate cyphonautes larva. (C) Gymnolaemate coronate larva. (D) Phylactolaemate larva. (E) Cyclostome larva. Abbreviations: ae – ­aboral epithelium, ai – aboral invagination, an – anus, ao – apical organ, ap – apical plate, at – atrium, bc – body cavity, cc – ciliated cleft, co – corona, cr – ciliated ridge, cy – ancestrular cystid, in – intestinal tract, is – internal sac, mf – mantle fold, mo – mouth, nm – neuromuscular strand, nn – nerve nodule, np – neural plate, oe – oral epithelium, pl – ancestrular polypid, po – pyriform organ, ps – pallial sinus, sh – shell, ve – vestibular pore, vp – vibratile plume.



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Postgastrulation development obviously differs between feeding and non-feeding larvae. Cyphonautes larvae take up active swimming once the corona and the apical organ are developed and their cilia break through the fertilization envelope (Fig. 5.3 J–L). The next steps in development include involution of the oral ectoderm to form the atrium, formation of ciliated ridges and other particle feeding structures, formation of the anus, differentiation of the gut, and formation of the lateral shells. The internal sac is the last structure to be completed – usually when larvae reach their full size and metamorphic competence. In starving larvae, the internal sac can also get reduced again resulting in loss of metamorphic competence (Strathmann et al. 2008). Non-feeding larvae may develop from macrolecithal eggs or receive extraembryonic nutrition during development (Mukai et al. 1997, Ostrovsky et al. 2009). However, these are two extremes and many combinations of yolk provision via the oocyte or via placental structures have been found (Ostrovsky 2013). Most lecithotrophic larvae completely lack a gut and feeding structures; thus, the oral surface does not involute or differentiate feeding structures. Shells are usually absent. Initially, gastrulation is comparable to that in feeding larvae – the vegetal macromeres are internalized as endodermal cells; however, in most species, they do not form a gut, but mesenchymal tissues. Few species (e.g. of genera Flustrellidra, Alcyonidium) develop a rudimentary, nonfunctional gut without clear blastopore or anal opening (Barrois 1877, d’Hondt 1977a, Gruhl 2009) (Fig. 5.3 N–Q). All non-feeding larvae emerge fully grown and differentiated and are more or less immediately competent for settlement and metamorphosis. Embryonic development in Cyclostomata and Phylactolaemata differs from that of Gymnolaemata and is, mainly due to technically challenging fixation and histological preparation, much less studied. All known phylactolaemates and cyclostomes are brooders; no free spawners with plankotrophic larvae have been described in these taxa. Cyclostome development has been studied by Harmer (1898), Calvet (1900), Robertson (1903), Borg (1926), and Nielsen (1970, 1971). It is dominated by polyembryony (vegetative division of primary embryos) (Fig. 5.3 C) and takes

5.2 Embryonic development 

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place in specialized, so-called gonozooids (but see Schwaha et al. 2018 for a potential deviation from this pattern). The zygote undergoes irregular cleavages, according to some reports even disintegration and subsequent reintegration, to form a primary embryo, which grows by receiving extraembryonic nutrition and buds off smaller secondary embryos. The latter are spherical, possibly bilayered, and develop both an apical and vegetal tissue invagination, which might correspond to gymnolaemate pallial sinus and internal sac, respectively. The outer surface becomes ciliated. Other organs known from gymnolaemate larvae, such as apical organ and pyriform organ never develop. Phylactolaemate development has been studied by, e.g., Davenport (1890), Kraepelin (1892), Braem (1897, 1908), Marcus (1934), Oka and Oda (1948), and Brien (1953). The embryos are brooded in so-called embryo sacs – invaginations of the body wall into the body cavity (Fig. 5.3 B). The small zygote undergoes irregular cleavages inside the embryo sac, developing a distinct polarity with the pole facing the maternal body wall referred to as distal, the opposite one as proximal. The distal pole seems to have more mitotic activity and represents the “growing edge” (Brien 1953). The embryo elongates toward the body wall and its proximal end becomes hollow (Fig. 5.3 M). At some stage, the embryo becomes bilayered, but reports on the origin of the inner layer give contradictory results. In many species, the bilayered embryo has a belt-like central constriction, which seems to be the contact point of a ring-like placenta structure. During later development, one to four polypide buds begin to form at the distal end and the whole outer surface becomes ciliated. Bud formation in larvae does not differ from the same process in adult colonies. Cleavage pattern, early blastomere fates, and mesoderm origin in gymnolaemates show many similarities to other lophotrochozoan embryos with spiral cleavages (Nielsen 2005, Gruhl 2010a), but later blastomere fates differ (Vellutini et al. 2017). Same is true for gene expression patterns, where certain elements are similar between bryozoans and spiral-cleavers (Fuchs et al. 2011). At the moment, it is difficult to decide whether bryozoans show an altered or reduced spiral cleavage or whether spiral

◂ Fig. 5.3: Bryozoan embryonic development. (A) Biradial cleavage in gymnolaemates (based on Corrêa 1948). (B) Phylactolaemate embryo in embryo sac attached to inside of maternal zooid’s body wall (based on Brien 1953). (C) Polyembryony in Cyclostomata (based on Robertson 1903). (D–L) Development of Membranipora membranacea (from Gruhl 2010a). (M) Embryo of Plumatella repens (Phylactolaemata), dissected out of maternal zooid. (N–O) Developmental stages of Flustrellidra hispida (Gymnolaemata, “Ctenostomata”). Abbreviations: ao – apical organ, at – atrium, bc – body cavity (= coelom), bd – polypide bud, bp – blastopore, ce – coelomic epithelium, co – corona, cr – ciliated ridge, dp – distal pole of embryo, du – duplicature, ed – ectodermal part of embryo sac, em – embryo, en – endocyst (=epidermis), fm – fertilization membrane, gz – gonozooid, in – intestinal tract, is – internal sac, mo – mouth opening, pb – polar body, pe – primary embryo, po – pyriform organ, pp – proximal pole of embryo, sa – sperm axoneme, se – secondary embryos. Scale bars: D–L, 50 µm; M, 200 µm; N–Q, 100 µm.

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and biradial cleavages evolved independently from an ancestral form that had only the basic similarities. This would ultimately also depend on the phylogenetic position of bryozoans with respect to spiralians (Hejnol 2010, Nielsen 2012, Laumer et al. 2015).

5.3 Larval morphology 5.3.1 General The main larval organs in gymnolaemate larvae include (1) the corona, which is the main ciliary band; (2) the apical organ, a complex of sensory and motory ciliated cells; (3) the internal sac, an invagination of the oral ectoderm facilitating contact with the substrate during settlement; (4) the pyriform organ, a complex of sensory, glandular, and locomotory cilia; (5) the digestive tract; (6) a neuromuscular strand connecting apical and pyriform organ; (7) a nerve nodule; and (8) blastemata (Fig. 5.2 A–C) It is difficult to correlate bryozoan larval body axes with those of adults, and even in the latter, it is controversial whether terms like dorsal and ventral make sense. For larvae, the following terminology was introduced by Zimmer and Woollacott (1977a): the corona divides the larva into two hemispheres; the one with the mouth opening, internal sac, and pyriform organ, is called oral, and the one with the apical organ is called aboral. The site of the pyriform organ determines the anterior end, whereas the opposite site, which includes the anus in planktotrophic larvae, is called posterior. The most prominent uniting feature of all gymnolaemate larvae is the corona – a row of ciliated cells, which, in a generalized larva, is almost equatorial. In many species, it consists of 32 cells, but higher numbers ( B > C etc.; Russ 1982, Lopez Gappa 1989, Barnes 2002, Centurion & Lopez Gappa 2011), or competitive networks (A > B > C, but C > A, etc.). Competitive networks such as those found in cryptic coral undersurfaces may lead to greater diversity in community

development on such patchy substrata where there is not a single dominant competitor that can monopolize a substratum (Buss & Jackson 1979, Karlson & Buss 1984, Taylor 2016). Fig. 6.24 D shows a coral undersurface on a Jamaican reef on which the actively budding frontal lobes of a Steginoporella colony are growing over an encrusting orange sponge, but at a senescent part of the Steginoporella colony, the sponge is winning. Competitive hierarchies may be more frequent at higher latitudes (Lopez Gappa 1989, Barnes & Rothery 1996, Barnes 2002). Phylogenetic constraints also matter in spatial competition. Encrusting cyclostomes have only two ways to block overgrowth by competing cheilostomes, elevation of the growing margin, and budding of subcolonies above the surface of the original colony. Stebbing (1972) found that colonies of Disporella hispida and Plagioecia patina, whose normal calcareous colony margin is adherent to the substratum, sometimes developed raised margins. The first type was found in isolated colonies; second type occurred in colonies near another sessile organism. In Adriatic cyclostomes studied by McKinney (1993)

6.6 Competition 

Diplosolen obelium was able to temporarily block cheilostome overgrowth in 10 of 120 encounters and Plagioeocia patina in 17 of 62 encounters, perhaps allowing the cyclostome colony time to reproduce and disperse larvae. Only Stebbing (1973) studied competition involving encrusting ctenostomes. Three of the four species found on Fucus serratus in Plymouth, England, were ctenostomes, Flustrellidra hispida (with chitinous spines), Alcyonidium hirsutum (also spiny), and Alcyonidium polyoum; the fourth was a spiny cheilostome, Electra pilosa. In most encounters between ctenostome species, forward growth stopped when colonies met (but was often diverted, continuing to fill in available bare space in another direction). Electra pilosa was frequently overgrown by the ctenostomes present, although, as its larvae tended to settle on the younger parts of fronds (Stebbing 1972), it could still control a fair amount of algal surface. In the few cases where Electra overgrew Flustrellidra, its normal sheet-like growth pattern was disrupted by the Flustrellidra spines and grew as narrow extensions between spines (Plate 4b in Stebbing 1973). The spines of Electra pilosa may sometimes have defended a colony margin against overgrowth by Alcyonidium polyoum, but they also occurred at nonthreatened margins where new zooids were being laid down (Fig. 2 in Stebbing 1973).

6.6.2 Feeding interference Another means to overcome competitors is by feeding interference. In laboratory studies of two encrusting species of Pacific cheilostomes, Buss (1981a, b) found that Onychocella alula, which had larger tentacle crowns and stronger feeding currents than competitor Antropora tincta, was able to capture marginal feeding currents from the Antropora colonies. But differing rates of flow can result in augmented feeding in bryozoan colonies, as shown by Okamura (1988). At low flow, feeding of Electra pilosa colonies was reduced when Alcyonidium hirsutum colonies were upstream, but in rapid flow, feeding was enhanced when Electra colonies were downstream of Alcyonidium and enhanced even more when the Electra colony was surrounded by Alcyonidium colonies. In Conopeum reticulum (Okamura 1985), feeding was enhanced by the presence of an actively feeding conspecific colony upstream.

6.6.3 Fighting buds Elongate stolon-like extensions of marginal zooid buds may also be deployed to overgrow a competitor (Fig. 6.24 C).

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This is shown in the Brazilian species Schizoporella pungens, a member of the Schizoporella errata group. A Schizoporella colony on panels being held in tanks at CEBIMar in Brazil was photographed in a battle with Hippopodina feegeensis. The long fingerlike protrusions of the marginal buds extended from the growing margin of the Schizoporella onto the surface of an adjacent competing portion of the Hippopodina colony (see enlarged area of Fig. 6.24 C). The use of such projecting outgrowths has also been documented in the Pacific Ocean species Stylopoma duboisii overgrowing Thalamoporella tubifera (Osborne 1984). Other encrusting cheilostomes with giant buds at the colony margins may be capable of this type of competitive behavior.

6.6.4 Molting Another form of defense against fouling and settlement is found in the free-living cupuladriids. These colonies, found on the surface on sandy bottoms, are subject to fouling by epibionts, especially microscopic algae, as well as to settlement by other bryozoans (e.g. Beania cupulariensis, Alcyonidium capronae, and entoprocts). The sweeping movements of the vibracula found distal to each zooid form a primary deterrent, but in late summer, Florida colonies of Cupuladria doma and Discoporella depressa were also found to molt their fouled frontal membranes. As the old membranes peeled off, pristine new membranes and opercula were revealed beneath. In Cupuladria doma, the extrazooidal colony base also molts its entire membrane (Winston & Håkansson 1989). Similar molting of frontal membranes has also been observed in a coral reef benthic encrusting species Crassimarginatella tuberosa from Belize and in a West African ctenostome, Alcyonidium sanguineum (Cook 1985). It may be more widespread among anascans and encrusting ctenostomes, especially in warm water habitats where seasonal cycles of activity and overwintering dormancy do not occur or are not as intense.

6.6.5 Chemical defenses Like other sessile organisms, bryozoans are capable of chemical defenses against potential predators and competitors, although less is known about their occurrence and use than for sponges and ascidians, the groups from which most biomedically important compounds have been derived. Sharp et al. (2007) reviewed the chemical ecology of bryozoans from an ecological perspective, noting that most of the relatively few studied so far were large, easily

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recognizable foliose species, abundant enough to collect the large quantities needed for natural product extraction. Encrusting colonies, with the need to prevent fouling of surfaces, offer great opportunity to find inhibitors of the kinds of predation that affect their colonies. Many of the secondary metabolites found so far do not come directly from the bryozoans, but from their microbial endosymbionts. Prinsep (2018) reviewed research on compounds isolated from bryozoans, especially those from the Australasia region. Only about 1% of bryozoans have so far been studied chemically. Types of compounds found include alkaloids, especially brominated alkaloids, such as amathamides (from Amathia wilsoni), polyketides (including the bryostatins), sterols, ceramides, peptides, and terpenes. A study of antibacterial activity in four species of bryozoans from Tasmania (Walls et al. 1993) showed that two species known to contain antibacterial secondary metabolites, Amathia wilsoni and Orthoscuticella ventricosa, had the lowest levels of fouling, and the two with no known active compounds, Cellaria pilosa and Bugularia dissimilis, showed only weak antibacterial activity and were heavily fouled. Many of the compounds found may not come directly from the bryozoans, but from the endosymbiotic microorganisms associated with them. Researchers have also sought chemical activity in Antarctic bryozoans. Winston and Bernheimer (1986) found moderate hemolytic activity in extracts of Carbasea curva, a weakly calcified Antarctic bryozoan with no physical deterrents against fouling or predation. The four other species tested had heterozooids ranging from spines to avicularia and vibracula and showed no cytotoxic activity. Figuerola et  al. (2014) analyzed 32 organic extracts from 13 Antarctic bryozoan species. Compounds found included both lipophilic and hydrophilic bioactive portions. In substratum preference assays, the chemical activity of 10 species was found to repel the omnivorous amphipod Cheirimedon femoratus. Amphipods are often found in large numbers living in Antarctic bryozoan colony masses (Winston & Heimberg 1988). Active compounds in 12 of 13 species at natural concentrations reduced the reproductive success of the sea urchin Sterechinus neumayeri, a grazer on bryozoans and other benthos, by affecting sperm viability (Figuerola et  al. 2014). Clearly, there is much more scope for research into chemical and ecological interactions of Antarctic bryozoans. A few studies (Lopanik et al. 2004, 2006) have focused on the chemical defenses of bryozoan larvae. Bugula neritina larvae are defended by high concentrations of bryostatins, cyclic polyketides produced by microbial endosymbionts and extracts deter pinfish predation, as

do extracts of early juveniles. However, extracts of adult colonies are palatable to fish. Extracts of the brooding portions of colonies also reduced fish predation by 54%, but those of non-brooding colonies were significantly (20%) less defended. Bugula neritina colonies have no avicularia; authors suggested that the physical structure of colony zooids might be enough of a deterrent, but the skeleton in this species is relatively light. In contrast, Bryan et  al. (1998) described compounds from Bugula neritina that caused larvae of the serpulid polychaete Hydroides elegans to metamorphose. They suggested that both metabolites of the bryozoan and its complex structure offer a refuge from predation to the polychaetes.

6.7 Interactions with other organisms Zooid level predation may well be the most important factor determining bryozoan evolution (Lidgard et  al. 2011). However, bryozoans interact with other organisms in both negative and positive ways, as Fig. 6.25 shows. Lidgard (2008a, b) reviewed the types of predation undergone by bryozoans and categorized 399 taxa involved. Mortality among young colonies is high, but as colonies grow bigger, the risk of lethal predation by large predators with broad diets (e.g. certain fish, decapods, echinoderms, chitons, and macro-gastropods) that graze, abrade, or bite to destroy large areas of colonies is lessened. However, the amount of sublethal predation by single zooid predators increases. Single zooid predators are small organisms themselves, certain nudibranchs, pycnogonids, turbellarians, micro-gastropods, and small polychaetes such as syllids, small arthropods and nematodes. These specialize in feeding on bryozoans by piercing and sucking out zooid contents. Cunha et al. (2018) observed feeding of the nudibranch Corambe carambola on the bryozoan Alcyonidium hauffi in Brazil. There were many empty zooids in the area of the attached nudibranch. It was observed on the bryozoan colony under a stereo microscope for about 3 hours, while its actions were documented with video. The nudibranch remained in one position, with its mantle lifted slightly. It moved with a short rocking motion that became faster and stronger from time to time. Movements of the radula and pumping apparatus were visible through the translucent skin of the nudibranch, as was food passing through its digestive system. Fig. 6.25 D shows another example – the nudibranch Okenia polycerelloides on the bryozoan Amathia verticillata on which it feeds and reproduces (Sales et al. 2019).



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Fig. 6.25: Interactions with other organisms. (A) Alcyonidium polypylum encrusting gastropod shell with live gastropod. (B) Alcyonidium polypylum encrusting a gastropod shell occupied by hermit crab and producing thick cylindrical branches. (C) Tube-building amphipod in branch fork of Bugula neritina. (D) Okenia nudibranch on its prey, Amathia verticillata. (E) Pycnogonid attached to Bugula neritina colony. (F) Didemnid ascidian colony settled on a Rhynchozoon colony. (G) Tube of a coronate scyphozoan attached to an encrusting cheilostome colony. (H) Celleporina sp. settled on an Amathia brasiliensis colony.

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6.7.1 Fouling There are many reports of bryozoans settling and living either on colonies of larger bryozoan species or on other organisms (Fig. 6.25 F–H), but the advantages and disadvantages to both parties are not always clear. In the case of vine-like colonies, e.g. Hippothoa, Aetea, and Aeverrillia, there seem to be nutritional advantages to settlement on larger sheet-like to erect species. McKinney (1988b) described a colony of Aetea curta that had grown along most branches of a large Cornucopina tuba colony with the distal portions of the Aetea zooids directed in such way that they could take advantage of the flow pattern created by the larger lophophores of the host. In this case, the Aetea would benefit, and the Cornucopina would not be negatively affected, but in cases in which recruits of encrusting species settled on non-specific colonies (e.g. Fig. 6.25 F, H), their growth would block zooids of the host species, apparently to the host’s detriment.

6.7.2 Commensalism Tamberg et al. (2013) reported that in the case of the solitary entoproct Loxosomella nordgaardi living on the surface of encrusting colonies of Tegella armifera, the entoprocts were found in areas of actively feeding polypides and strong colony currents. Since there was also an overlap in diet, especially in smaller particles (300 m). Light as a possible driving factor of community changes disappears in deep waters, and spatial competition seems less important at the low colony densities that, with few exceptions, are present there (Hughes 2001, Souto & Albuquerque 2019).

11.2.2 Diet of cheilostomes Like all bryozoans, cheilostomes are suspension feeders, but it remains unclear what their most important food sources are. Non-skeletized phytoplankton is generally considered as one of the main resources for bryozoans (Winston 1977; chapter 6). Diatom frustules have often been observed in the gut of actively feeding zooids, but in many cases appear to be released mostly undigested. Phytoplankton seems not to be the only food source, and species with large lophophores are able to capture ciliates, small eggs, and tiny zooplankton organisms. In some species, zooids catch prey by forming a cage with their tentacles that then move captured items toward the mouth with the tentacle tips (Winston 1978, Shunatova & Ostrovsky 2001; chapter 6). Bacteria are also taken, as demonstrated experimentally for both freshwater and marine Bryozoa (Richelle et al. 1994, Gosselin & Qian 2000). Tentacles are also able to directly absorb organic compounds dissolved in the water. Membranipora membranacea (Linnaeus, 1767), and possibly other species, can absorb dissolved organic substances originating from kelp mucus (De Burgh & Fankboner 1978). In deep waters, which lack phytoplankton, cheilostomes presumably feed on a combination of heterotrophic protists, non-living particulate detritus, and possibly dissolved organic matter (Hughes 2001). The presence of calcite in fecal pellets is indicative of ingestion, but not digestion, of detritus (McKinney & Jackson 1989).

11.2.3 Predation on Cheilostomata Predation of cheilostomes occurs at all stages of their life cycle, either by invertebrates or vertebrates. Liberated sperm can be ingested by suspension feeders such as sponges and ascidians and bryozoan larvae are part of the diet of hydrozoans, ctenophores, amphipods, cirripedes, and larvae/small fish. Bryozoan embryos in ovicells are highly nutritious and can be targeted by some predators (Lidgard 2008b). A range of animals has been observed to feed on bryozoan zooid and colonies.

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In marine environments, these include turbellarians, nematodes, polychaetes, crustaceans (decapods, amphipods, isopods, and copepods), mites, pycnogonids, mollusks (polyplacophorans, prosobranchs and especially nudibranchs), echinoderms, and some fish (e.g. Lidgard 2008a, b, Gordon et al. 2009). It is evident that bryozoans are part of the diet of some not very selective carnivorous and omnivorous taxa such as polyplacophorans, echinoderms, and fish (Lidgard 2008a, b, Gordon et  al. 2009, Wangensteen et  al. 2011). Several kinds of organisms found on bryozoan colonies, like mites and crustaceans, feed on surface microbiota and detritus and are either selectively eaten or ingested along with parts of the bryozoan (Hayward 1985, Hayward & Ryland 1985). Bryozoans and cnidarians are the main food of pycnogonids, although the exact relationship is difficult to assess, because their presence on bryozoan colonies does not necessarily imply that they are predators. Anascans lacking frontal-wall calcification are more vulnerable to their attacks. Adults of Achelia echinata feed on Flustra foliacea (Linnaeus, 1758) by extending their proboscis into the zooidal orifice when the operculum is open and sucking out internal contents (Wyer & King 1973). Other pycnogonids, like Austrodecus glaciale, are capable of introducing the long and thin proboscis through the frontal pores of Cellarinella foveolata Waters, 1904 avoiding damage to the proboscis that could be caused by the closing of the zooidal opercula or the mandibles of avicularia (Fry 1965). Nudibranchs are more-specialized predators of bryozoans with the structure of their feeding system being decisive in prey selection (Thompson & Brown 1976, Hayward & Ryland 1999). Some suctorial nudibranchs feed only on anascans with vulnerable frontal walls (Hayward 1985, Hayward & Ryland 1985). Others also feed on more-calcified bryozoans, drilling the area next to the opercula by radular abrasion and sucking out internal contents (Ryland & Hayward 1977). Some species even directly drill the frontal wall of zooids (Lidgard 2008b). Species of doridoidean nudibranchs are reported to feed on several cheilostomes, including Scruparia chelata (Linnaeus, 1758), Membranipora membranacea (Linnaeus, 1767), Electra pilosa (Linnaeus, 1767), Cradoscupocellaria reptans (Linnaeus, 1758), Bugula sp., Cryptosula pallasi­ ana (von Moll, 1803), Escharoides coccinea (Abildgaard, 1806), Schizomavella linearis (Hassall, 1841), Microporella ciliata (Pallas, 1766), Chorizopora brongniartii (Audouin, 1826), and Turbicellepora magnicostata (Barroso, 1919). Often, distinct specificity exists as seen in some species of Polycera, e.g. Polycera faeroensis and P. quadriline­ ata, which are usually associated with M. membranacea

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colonies. Also, some species of Arminoidea, like Jalonus hyalinus or J. cristatus, feed on the bryozoans S. chelata, S. reptans, and Bugula spp. (Thompson & Brown 1976). Generally, recently formed and young colonies are more vulnerable to predation than older, larger colonies (Lidgard 2008b). The effects of predation can be lethal or sublethal depending on the size of the predators and the mechanism of predation. Larger predators destroy many zooids, or even entire colonies, while small predators damage one or few zooids, which, however, are able to be repaired by regeneration (Berning 2008, Lidgard 2008a).

11.3 Systematics of Cheilostomata Cheilostome systematics are currently based mainly on the morphological key characters of the skeleton, while phylogenetically informative morphological characters of the soft parts are largely neglected. Traditionally, Cheilostomata has been subdivided into two morphologically distinct suborders, Anasca Levinsen, 1909 and Ascophora Levinsen, 1909 (e.g. Bassler 1953). Several classification concepts for Cheilostomata have been proposed more recently (e.g. d’Hondt 1985a, Viskova & Morozova 1988, Bock & Gordon 2013, d’Hondt 2016), but none of these concepts could gain universal acclaim, while Voigt (1991) showed that a systematic separation relying on the traditionally used groups to subdivide Cheilostomata is arbitrary as the characteristic morphological features associated with each group may be convergencies. Molecular data suggest that the currently employed morphological concepts to subdivide Cheilostomata may not withstand a molecular phylogeny (e.g. Knight et  al. 2011, Waeschenbach et  al. 2012). Therefore, combined molecular and morphological works are needed to better understand phylogenetic significance of morphological characters and homologies among structures. In the light of a wanting comprehensive and stable family-level molecular phylogeny of cheilostome bryozoans, we decided to not push another morphology-based classification but largely follow the classifications of Bock and Gordon (2013), Ostrovsky (2013), and Cook et  al. (2018) for convenience. However, Skylonina Viskova in Viskova and Morozova (1988), a valid but neglected name for an enigmatic suborder comprising three cheilostome families with mainly (or exclusively) fossil representatives, is reintroduced here. Thus, we recognize eight nominal suborders of Cheilostomata. The systematic account presented here lists only families and genera that include nominally Recent species, while cheilostome families and genera based on extinct

species were not considered. Taxa in the systematic listing (families, genera and type species) followed by a dagger (†) are based on a fossil nominal type specimen but the genus or family includes at least one extant representative. The appendices of this chapter provide a glossary of terms and a systematic list of taxa as presented in this chapter. A new replacement name for a secondary homonym is Bryobuchneria nom. nov. (replacing Buchneria Harmer, 1957, a secondary homonym of the diplopod genus Buch­ neria Verhoeff, 1941).

Phylum Bryozoa Ehrenberg, 1831 Order Cheilostomata Busk, 1852a 11.3.1 Suborder Membraniporina Ortmann, 1890 Remarks: Malacostegina Levinsen, 1902 has been synonymized with Membraniporina after the concept of Malacostegina was narrowed (Kubanin 2001). 11.3.1.1 Superfamily Membraniporoidea Busk, 1852b Family Membraniporidae Busk, 1852b Type genus: Membranipora de Blainville, 1830. Type species: Flustra membranacea Linnaeus, 1767. Other genera: Acanthodesia Canu & Bassler, 1919, Biflus­ tra d’Orbigny, 1852, Jellyella Taylor & Monks, 1997. Diagnosis: Colonies encrusting, unilamellar, multiserial; or erect, bilamellar. Autozooids lacking frontal-wall calcification, longitudinally rectangular. Gymnocyst absent or reduced, smooth. Cryptocyst narrow or negligible, steeply sloping into the opesia. Spinules (or spine-like denticles) or adoral tubercles may be present, occurring under the membranous frontal wall on lateral walls. Intertentacular organ present. Embryos non-brooded, planktotrophic cyphonautes. Ancestrula twinned. Kenozooids and avicularia-like zooids may occur. Brood chambers absent (Fig. 11.5 A, B). Distribution: Cosmopolitan. Remarks: Busk (1854) is often erroneously indicated as the author of Membraniporidae. However, the family was first cited in Busk (1852b, pp. 2, 47). Biflustridae d’Orbigny, 1852 was proposed the same year (d’Orbigny 1852, p. 217), and although it may be a senior synonym, the name has not been used with d’Orbigny as an author by subsequent authors. Membraniporidae Busk, 1852b is therefore considered valid in accordance with Article 23.9 of the International Code on Zoological Nomenclature



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Fig. 11.5: (A, B) Membraniporidae: Membranipora villosa Hincks, 1880a. Ketchikan, Alaska, USA, NE Pacific Ocean, intertidal. Peabody Museum of Natural History, New Haven, YPM IZ 100285. (C, D) Electridae: Electra asiatica Grischenko, Dick & Mawatari, 2007. Akkeshi Bay, Hokkaido, Japan, NW Pacific Ocean, intertidal. Natural History Museum, London, 2006.2.27.20 and 2006.2.27.21. Scale bars: A, B, 500 µm; C, 1 mm; D, 250 µm.

(ICZN 1999). Smitt (1873) also proposed the family Biflustridae, unaware of Busk’s and d’Orbigny’s earlier proposals. Biflustridae Smitt, 1873 is regarded as a junior synonym of Membraniporidae. The genera Biflustra and Membranipora have long been used by many authors for both fossil and Recent “anascan” species with little or no frontal-wall calcification. Some still require proper revision and especially the fossil material may not represent true membraniporids. In contrast to Electridae Stach, 1937 (1851), membraniporids usually have no gymnocyst at all or a moderately developed proximal gymnocyst in Jellyella. Family Electridae Stach, 1937 (1851) Type genus: Electra Lamouroux, 1816. Type species: Flustra verticillata Ellis & Solander, 1786.

Other genera: Arbocuspis Nikulina, 2010, Arbopercula Nikulina, 2010, Aspidelectra Levinsen, 1909, Bathy­ pora MacGillivray, 1885a, Conopeum Gray, 1848, Einhor­ nia Nikulina, 2007, Harpecia Gordon, 1982, Lapidosella Gontar, 2010, Miravitrea Gontar, 2014, Mychoplectra Gordon & Parker, 1991a, Osburnea Nikulina, 2010, Pyri­ pora d’Orbigny, 1852, Tarsocryptus Tilbrook, 2011, Villich­ arixa Gordon, 1989a. Diagnosis: Colonies encrusting, unilamellar, uniserial, oligoserial or multiserial; or erect, unilamellar to multi­ lamellar. Autozooids usually rectangular, elliptical or pyriform. Gymnocyst completely surrounding opesia, smooth or perforate, occasionally caudate. Cryptocyst steeply sloping into the opesia, reduced. Periopesial spines absent or present. Intertentacular organ present. Embryos non-brooded, planktotrophic cyphonautes. Ancestrula

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autozooidal, single. Kenozooids may be present. Avicularia, pore-chambers, and brood chambers absent (Fig. 11.5 C, D). Distribution: Cosmopolitan. Remarks: Stach (1937) proposed a change in the family name from Electrinidae d’Orbigny, 1851 to the new name Electridae as Electrina d’Orbigny, 1851 has previously been recognized as a junior synonym of Electra Lamouroux, 1816 by Harmer (1926). While some authors continued to use Electrinidae as a family name (e.g. Osburn 1950), some later authors mistakenly used the wrong combination ‘Electridae d’Orbigny, 1851’ (e.g. Martha et al. 2019c). As this family-group name was replaced before 1961, in accordance with Article 40.2 of the International Code of Zoological Nomenclature (ICZN 1999), the substitute name Electridae is maintained in prevailing usage and should be cited with its original author and date followed by the date of its priority enclosed in parentheses, thus Electridae Stach, 1937 (1851). Electrids are similar to membraniporids, but the gymncyst is usually circumopesial and spines are usually present, while avicularia-like polymorphs are always lacking. The most important feature used to distinguish electrids from membraniporids is currently the ancestrula that is twinned in membraniporids but single in electrids.

zooids of Sinoflustra annae (Osburn, 1953), which also indicates planktotrophic larvae.

Family Sinoflustridae Gordon, 2009a Type genus: Sinoflustra Liu & Yang, 1995. Type species: Membranipora amoyensis Robertson, 1921. Other genera: Membraniporopsis Liu in Liu et al. (1999). Diagnosis: Colonies encrusting, unilamellar, multiserial; or erect, unilamellar, multiserial or bilamellar, multiserial. Autozooids lacking frontal wall calcification, longitudinally rectangular. Gymnocyst absent or negligible. Cryptocyst narrow, usually granular, occasionally with spinous processes. Avicularia-like zooids larger than autozooids, bearing large, mandible-like operculum. Kenozooids paired, distolaterally. Ancestrula single, bearing distolateral kenozooids. Brood chambers absent. Inter­ tentacular organ may be present (Fig. 11.6 A, B). Distribution: Tropical waters. Remarks: Gordon (2009a) introduced this family for two membraniporiform genera with disolateral spine- or funnel-like kenozooids. The type genus, Sinoflustra, also has vicarious avicularium-like heterozooids. Sinoflustrids further have a single, autozooidal ancestrula, while “true” membraniporids have a twinned ancestrula. Vieira and Migotto (2015, fig. 1c) recently showed an intertentacular organ in Membraniporopsis tubigera (Osburn, 1940), indicating that this family consists of non-brooding taxa with planktotrophic cyphonautes larvae. Karande and Udhayakumar (1992) noted six to seven small eggs in reproductive

11.3.2.1 Superfamily Aeteoidea Smitt, 1868a

11.3.2 Suborder Inovicellina Jullien, 1888 Remarks: Gordon and Bock proposed in Cook et al. (2018) the name Aeteina as a replacement for Inovicellina in order to typify suborder names with family names. Viskova and Morozova (1988) already used “Aeteida Smitt, 1867” (p. 18) as a name for an order that they based on Aeteidae Smitt, 1868a. However, such a name above family-level was not proposed in Smitt (1868a), and thus, Smitt cannot be considered author of the Aeteina. Viskova and Morozova (1988) considered Inovicellina as a junior synonym of their “Aeteida” and authorship of Aeteina may be attributed to them. Nonetheless, there is no provision for coordination of order-names in the International Code of Zoological Nomenclature (ICZN 1999), and many names of animal orders are not coordinated with family-group names. This holds also true for most orders of cyclostome Bryozoa, and Inovicellina is a well-established and stable name. Both Aeteina Viskova & Morozova, 1988 and Aeteina Gordon & Bock in Cook et al., 2018 are therefore suppressed.

Family Aeteidae Smitt, 1868a Type genus: Aetea Lamouroux, 1812. Type species: Sertularia anguina Linnaeus, 1758. Other genera: Callaetea Winston, 2008. Diagnosis: Colonies encrusting, rarely erect, unilamellar, uniserial. Autozooids weakly calcified, tubular, with decumbent proximal portion and free, erect distal portion. Adherent parts contribute to the formation of a stolon. Frontal membrane differentiated terminally as an operculum. Single embryos (sometimes two) incubated in external membranous brood sac attached to the zooidal wall. Larva non-feeding. Pore-chambers or septula present. Ancestrula single, a smaller version of normal autozooids and with reduced number of tentacles. Spines, avicularia, and ovicells absent. Kenozooids (zoeciules or sacculi) rarely present (Fig. 11.6 C, D). Distribution: Cosmopolitan. Remarks: Aeteids are very simple and weakly calcified cheilostomes that have bipartite autozooids consisting of a decumbent proximal portion and a free, erect distal portion. Despite the presence of zoeciules in Aetea cul­ trata Vieira, Almeida & Winston, 2016 and sacculi in Aetea sica (Couch, 1844) (see Simma-Krieg 1969), no other polymorphs are present and colonies are always uniserial.



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Fig. 11.6: (A, B) Sinoflustridae: Membraniporopsis tubigera (Osburn, 1940). São Sebastião, São Paulo, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, unregistered. (C, D) Aeteidae: Aetea cultrata Vieira, Almeida & Winston, 2016. Maceió, Alagoas, Brazil, W Atlantic Ocean. Universidade Federal de Pernambuco, 076, holotype. Scale bars: A, 500 µm; B, C, 100 µm; D, 50 µm.

Aetea truncata (Landsborough, 1852) can form erect colonial branches from the distal portions of a zooid. All mentionings of fossil Aeteidae except for possibly Canu (1912) and bioimmured material described by Voigt (1983) are dubious.

11.3.3 Suborder Scrupariina Silén, 1941 Remarks: There is some ambiguity regarding the correct date of publication of Silén (1941). While it is common to take 1941 as the date of publication, some authors rather indicated 1942 (e.g. Bassler 1953). The correctness of 1941 is suggested by the front cover of the offprint, as well as by a note on p. 130 that reads “Tryckt den 23 september 1941. Uppsala 1941” (“Printed on 23 September 1941. Uppsala 1941”). Silén (1942) supports this by citing the publication as 1941, which is used here for all associated taxa.

11.3.3.1 Superfamily Scruparioidea Gray, 1848 Family Scrupariidae Gray, 1848 Type genus: Scruparia Oken, 1815. Type species: Sertularia chelata Linnaeus, 1758. Other genera: Brettiopsis López Gappa, 1986. Diagnosis: Colonies erect, rising from creeping base, dichotomously branching. Autozooids (and kenozooids) arranged in a uniserial series, weakly calcified, tubular, with subterminal frontal membrane. Opesia less than half the autozooidal length, elliptical. Budding of zooids from distal margin of an autozooid or proximal to the opesia. Polypide with about 12 tentacles. Tentacle sheath encircled by a ring of setose teeth. Several embryos incubated simultaneously in a bivalved terminal ovicell-like brood chamber, semicleithral or acleithral. Larva lecithotrophic. Ancestrula single, a smaller version of normal autozooids. Spines and avicularia absent (Fig. 11.7 A, B).

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Fig. 11.7: (A, B) Scrupariidae: Scruparia ambigua (d’Orbigny, 1841). Caniçal, Maderia Island, E Atlantic. University of Vienna. (C, D) Eucrateidae: Eucratea loricata (Linnaeus, 1758). Barents Sea. Perm State National Research University, Kluge Collection, 810.03. Scale bars: A, C, 500 µm; B, 100 µm; D, 250 µm.

Distribution: Cosmopolitan. Remarks: Gray (1848) consistently misspelled the generic name Scruparia as “Scuparia” and accordingly incorrectly proposed the family “Scupariadæ.” The first to correctly spell the family name was Busk (1852b), who is sometimes indicated as the author of both the family and the superfamily. Still, according to Article 35.4.1 of the Inter­ national Code of Zoological Nomenclature (ICZN 1999), a family-group name based upon an incorrect spelling of the name of the type genus must be corrected. Therefore, Gray (1848) is deemed as the author of both Scrupariidae and Scruparioidea. Family Eucrateidae Johnston, 1847 Type genus: Eucratea Lamouroux, 1812. Type species: Sertularia loricata Linnaeus, 1758. Other genera: None.

Diagnosis: Colonies erect, attached to substratum by kenozooidal rhizoids, dichotomously branching. Autozooids mostly arranged in back-to-back pairs, effectively forming biserial branches, weakly calcified, long and slender. Opesia less than half the autozooidal length, elliptical. Cryptocyst negligible. Budding of zooids distobasal. Embryos incubated singly in external membranous brood sacs attached above zooidal orifice. Ancestrula attached by distally budded initial rhizoid to substratum. Spines, avicularia, and ovicells absent (Fig. 11.7 C, D). Distribution: Polar waters of the Arctic Ocean. Remarks: Eucrateids mainly differ from scrupariids in brooding their embryos externally in membranous sacs, while the latter have bivalved ovicell-like brood chambers. Furthermore, the four known species all show back-toback orientation of the autozooids.



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Fig. 11.8: (A, B) Leiosalpingidae: Leiosalpinx australis (Busk, 1884). Station E800, Dagg Sound, South Island, New Zealand, SW Pacific Ocean, National Institute of Water & Atmospheric Research, unregistered. Modified after Gordon (1986). (C, D) Tendridae: Tendra zostericola de Nordmann, 1839. Karantinnaya Bay, Sevastopol, Crimean Peninsula, Black Sea. Saint Petersburg State University, Russian Federation, unregistered.

Family Leiosalpingidae d’Hondt & Gordon, 1996 Type genus: Leiosalpinx Hayward & Cook, 1979. Type species: Alysidium inornata Goldstein, 1882. Other genera: Astoleiosalpinx d’Hondt & Gordon, 1996. Diagnosis: Colonies erect, dichotomously branching, attached to substratum by kenozooidal rhizoids. Autozooids arranged in a uniserial series, weakly calcified, long and slender. Opesia less than half the autozooidal length, elliptical. Budding of zooids distobasal. One or two embryos incubated externally in membranous brood sac attached to zooidal frontal membrane near zooidal orifice. Ancestrula erect, attached to substratum by initially budded rhizoid. Spines, avicularia, and ovicells absent (Fig. 11.8 A, B). Distribution: Tropical Indo-Pacific.

Remarks: Leiosalpingidae comprises three species that are all from the Indo-Pacific of the Southern Hemisphere. Leiosalpingids show no back-to-back orientation of the zooids and have a different budding pattern than eucrateids (d’Hondt & Gordon 1996), but the two families compare well in most other characters, especially in the reproductive pattern (Ostrovsky 2013).

11.3.4 Suborder Tendrina Ostrovsky, 2013 Remarks: Ostrovsky (2013) proposed the suborder Tendrina for the sole-included family Tendridae based on the acanthostegal brood chambers and the non-feeding larva, which probably evolved from a membraniporine ancestor independently of the flustrine cheilostomes that

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have ovicells and a non-feeding larva (Ostrovsky 2013). The presence of an intertentacular organ, known only from some membraniporines and thalamoporellines, suggests a possible evolution of Tendrina from a cheilostome ancestor with an intertentacular organ and non-articulated mural spines (Ostrovsky 2013). 11.3.4.1 Superfamily Tendroidea Vigneaux, 1949 Family Tendridae Vigneaux, 1949 Type genus: Tendra de Nordmann, 1839. Type species: Tendra zostericola de Nordmann, 1839. Other genera: Heterooecium Hincks, 1892. Diagnosis: Colonies encrusting and unilamellar, uniserial to oligoserial or multiserial. Autozooids lacking frontal-wall calcification, longitudinally rectangular. Gymnocyst well developed, smooth. Cryptocyst narrow or negligible, steeply sloping into the opesia. Intertentacular organ may be present. Spines oral and circumopesial, articulated or non-articulated, overarching the opesia. Several embryos incubated simultaneously in acanthostegal brood chambers formed by periopesial non-articulated spines formed by autozooid or kenozooid distal to fertile autozooid. Larva lecithotrophic. Septula in vertical walls, multiporous. Ancestrula autozooidal. Kenozooids and avicularia absent (Fig. 11.8 C, D). Distribution: Temperate waters of the Black Sea and tropical waters of the Indo-Pacific. Remarks: The two genera of Tendridae each comprise two species with those of Tendra occurring in the Black Sea (Gryncharova 1980) and the Mediterranean Sea (Occhipinti Ambrogi & d’Hondt 1981) and the species of Heterooecium being reported from waters around western and southern Australia (Hincks 1881d, Levinsen 1909, Hastings 1966).

11.3.5 Suborder Thalamoporellina Ostrovsky, 2013 Remarks: Ostrovsky (2013) erected this suborder because of the bivalved ovicell-like brood chambers found in some thalamoporellids. Steginoporellids and some thalamoporellids, however, incubate their larvae in internal brood sacs (which may have been accomplished through an evolutionary substitution of ovicells by internal brood sacs). 11.3.5.1 Superfamily Thalamoporelloidea Levinsen, 1902 Family Thalamoporellidae Levinsen, 1902 Type genus: Thalamoporella Hincks, 1887.

Type species: Flustra rozieri Audouin, 1826. Other genera: Dibunostoma Cheetham, 1963, Diploporella MacGillivray, 1885b, Hesychoxenia Gordon & Parker, 1991b, Marsupioporella Soule, Soule & Chaney, 1991a, Thairopora MacGillivray, 1882. Diagnosis: Colonies encrusting, unilamellar to multilamellar, multiserial; or erect, stalk-like, multiserial, articulated or non-articulated, dichotomously branching. Autozooids longitudinally rectangular to hexagonal. Gymnocyst negligible to absent, smooth. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, granular, pseudoporous, depressed. Opesiules may occur, often asymmetrical, circular, as a single pair or in series. Intertentacular organ may be present. Adoral tubercles may be present. Internal spicules may be present, projecting into body cavity. Up to seven embryos incubated simultaneously in bivalved ovicell-like brood chambers or in internal brood sacs. Ovicell-like brood chambers, if present, hyperstomial, cleithral, bivalved, ooecium with median suture. Septula in vertical walls, multiporous. Intertentacular organ may be present. Larva lecithotrophic. Ancestrula autozooidal or kenozooidal. Avicularia vicarious, irregularly interspersed among autozooids. Kenozooids may be present (Fig. 11.9 A, B). Distribution: Tropical to temperate waters. Remarks: Thalamoporellids usually have bivalved ovicell-like brood chambers superficially similar to those in scrupariids and alysidiids but apparently evolved independently (Ostrovsky 2013). The cryptocyst in thalamoporellids is pseudoporous and often pierced by usually asymmetrical opesiules. Furthermore, Thalamoporellidae is the only cheilostome family known to possess minute, caliper-shaped spicules projecting into the body cavity. Large vicarious avicularia are present in most species but are lacking in the type species, T.  rozieri, a neotype for which was selected by Soule et al. (1992). Family Steginoporellidae Hincks, 1884a Type genus: Steginoporella Smitt, 1873. Type species: Membranipora magnilabris Busk, 1854. Other genera: Labioporella Harmer, 1926, Siphonoporella Hincks, 1880b. Diagnosis: Colonies encrusting, unilamellar to multilamellar, multiserial; or erect from an encrusting base, cylindrical to frondose, multiserial, dichotomously branching and anastomosing. Autozooids longitudinally rectangular to hexagonal. Gymnocyst negligible to absent, smooth. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, pseudoporous, depressed, forming a median process with an associated polypide tube. Embryos incubated singly in internal brood sacs. Septula in vertical walls, multiporous. Ancestrula autozooidal. B-zooids vicarious, irregularly interspersed



11.3 Systematics of Cheilostomata 

 333

Fig. 11.9: (A, B) Thalamoporellidae: Thalamoporella floridana (Osburn, 1940). Maceió, Alagoas, Brazil, W Atlantic Ocean. Universidade Federal de Pernambuco, 079, Vieira Collection. (C, D) Steginoporellidae: Steginoporella magnilabris (Busk, 1854). Maceió, Alagoas, Brazil, W Atlantic Ocean. Universidade Federal de Pernambuco, 082. Scale bars: A, C, 500 µm; D, 250 µm; B, 100 µm.

among autozooids. Avicularia usually absent, sometimes vicarious or interzooidal, irregularly interspersed among autozooids. Kenozooids may be present (Fig. 11.9 C, D). Distribution: Cosmopolitan. Remarks: B-zooids are characteristic for most stegino­ porellid species. They are larger than ordinary autozooids (“A-zooids”). In contrast to avicularia, B-zooids possess a lophophore and are thus capable of feeding. They further possess augmented opercula and opercular muscles, which are lacking in A-zooids (see Banta 1973). Some steginoporellid species may also have interzooidal avicularia (Banta 1973).

11.3.6 Suborder Belluloporina Ostrovsky, 2013 Remarks: Like Tendrina and Thalamoporellina, Ostrovsky (2013) erected this suborder for its unique brooding

features that have evolved independently and are incomparable to those of any other cheilostome taxa.

11.3.6.1 Superfamily Belluloporoidea Ostrovsky, 2013 Family Belluloporidae Ostrovsky, 2013 Type genus: Bellulopora Lagaaij, 1963. Type species: Colletosia bellula Osburn, 1950. Other genera: None. Diagnosis: Colonies encrusting, unilamellar, multiserial. Autozooids arranged in irregular series, longitudinally hexagonal to elliptical. Gymnocyst narrow, smooth. Frontal shield made of kenozooidal costae overarching the frontal membrane. Orifice terminal. Spines oral. Pore-chambers basal. Embryos incubated in ovicell-like cleithral brood chambers composed of fused costae-like

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Fig. 11.10: (A, B) Belluloporidae: Bellulopora bellula (Osburn, 1950). Florida, USA, W Atlantic Ocean. Natural History Museum, London, 1986.8.14.20. (C, D) Flustridae: Isosecuriflustra pinniformis Vieira, Gordon, Souza & Haddad, 2010. São Paulo, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, 0277, holotype. Scale bars: A, B, 500 µm; C, 2.5 mm; D, 250 µm.

kenozooids. Ancestrula tatiform, bearing mural spines. Avicularia absent (Fig. 11.10 A, B). Distribution: Eastern Pacific and Western Atlantic. Remarks: Belluloporidae is a monospecific family, the type species of which differs from other cheilostomes in having unique brood chambers composed of fused costa-like kenozooids. Ostrovsky (2013) used this character to justify a new family and suborder for Bellulopora bellula, while previous authors had included the species in the family Cribrilinidae Hincks, 1879 owing to the costate frontal shield (e.g. Winston 1982).

Ascophora Levinsen, 1909 has just recently come out of use and all taxa (except for Bellulopora bellula (Osburn, 1950)) previously included in Ascophora are now included in Flustrina. Neocheilostomina d’Hondt, 1985a, which some authors still use, is a junior synonym of Flustrina in the current concept of cheilostome systematics (see Cook et al. 2018). For convenience, we group the original flustrine superfamilies (Flustroidea, Buguloidea, Calloporoidea, Cellarioidea, Lunulitoidea, Microporoidea, and Monoporelloidea) at the beginning of the account, followed by the “ascophoran”-grade superfamilies in alphabetical order.

11.3.7 Suborder Flustrina Smitt, 1867

11.3.7.1 Superfamily Flustroidea Lamoroux, 1816

Remarks: Flustrina is a highly heterogeneous group that now includes most cheilostome taxa with a high variability of morphological characters. The unaccepted suborder

Family Flustridae Lamouroux, 1816 Type genus: Flustra Linnaeus, 1761. Type species: Eschara foliacea Linnaeus, 1758.



Other genera: Austroflustra López Gappa, 1982, Carbasea Gray, 1848, Chartella Gray, 1848, Gontarella Grischenko, Taylor & Mawatari, 2002, Gregarinidra Barroso, 1948, Hincksina Norman, 1903a, Hincksinoflustra Bobin & Prenant, 1961, Isosecuriflustra Liu & Hu, 1991, Kenella Levinsen, 1909, Klugeflustra Moyano G., 1972, Nematoflustra Moyano G., 1972, Neoflustra López Gappa, 1982, Retiflus­ tra Levinsen, 1909, Sarsiflustra Jullien in Jullien & Calvet, 1903, Securiflustra Silén, 1941, Serratiflustra Moyano G., 1972, Spiralaria Busk, 1861a, Terminoflustra Silén, 1941. Diagnosis: Colonies erect, unilamellar, multiserial, nonarticulated, dichotomously branching; or erect, frondose, bilamellar, multiserial; sometimes encrusting, unilamellar, multiserial. Erect colonies attached to substratum by a narrow encrusting base. Autozooids rectangular, weakly calcified. Gymnocyst absent. Cryptocyst absent or weakly developed proximo-laterally as a narrow rim, smooth. Opesia occupying almost the whole frontal area. Spines usually absent, but may occur at the distal corners or around the lateral rim. Embryos incubated singly in ovicells or internal brood sacs. Ovicells usually endozooidal or, sometimes, hyperstomial, acleithral, semicleithral, or cleithral. Larva non-feeding. Septula in vertical walls, uni- or multiporous. Ancestrula autozooidal. Avicularia may be present, usually vicarious, irregularly interspersed among autozooids, often at row bifurcations. Kenozooids may be present (Fig. 11.10 C, D). Distribution: Cosmopolitan. Remarks: Fleming (1828) or Smitt (1867), respectively, are often recognized as authors of Flustridae. Gordon (1984) appears to be the first who recognized the authorship of Lamouroux (1821) based on Lamouroux’ “Flustrées/Flustreæ” (1821, p. 2). “Flustrées” was used as a taxonomic name to group genera (i.e. at a family level) and therefore fulfills the provisions of Article 11.7 of the International Code on Zoological Nomenclature (ICZN 1999). Since Lamouroux used the name “Flustrées/Flustreæ” already in an earlier publication (Lamouroux 1816, p. 84), the correct date of this family is 1816 rather than 1821 according to Article 11.7.2 of the International Code on Zoological Nomencla­ ture. Although Lamouroux called “Flustrées/Flustreæ” an “ordre” in both Lamouroux (1816, 1821), it is evident that he denoted a suprageneric taxon. Lamouroux’ (1816, 1821) use of the word “ordre” to denote a family is further corroborated when comparing his later works with Lamouroux (1812). In this publication, he subdivided the zoophytes into six “familles” (French for families) with Flustra and other cheilostome genera having been included in the family “Sertulariées/Sertularieæ.” Both the French (-ées) and the Latin (-eæ) endings used in Lamouroux (1812) for his “familles” correspond with the endings used in Lamouroux (1816, 1821) for his “ordres.” The six “familles” included in Lamouroux (1812) do also appear in Lamouroux (1816, 1821), now

11.3 Systematics of Cheilostomata 

 335

as “ordres.” So, “Flustrées/Flustreæ” and the other “ordre” names first mentioned in Lamouroux (1816) must be considered available family names according to Article 11.7.1.2 of the International Code on Zoological Nomenclature. Flustridae is a highly heterogeneous family of simple and weakly calcified anascan cheilostomes (e.g. Silén 1941). Its composition will change pending proper revision. Especially the placement of the genera Austroflustra, Klugeflustra, and Neoflustra in Flustridae is very uncertain. Liu and Hu (1991) introduced the new genus Isoseculiflus­ tra, which must be considered a lapsus calami and has to be corrected to Isosecuriflustra based on Article 32.4.1 of the International Code on Zoological Nomenclature (ICZN 1999).

11.3.7.2 Superfamily Buguloidea Gray, 1848 Family Bugulidae Gray, 1848 Type genus: Bugula Oken, 1815. Type species: Sertularia neritina Linnaeus, 1758. Other genera: Beanodendria d’Hondt & Gordon, 1996, Bicellariella Levinsen, 1909, Bicellarina Levinsen, 1909, Brettiella Gordon, 1984, Bugularia Levinsen, 1909, Bugule­ lla Verrill, 1879a, Bugulina Gray, 1848, Calyptozoum Harmer, 1926, Camptoplites Harmer, 1923, Caulibugula Verrill, 1900, Cornucopina Levinsen, 1909, Corynoporella Hincks, 1888, Crisularia Gray, 1848, Cuneiforma d’Hondt & Schopf, 1985, Dendrobeania Levinsen, 1909, Dimetopia Busk, 1852b, Fal­ sibugulella Liu, 1984, Farciminellopsis Silén, 1941, Haloph­ ila Gray, 1843, Himantozoum Harmer, 1923, Himantozoume­ lla d’Hondt & Schopf, 1985, Kinetoskias Danielssen, 1868, Klugella Hastings, 1943, Luguba Gordon, 1984, Nordgaardia Kluge, 1962, Semidendrobeania d’Hondt & Schopf, 1985, Semikinetoskias Silén, 1941, Sessibugula Osburn, 1950, Thaminozoum d’Hondt & Gordon, 1996, Uschakovia Kluge, 1946, Virididentula Fehlauer-Ale, Winston, Tilbrook, Nascimento & Vieira, 2015, Xenoflustra Moyano G., 2011. Diagnosis: Colonies erect, unilamellar, bi- to multiserial, non-articulated, dichotomously branching, attached to substratum by rhizoids; sometimes encrusting, unilamellar, unito multiserial, repent, supported above the substratum by rhizoids. Autozooids rectangular, weakly calcified. Frontal gymnocyst absent or moderately to little developed proximally, smooth. Cryptocyst very narrow, vestigial or absent. Opesia occupying much or almost the whole frontal area. Spines usually present, oral and circumopesial, articulated or non-articulated. Embryos incubated in hyperstomial or, in few instances, immersed ovicells; sometimes terminal, acleithral, semicleithral, or cleithral. Larva non-feeding. Ancestrula autozooidal. Avicularia may be present, usually of bird’s-head form, associated with many autozooids. Kenozooids may be present (Fig. 11.11 A, B).

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 11 Gymnolaemata, Cheilostomata

Fig. 11.11: (A, B) Bugulidae: Bugula neritina (Linnaeus, 1758). Australia. Museu de Zoologia, Universidade de São Paulo, unregistered. (C, D) Beaniidae: Beania correiae Vieira, Migotto & Winston, 2010. Maceió, Alagoas, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, 0402, holotype. Scale bars: A, C, 500 µm; B, 250 µm; D, 100 µm.

Distribution: Cosmopolitan. Remarks: Characteristic for this family are the so-called bird’s-head avicularia – pedunculate avicularia shaped like a bird’s-head. However, these avicularia are lacking in many species, including also the type species of the family. Family Beaniidae Canu & Bassler, 1927 Type genus: Beania Johnston, 1840. Type species: Beania mirabilis Johnston, 1840. Other genera: Stolonella Hincks, 1883. Diagnosis: Colonies encrusting, unilamellar, uni- to multiserial, repent, supported above the substratum by rhi­zoids; or erect, bilamellar, frondose, multiserial, attached to substratum by rhizoids. Autozooids rectangular, weakly calcified, disjunct but interconnected by calcified tubes. Gymnocyst and cryptocyst absent. Opesia occupying almost the whole frontal area. Spines usually present, oral or circumopesial and sometimes basal, articulated or non-articulated.

Embryos incubated in immersed terminal ovicell or in internal brood sacs with vestigial ooecium. Septula in vertical walls, multiporous. Avicularia may be present, usually pedunculate, often in an oral position (Fig. 11.11 C, D). Distribution: Cosmopolitan (except for polar waters of the Northern Hemisphere). Remarks: Characteristic of beaniids are the disjunct zooids that are interconnected by calcified tubes originating from pore-plates inside the vertical walls as well as pedunculate avicularia, the latter, however, lacking, in many species. The genus Amphibiobeania Metcalfe, Gordon, & Hayward, 2007, originally included in Beaniidae, may be a ctenostome (Cook et al. 2018). This family is highly diverse and awaits proper revision. Despite morphological differences, at least three named genera, i.e. Diachoris Busk, 1852a, Chaunosia Busk, 1867, and Dimor­ phozoum Levinsen, 1909, are regarded as junior synonyms of Beania (Hastings 1939, Vieira et al. 2010b).



11.3 Systematics of Cheilostomata 

 337

Fig. 11.12: (A, B) Candidae: Scrupocellaria scrupea Busk, 1852b. Guernsey, United Kingdom, English Channel. Natural History Museum, London, 1911.10.1.360 01. (C, D) Epistomiidae: Synnotum aegyptiacum (Audouin, 1826). Maceió, Alagoas, Brazil, W Atlantic Ocean. Universidade Federal de Pernambuco, unregistered. Scale bars: A, B, 250 µm; C, 500 µm; D, 100 µm.

Family Candidae d’Orbigny, 1851 Type genus: Canda Lamouroux, 1816. Type species: Canda arachnoides Lamouroux, 1816 (= Cellaria filifera Lamarck, 1816). Other genera: Amastigia Busk, 1852b, Aquiloniella Vieira, Spencer Jones, Winston, Migotto & Marques, 2014, Aspis­ cellaria Vieira, Spencer Jones, Winston, Migotto & Marques, 2014, Bathycellaria Vieira, Spencer Jones, Winston, Migotto & Marques, 2014, Bobinella d’Hondt, 1981, Bugulopsis Verrill, 1879b, Caberea Lamouroux, 1816, Cabereopsis Hasenbank, 1932, Candomenipea d’Hondt & Gordon, 1996, Cando­ scrupocellaria d’Hondt & Gordon, 1996, Cradoscrupocellaria Vieira, Spencer Jones, & Winston, 2013, Diplobicellariella d’Hondt, 1985b, Emma Gray, 1843, Eupaxia Hasenbank, 1932, Hoplitella Levinsen, 1909, Licornia van Beneden, 1850, Maplestonia MacGillivray, 1885b, Menipea Lamouroux, 1812, Monartron Canu & Bassler, 1929a, Notoplites Harmer,

1923, Notoplitesigia d’Hondt, 1987, Paralicornia Vieira, Spencer Jones, Winston, Migotto & Marques, 2014, Penemia Gordon, 1986, Pomocellaria Vieira, Spencer Jones, Winston, Migotto & Marques, 2014, Pseudoporicellaria d’Hondt, 1987, Scrupocaberea Vieira, Spencer Jones, Winston, Migotto & Marques, 2014, Scrupocellaria van Beneden, 1845, Scrupo­ cellarinella d’Hondt & Schopf, 1985, Semibugula Kluge, 1929, Sinocellaria Vieira, Spencer Jones, Winston, Migotto & Marques, 2014, Tricellaria Fleming, 1828. Diagnosis: Colonies erect, bi-, oligo- or multiserial, unilamellar, weakly jointed or unjointed, bifurcating, supported above the substratum by rhizoids. Autozooids rectangular to elliptical. Gymnocyst circumopesial, smooth. Cryptocyst weakly developed, sometimes granular and crenulated. Opesia occupying most of the frontal area. Spines may occur, usually in an oral position; some species with simple or enlarged, flattened, branching, spine (scutum) on one

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side of and protecting membranous frontal wall. Septula in vertical walls, multiporous. Embryos incubated in hyperstomial, endozooidal, or immersed ovicells, acleithral, semicleithral, or cleithral. Ectooecium pseudoporous or with frontal membranous area. Larva non-feeding. Ancestrula autozooidal, semierect. Avicularia may be present, adventitious, associated with many autozooids, frontally or laterally. Avicularium-like vibracular polymorphs with long setae may be present axially, distolaterally, or basally. Kenozooids may be present (Fig. 11.12 A, B). Distribution: Cosmopolitan. Remarks: Among the characteristics of this family are erect colonies with all autozooids facing in one direction. Branches may be unjointed (e.g. Caberea) or relatively weakly jointed across the middle of zooids distal to bifurcations. In some taxa (e.g. Canda), stolon-like kenozooids connect adjacent branches. Many species possess a scutum, i.e. a modified, mostly enlarged, flattened, and sometimes branching lateral opesial spine that overarches the frontal area. The type species of the type genus Canda Lamouroux, 1816, Canda arachnoides Lamouroux, 1816, is a junior synonym of Cellaria filifera Lamarck, 1816 according to d’Hondt (1988). Bugulicellaria Mawatari, 1957 is a junior subjective synonym of Semibugula Kluge, 1929. Family Epistomiidae Gregory, 1893 Type genus: Epistomia Fleming, 1828. Type species: Sertularia bursaria Linnaeus, 1758. Other genera: Synnotum Pieper, 1881. Diagnosis: Colonies erect, biserial with zooids arranged in obliquely shifted pairs, articulated, bifurcating, attached to substratum by rhizoids. Autozooids weakly calcified, tubular, disjunct but interconnected by cuticular joints. Gymnocyst and cryptocyst absent. Opesia occupying almost the whole frontal area. Viviparous. Single embryo incubated intracoelomically in swollen or enlarged polymorphic female zooids. Ancestrula not reported. Avicularia adventitious and pedunculate, associated with all autozooids. Spines and ovicells absent (Fig. 11.12 C, D). Distribution: Tropical and temperate waters. Remarks: There are five epistomiid species reported. The family is characterized by being viviparous. Embryos are incubated intracoelomically in female zooids lacking brood sacs. The reproduction mode of epistomiids is unique among cheilostomes. Family Euoplozoidae Harmer, 1926 Type genus: Euoplozoum Harmer, 1923. Type species: Cellularia cirrata Busk, 1884. Other genera: None. Diagnosis: Colonies erect, biserial, unilamellar, articulated, bifurcating, attached to substratum by rhizoids.

Autozooids arranged in obliquely shifted pairs, weakly calcified, tubular, disjunct but interconnected by cuticular joints. Gymnocyst and cryptocyst absent. Opesia occupying almost the whole frontal area. Embryos incubated in hyperstomial ovicells. Ancestrula not reported. Avicularia adventitious and pedunculate, associated with all autozooids. Spines absent. Distribution: Tropical Indo-Pacific. Remarks: This monospecific family compares well with Epistomiidae Gregory, 1893, but the main difference is that the only representative incubates its embryos in large ovicells rather than intracoelomically in polymorphic zooids. Family Jubellidae Reverter-Gil & Fernández-Pulpeiro, 2001 Type genus: Jubella Jullien, 1882a. Type species: Jubella enucleata Jullien, 1882a. Other genera: None. Diagnosis: Colonies erect, unilamellar, tri- to (rarely) tetraserial, articulated, bifurcating, attached to substratum by rhizoids. Autozooids rectangular. Gymnocyst negligible, smooth. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, covered by tubercles. Opesia subquadrangular. Embryos presumably incubated internally, in enlarged polymorphic zooids. Septula basal, uniporous. Ancestrula autozooidal. Avicularia adventitious, associated with all autozooids, distal to the orifice. Spines and ovicells absent (Fig. 11.13 A, B). Distribution: Temperate Atlantic Ocean. Remarks: Reproduction in Jubella enucleata Jullien, 1882a, the single representative of this family, remains unexplored and incubation of embryos may take place in specialized zooids (Souto et al. 2011). The taxonomic position of Jubellidae remains uncertain and only molecular work may provide a clear answer concerning how to classify this taxon. Family Rhabdozoidae MacGillivray, 1887a Type genus: Rhabdozoum Hincks, 1882a. Type species: Rhabdozoum wilsoni Hincks, 1882a. Other genera: None. Diagnosis: Colonies erect, bilamellar, multiserial, bifurcating, attached to substratum by rhizoids. New branches arising from occluded autozooid and budding a specialized, calyciform, spinose zooid that is the first zooid of the new branch. Autozooids rectangular to slightly hexagonal. Gymnocyst extensive, smooth. Cryptocyst absent. Opesia longitudinally elliptical, about one third of the frontal area. Spines present, proximal of the opesia, usually paired. Embryonic incubation in hyperstomial, acleithral ovicells; ectooecium with large membranous area. Ancestrula erect, vase shaped, attached to substratum by basal disc. Avicularia adventitious, may replace one or both spines.



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Fig. 11.13: (A, B) Jubellidae: Jubella enucleata Jullien, 1882a. Travailleur, Dr. 42, 44° 01′ 20′′ N, 9° 25′ 00′′ W, Bay of Biscay, NE Atlantic Ocean off the Spanish coast. 896 m, 16/8/1881. Muséum National d’Histoire Naturelle, Paris, IB-2008-2634, lectotype. (C, D) Calloporidae: Callopora lineata (Linnaeus, 1758). NE Atlantic Ocean off the British coast. Natural History Museum, London, 1911.10.1.512. Scale bars: A, 1 mm; B, C, 250 µm; D, 50 µm.

Distribution: Temperate Indo-Pacific. Remarks: Rhabdozoidae contains only two species from the waters off southeastern Australia and New Zealand (Hincks 1882a) and South Africa (O’Donoghue & de Watteville 1944). The branching pattern of rhabdozooids is unique as an autozooid in a branch becomes occluded and buds a tubular stalk frontally that forms a calyciform, spinose zooid that gives rise to a new branch. Revisions of the type species Rhabdozoum wilsoni Hincks, 1882a were provided by Gordon (1989a) and Cook and Bock (1994).

11.3.7.3 Superfamily Calloporoidea Norman, 1903a Family Calloporidae Norman, 1903a Type genus: Callopora Gray, 1848. Type species: Flustra lineata Linnaeus, 1758.

Other genera: Adenifera Canu & Bassler, 1917, Alderina Norman, 1903a, Allantocallopora d’Hondt & Schopf, 1985, Allantopora† Lang, 1914, Ammatophora Norman, 1903b, Amphiblestrum Gray, 1848, Apiophragma Hayward & Ryland, 1993, Aplousina Canu & Bassler, 1927, Aviculam­ phiblestrum Rosso, 1999, Barrosia Souto, Reverter-Gil & Fernández-Pulpeiro, 2010, Bidenkapia Osburn, 1950, Bry­ ocalyx Cook & Bock, 2000, Cauloramphus Norman, 1903a, Cavalliella Gordon, 2014, Clavodesia Harmelin & d’Hondt, 1992, Concertina Gordon, 1986, Copidozoum Harmer, 1926, Corbulella Gordon, 1984, Cranosina Canu & Bassler, 1933, Crassimarginatella Canu, 1900, Flustrellaria† d’Orbigny, 1853, Hemiseptella Levinsen, 1909, Leptinatella Cook & Bock, 2000, Marssonopora† Lang, 1914, Membrani­ poridra† Canu & Bassler, 1917, Olisthella Gordon & Taylor, 2017, Onychoblestrum† Gordon, 1984, Parellisina Osburn, 1940, Platypyxis De Blauwe & Gordon, 2014, Pyriporoides

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Hayward & Thorpe, 1989, Ramphonotus Norman, 1894, Recapitulator Gordon, 2014, Septentriopora Kukliński & Taylor, 2006, Tegella Levinsen, 1909, Valdemunitella Canu, 1900, Xylochotridens Hayward & Thorpe, 1989. Diagnosis: Colonies encrusting or (rarely) erect, uni- to multilamellar, multiserial. Autozooids with weak to fairly extensive frontal skeletal calcification, longitudinally rectangular to elliptical. Gymnocyst often extensive, surrounding the opesia from all sides, smooth. Cryptocyst narrow, steeply sloping into the opesia, usually pustulose, in some genera forming a proximal shelf extending over half or more of the frontal area, encircling opesia. Spines present or absent, oral or circumopesial, sometimes overarching the frontal membrane. Basal pore-chambers or septula present. Embryos incubated singly in internal brood sacs with or without vestigial ooecium, or in ovicells. Ovicells hyperstomial, subimmersed or immersed, acleithral, semicleithral, or cleithral; ectooecium calcified with membranous window or mostly membranous. Ancestrula autozooidal. Avicularia present or absent, adventitious, or interzooidal or vicarious, sometimes of more than one type. Kenozooids may be present (Fig. 11.13 C, D). Distribution: Cosmopolitan. Remarks: Calloporidae is a broadly defined family combining many simple cheilostome genera and species with widely varying expressions of frontal skeletal calcification (limited to extensive), avicularian types, and embryonic incubation modes. Despite the obvious variability, it has proven difficult to split the family into subgroups because of numerous intermediate morphologies that link the included genera. Uniting features include both gymnocyst and cryptocyst in varying proportions and the incubation of embryos in ovicells in a majority of genera. Probably only molecular data will resolve splitting of calloporids into several families. The type species of the family, Cal­ lopora lineata, has 8–11 circumopesial spines and distal and lateral pore-chambers. Ovicells are hyperstomial with the ectooecium partly or wholly membranous (Ostrovsky & Schäfer 2003, Ostrovsky et al. 2003, 2009a). Avicularia are adventitious and have a triangular mandible. The ancestrula is single and autozooidal. Apiophragma has a very extensive cryptocyst and is only doubtfully included in Calloporidae. Family Antroporidae Vigneaux, 1949 Type genus: Antropora Norman, 1903b. Type species: Membranipora granulifera Hincks, 1880b. Other genera: Akatopora† Davis, 1934, Parantropora Tilbrook, 1998, Rosseliana Jullien, 1888. Diagnosis: Colonies encrusting, uni- to (rarely) multilamellar, multiserial. Autozooids lacking or (rarely)

moderate frontal wall calcification, longitudinally rectangular to elliptical. Gymnocyst absent or negligible, smooth. Cryptocyst narrow, steeply sloping into the opesia, usually pustulose, in some genera forming a proximal shelf extending over half or more of the frontal area, encircling opesia. Pore-chambers basal. Embryos incubated in immersed cleithral ovicells. Larva non-feeding. Ancestrula autozooidal. Avicularia usually present, adventitious or interzooidal or vicarious. Kenozooids may be present. Spines typically absent (Fig. 11.14 A, B). Distribution: Cosmopolitan. Remarks: The validity of Antroporidae is disputed and it only recently has been accepted as a family separate from Calloporidae (e.g. Cook et  al. 2018). Discrimination from the latter may be difficult as most morphological characters found in antroporids can be observed in some calloporids. However, antroporids have immersed ovicells and spines are typically lacking, in contrast to the large, hyperstomial ovicells and periopesial spines of the type calloporid Callopora lineata. Antroporidae is a useful family for subdivision of the calloporid complex. Family Bryopastoridae d’Hondt & Gordon, 1999 Type genus: Bryopastor Gordon, 1982. Type species: Heterocella pentagonus Canu & Bassler, 1929a. Other genera: Omoiosia Canu & Bassler, 1927, Pseudothy­ racella† Labracherie, 1975. Diagnosis: Colonies erect, vincularian, oligo–multiserial, unbranched or with jointed branches, attached to substratum by rhizoids. Autozooids in alternating longitudinal series, longitudinally rectangular to pyriform. Gymnocyst absent. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, granular, encircling opesia. Opesia longitudinally elliptical to pyriform. Deep recesses for operculum occlusor muscles common, paired. No spines. Septula present, multiporous, in vertical walls. Embryos incubated internally in polymorphic zooids (presumably in brood sacs) or in immersed ovicells. Ancestrula autozooidal, rooted by basal rhizoids. Avicularia usually absent, if present, vica­rious. Spines absent (Fig. 11.14 C, D). Distribution: Tropical Pacific Ocean. Remarks: Bryopastoridae was proposed for erect, rooted cheilostomes with shelf-like cryptocyst, immersed ovicells, no spines, and usually lacking avicularia. The status of Omoiosia Canu & Bassler, 1927 is uncertain, pending revision. Cookinella d’Hondt, 1981 (currently in Membranicellariidae) and Acanthodesiomorpha d’Hondt, 1981 (unassigned) may possibly also be included in Bryopastoridae when revised (d’Hondt & Gordon 1999).



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Fig. 11.14: (A) Antroporidae: Antropora minor (Hincks, 1880a). Bahia, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade Federal da Bahia, 0408. (B) Antroporidae: Akatopora leucocypha (Marcus, 1937). São Paulo, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, 0702. (C, D) Bryopastoridae: Bryopastor pentagonus (Canu & Bassler, 1929a). Celebes Sea, W Pacific Ocean. Muséum National d’Histoire Naturelle, unregistered. © http://bryozoa.net. Scale bars: A, B, D, 250 µm; C, 500 µm.

Family Chaperiidae Jullien, 1888 Type genus: Chaperia Jullien, 1881. Type species: Chaperia australis Jullien, 1881. Other genera: Chaperiopsis Uttley, 1949, Clipeochaperia Uttley & Bullivant, 1972, Exallozoon Gordon, 1982, Exos­ tesia Brown, 1948, Icelozoon Gordon, 1982, Larnacicus Norman, 1903b, Notocoryne Hayward & Cook, 1979, Pat­ syella† Brown, 1948, Pyrichaperia Gordon, 1982. Diagnosis: Colonies encrusting, unilamellar, uni- to multiserial, loosely attached to substratum; or erect from an encrusting base, vincularian or bilamellar, nonarticulated, dichotomously branching. Autozooids longitudinally elliptical to hexagonal. Gymnocyst present or restricted to area around spine bases, smooth, sometimes bearing adventitious avicularia. Cryptocyst forming

a proximal shelf extending over half or more of the frontal area, smooth to granular, encircling opesia. Opesia longitudinally elliptical. Occlusor laminae paired. Spines present or absent, oral, articulated, sometimes bifurcating. Septula present, multiporous, in vertical walls. Incubation of embryos in hyperstomial or endozooidal acleithral ovicells or internally (presumably in brood sacs, sometimes associated with vestigial ooecia). Ancestrula autozooidal. Avicularia present or absent, adventitious or interzooidal. Kenozooids may be present (Fig. 11.15 A, B). Distribution: Cosmopolitan; predominantly temperate to polar waters of the Southern Hemisphere. Remarks: All chaperiids have paired occlusor laminae, i.e. skeletal plates for the accommodation of the opercular occlusor muscles. This is the main character for

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Fig. 11.15: (A, B) Chaperiidae: A, Chaperia n. sp. NIWA 65975, NIWA Stn TRIP3135/6, 48.3433° S, 168.1567° E. B, Chaperiopsis n. sp, Three Kings shelf, northern New Zealand. (C, D) Cupuladriidae: C, Cupuladria biporosa (Canu & Bassler, 1923), Caribbean, AT-05-186-2, photo: P.D. Taylor. D, Cupuladria guineensis (Busk, 1854) NHMUK, Queensland. Photo: P.E. Bock. Scale bars: C, 1 mm; A, B, 500 µm; D, 250 µm.

identification of the family. The occurrence of occlusor laminae, however, is not unique for chaperiids, but they can also be found in bryopastorids (weakly developed) and some onychocellids, e.g. Rhagasostoma hexagonum Koschinsky, 1885 from the Lutetian of Bavaria, Germany (see Taylor et al. 2018). Family Cupuladriidae Lagaaij, 1952 Type genus: Cupuladria Canu & Bassler, 1919. Type species: Cupularia canariensis Busk, 1859a. Other genera: Discoporella† d’Orbigny, 1852, Reussirella† Bałuk & Radwański, 1984. Diagnosis: Colonies cup-shaped, convex frontally, concave basally, unilamellar, multiserial, free-living, motile, supported by vibracular mandibles. Autozooids

arranged in alternating series radiating from the center, longitudinally hexagonal to elliptical, opening on the convex surface. Gymnocyst negligible, smooth. Cryptocyst narrow, steeply sloping into the opesia, usually pustulose, in some genera extending over half or more of the frontal area, encircling opesia. Basal surface bounded by cuticle covering extrazooidal coelom, sometimes thickened by secondary calcification. Septula present, multiporous, in vertical and basal walls. Embryos brooded in internal brood sacs. Larva non-feeding. Reproduction also by asexual peripheral budding and regeneration after fragmentation. Ancestrula triad, located in the center of the colony, may be replaced by vicarious avicularia. Other vicarious avicularia may be present. Vibracula distal of each autozooid, asymmetrical, bearing setiform



mandibles. Kenozooids may be present, usually on the basal side. Spines absent (Fig. 11.15 C, D). Distribution: Tropical waters. Remarks: Cupuladriids show convergent evolution with the lunulitiform morphotype but are systematically distinct from Lunulitoidea Lagaaij, 1952 and Mamilloporoidea Canu & Bassler, 1927. Cupuladriids are distinguished from other lunulitiform taxa in having a triad ancestrula, no ovicells and the basal surface being divided into porous sectors. The oldest known cupuladriid species, Cupuladria ovalis Gorodiski & Balavoine, 1962, was found in Palaeocene sediments from Senegal and was already highly integrated. The family therefore probably originated in the Latest Cretaceous (Cook & Chimonides 1983). Cupuladriids spread during the Palaeogene and Neogene and are nowadays the most widespread lunulitiform bryozoans with high abundances in tropical and subtropical seas. Lunulitoids, on the other hand, are now restricted to Australasia. A revision of Cupuladriidae was provided by Cook & Chimonides (1994a), who assigned three genera to the family. Colonial and zooidal characteristics of species attributed to Vibracellina Canu & Bassler, 1917, a genus previously assigned to Cupuladriidae by Winston & Vieira (2013), suggest it to be better assigned to Heliodomidae Vigneaux, 1949 than Cupuladriidae (see below). Family Cymuloporidae Winston & Vieira, 2013 Type genus: Cymulopora Winston & Håkansson, 1986. Type species: Cymulopora uniserialis Winston & Håkansson, 1986. Other genera: Crepis Jullien, 1882a. Diagnosis: Colonies encrusting, unilamellar, uniserial. Autozooids arranged in a single row, longitudinally elliptical to pyriform. Gymnocyst surrounding opesia on all sides, smooth, forming a short or usually long cauda proximally. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, granular. Opesia longitudinally semielliptical to bell shaped; may have opesiular constrictions or indentations proximolaterally. Septula present, uniporous, in vertical walls. Embryos incubated in immersed or hyperstomial cleithral ovicells. Ancestrula unknown in most species, presumably autozooidal. Avicularia, kenozooids, and spines absent (Fig. 11.16 A, B). Distribution: Tropical waters. Remarks: Previously included in Calloporidae, Winston & Vieira (2013) separated Cymulopora and Crepis from the latter to create a new family. Cymuloporid colonies are uniserial, and autozooids have an extensive gymnocyst that forms a cauda proximally. The cryptocyst always extends over the frontal area as a granular shelf proximal

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to the opesia. Cymuloporidae are distinctive in lacking spines, kenozooids and avicularia. Family Doryporellidae Grischenko, Taylor, & Mawatari, 2004 Type genus: Doryporella Norman, 1903b. Type species: Lepralia spathulifera Smitt, 1868b. Other genera: Doryporellina Grischenko, Mawatari & Taylor 2000. Diagnosis: Colonies encrusting, unilamellar, multiserial. Autozooids longitudinally elliptical to hexagonal. Gymnocyst negligible, smooth. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, ornamented with a reticulate pattern of polygonal ridges, encircling opesia. Opesia longitudinally semielliptical to bell shaped. Spines oral, articulated, sometimes bifurcating. Pore-chambers basal. Embryos incubated in hyperstomial ovicells with mostly membranous ectooecium and calcified endooecium. Ancestrula modified tatiform. Avicularia present, adventitious. Kenozooids not reported (Fig. 11.16 C, D). Distribution: Polar and temperate waters of Northern Hemisphere. Remarks: Doryporellidae was established for two genera previously included in Calloporidae. The main character distinguishing doryporellids from other calloporoid families is the reticulate ornamentation of the cryptocyst, comprising polygonal ridges. Ovicells show the same type of ornamentation on the endooecium. Family Ellisinidae Vigneaux, 1949 Type genus: Ellisina Norman, 1903a. Type species: Membranipora levata Hincks, 1882c. Other genera: Kenoaplousina López Gappa & Liuzzi, 2013, Lamourouxia d’Hondt & Gordon, 1999, Retevirgula Brown, 1948. Diagnosis: Colonies encrusting, unilamellar, multiserial; or erect, unilamellar, bi- to oligoserial, directly attached to substratum. Autozooids longitudinally rectangular to elliptical. Gymnocyst negligible laterally, more developed proximally, smooth. Cryptocyst narrow, steeply sloping into the opesia, usually pustulose, encircling opesia. Spines present or absent, oral or circumopesial. Pore-chambers basal. Embryos incubated in hyperstomial to subimmersed cleithral ovicells, often terminal. Ectooecium completely or mostly calcified. Ancestrula autozooidal. Avicularia interzooidal or absent. Kenozooids may be present (Fig. 11.17 A, B). Distribution: Cosmopolitan. Remarks: Ellisinidae Vigneaux, 1949, a family previously regarded as a junior synonym of Calloporidae Norman,

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Fig. 11.16: (A, B) Cymuloporidae: Cymulopora uniserialis Winston & Håkansson, 1986. São Sebastião, São Paulo, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, 0709. (C, D) Doryporellidae: Doryporella smirnovi Grischenko, Taylor & Mawatari, 2004. Bering Sea, NW off Bering Island, Commander Islands, 55° 25.2′ N, 165° 33.8′ E, depth 152 m. Zoological Institute, Russian Academy of Science, Saint Petersburg, ZIRAS 1/50130, holotype. Scale bars: A, D, 250 µm; B, 50 µm; C, 500 µm.

1903a, has recently been revalidated for cheilostome species with ooecia associated with an avicularium (Cook et al. 2018) or kenozooid. Family Farciminariidae Busk, 1852b Type genus: Farciminaria Busk, 1852b. Type species: Farciminaria aculeata Busk, 1852b. Other genera: Columnella Levinsen, 1914, Didymozoum Harmer, 1923, Farciminellum Harmer, 1926. Diagnosis: Colonies erect, unilamellar or bilamellar, biserial, quadriserial or oligoserial, articulated or non-articulated, bifurcating, attached to substratum by rhizoids. Zooids arranged back to back in bilamellar colonies, weakly calcified, longitudinally rectangular. Gymnocyst lacking or vestigial, smooth. Cryptocyst absent in autozooids, but present in kenozooids, pustulose. Opesia

typically occupying the whole frontal area. Orifice subterminal, transversely semielliptical. Spines present or absent, oral or circumopesial. Embryos incubated in internal brood sacs of enlarged autozooids or in hyperstomial acleithral ovicells. Ancestrula autozooidal. Avicularia usually absent: if present, adventitious, associated with gymnocyst. Kenozooids present, accumulated on the basal surface in bilamellar colonies. Distribution: Cosmopolitan. Remarks: Farciminariidae comprises only erect taxa that are attached to the substratum by rhizoids. Kenozooids form the basal surface in bilamellar colonies. Most characters are largely variable in this family and the type genus has cylindrical, oligoserial, non-articulated branches, hyperstomial ovicells and may or may not have marginal spines. Avicularia are lacking.



11.3 Systematics of Cheilostomata 

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Fig. 11.17: (A, B) Ellisinidae: Ellisina gautieri (Fernández‐Pulpeiro & Reverter-Gil, 1993). Ría de Ferrol, Spain, E Atlantic Ocean, unregistered. (C, D) Heliodomidae: Setosellina roulei Calvet, 1906. Morocco, 33° 9′ 0.0072′′ N, 9° 37′ 59.988′′ E, NE Atlantic Ocean. Muséum National d’Histoire Naturelle, Paris, IB-2008-1979, syntype. Scale bars: A, C, 500 µm; B, 250 µm; D, 100 µm.

Family Foveolariidae Gordon & Winston, 2005 in Winston (2005) Type genus: Foveolaria Busk, 1884. Type species: Foveolaria elliptica Busk, 1884. Other genera: Amplexicamera Winston, 2005, Dacty­ lostega Hayward & Cook, 1983, Mangana Gordon, 2014, Odontionella Canu & Bassler, 1917. Diagnosis: Colonies encrusting, unilamellar, oligo- to multiserial; or erect, cylindrical, articulated; or erect from an encrusting base, bilamellar, dichotomously branching. Autozooids longitudinally elliptical. Gymnocyst negligible, smooth. Cryptocyst narrow to extensive, steeply sloping into the opesia, pustulose, encircling opesia; or scattered areas of the interior (cryptocystal) wall on the sides of avicularia and ovicells. Spines in some species, oral, ephemeral. Pore-chambers basal. Embryos incubated in hyperstomial, subimmersed or immersed ovicells. In some species ooecia sunken in secondary calcification in old colonies.

Ancestrula not reported. Avicularia adventitious or interzooidal or absent. Kenozooids may be present. Distribution: Predominantly Southern Hemisphere. Remarks: Mangana was originally assigned to the Calloporidae (Gordon 2014). However, it should be included in Faveolariidae owing to the presence of a large oval opesia, surrounded by a narrow cryptocyst, proximally adventitious avicularia and scattered areas of interior (cryptocystal) wall on avicularia and ovicells. Family Heliodomidae Vigneaux, 1949 Type genus: Heliodoma Calvet, 1906. Type species: Heliodoma implicata Calvet, 1906. Other genera: Setosellina Calvet, 1906, Vibracellina† Canu & Bassler, 1917. Diagnosis: Colonies encrusting, unilamellar, multiserial, sometimes becoming free at the periphery. Autozooids arranged in spirals around the ancestrula during early

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astogeny and in regular series in later astogeny, longitudinally hexagonal to elliptical. Gymnocyst negligible, smooth. Cryptocyst narrow, pustulose, steeply sloping into the opesia and encircling opesia. Pore-chambers usually present, basal. Embryos incubated in immersed or subimmersed ovicells or internally (presumably in brood sacs). Ancestrula autozooidal, budding two autozooids and two avicularia. Avicularia interzooidal, distal or distolateral to each autozooid, bearing setiform mandible. Spines and kenozooids absent (Fig. 11.17 C, D). Distribution: Tropical to temperate waters of Atlantic and Indian oceans. Remarks: Ovicells may be present or absent (Harmer 1926, Harmelin 1977). The zooidal and avicularian characters of Vibracellina capillaria Canu & Bassler, 1917, the type species of Vibracellina, suggest that the genus is better attributed to Heliodomidae than to Cupuladriidae.

Interzooidal avicularia with setiform mandibles (vibracula) are also present in Cupuladriidae, but this family is distinct in having a triad ancestrula, no ovicells (Ostrovsky et  al. 2009b) and a basal surface 90° divided into porous sectors. Family Hiantoporidae Gregory, 1893 Type genus: Hiantopora MacGillivray, 1887a. Type species: Lepralia ferox MacGillivray, 1869. Other genera: Tremopora Ortmann, 1890. Diagnosis: Colonies loosely encrusting, unilamellar, multiserial, supported above the substratum by rhizoids emanating from septular pores. Autozooids longitudinally polygonal, contiguous or interconnected by calcified tubes. Gymnocyst surrounding opesia on all sides, smooth. Cryptocyst narrow, steeply sloping into the opesia, pustulose, encircling opesia. Spines single or pairwise, generally simple but a proximal spine may also branch, partly covering the

Fig. 11.18: (A, B) Hiantoporidae: Hiantopora sp. Bahia, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade Federal da Bahia, unregistered. (C, D) Mourellinidae: Mourellina decussata (Harmer, 1926). Siboga, Station 227, western end of Ceram, Banda Sea, Indonesia. Zoölogisch Museum, Universiteit van Amsterdam, V.Bry. 1811, lectotype. Scale bars: A, C, 500 µm; B, 250 µm; D, 100 µm.



opesia. Pore-chambers multiporous. Embryos incubated in hyperstomial or subimmersed cleithral ovicells, sometimes reminiscent of immersed or endozooidal ovicells because of frontal spine development above ooecium; ectooecium with large membranous area (fenestra). Ancestrula not reported. Avicularia adventitious, single or paired and unequal in size, sometimes spinose, partly or mostly concealing membranous frontal wall (Fig. 11.18 A, B). Distribution: Tropical to temperate Indo-Pacific. Remarks: Harmer (1926) regarded Hiantopora and Tremopora as synonymous, but differences in ovicell structure suggest they comprise two different genera (Cook et al. 2018). Family Mourellinidae Reverter-Gil, Souto, & FernándezPulpeiro, 2011 Type genus: Mourellina Reverter-Gil, Souto & Fernández-Pulpeiro, 2011.

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 347

Type species: Crepis decussata Harmer, 1926. Other genera: None. Diagnosis: Colonies erect, cylindrical, uniserial, dichotomously branching; or loosely encrusting, unilamellar, uniserial. Autozooids weakly calcified, tubular, with decumbent proximal portion and projecting distal portion. Cryptocyst reduced, projecting into body cavity. Operculum terminal, bearing a crescentic marginal sclerite. Internal spicules may be present, projecting into body cavity. Septula uniporous, in vertical and basal walls. Embryonic incubation uncertain. Ancestrula not reported. Avicularia present or absent, vicarious. Spines absent (Fig. 11.18 C, D). Distribution: Banda Sea. Remarks: This family contains only two closely related species, both of which are endemic to the Banda Sea. Colonies are weakly calcified and uniserial, while the autozooids may possess spicules projecting into the body cavity.

Fig. 11.19: (A, B) Pyrisinellidae: A, Spinisinella zagorseki Di Martino & Taylor, 2012. Dresden-Plauen, Germany, late Cenomanian. Senckenberg Naturhistorische Sammlungen Dresden, SaK 15776. B, Microblestrum imitator Gordon, 2014, NIWA Stn TAN0803/98, 700m. (C, D) Quadricellaria bocki (Silén, 1942) NIWA 26701, NIWA Stn TAN0408/22, 42.7470° S, 177.8978° W, 963 m. Scale bars: A, C, 500 µm; D, 200; B, 150 µm.

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Family Pyrisinellidae† Di Martino & Taylor, 2012 Type genus: Pyrisinella† Di Martino & Taylor, 2012. Type species: Setosinella meniscacantha† Taylor & McKinney, 2006. Other genera: Megapora Hincks, 1877a, Microblestrum Gordon, 2014, Ristedtia Matsuyama, Martha, Scholz, & Hillmer, 2017, Stolomicropora Gordon, 2014. Diagnosis: Colonies encrusting, unilamellar, multiserial. Autozooids longitudinally hexagonal to oval. Gymnocyst narrow, surrounding the whole cryptocyst, tapering distally, smooth. Cryptocyst forming a proximal shelf extending to the orifice, granular, surrounded by a mural rim. Opesia trifoliate or semielliptical, subterminal. Opesiules may occur, circular, as a single pair or in series. Spines oral, distal to the orifice. Embryos incubated in hyperstomial ovicells. Pore-chambers basal. Ancestrula autozooidal, but with variable number of spines, budding one autozooid distally. Avicularia absent or present, adventitious or interzooidal or vicarious, irregularly interspersed among autozooids. Kenozooids may be present (Fig. 11.19 A, B). Distribution: Polar waters of the Atlantic and Pacific oceans. Remarks: Both known species of the type genus are Maastrichtian and Ypresian, respectively. The family is closely related to Onychocellidae but differs from the latter in always having oral spines and in showing a smooth gymnocyst around the opesia. However, the boundaries between the two families vanish in some fossil material as species of the genus Holpitaechmella Voigt, 1949 have six oral spines and hyperstomial ovicells and they may show a salient mural rim that separates the granular cryptocyst from the surrounding, smooth gymnocyst. A manuscript by Ehrhard Voigt reviewing the species and characters of Hoplitaechmella and entitled “Das Genus Hoplitaechmella Voigt (Bryozoa, Cheilostomata) in der oberen Kreide und im ältesten Tertiär (Danium)” remained unfortunately unfinished. Apiophragma Hayward & Ryland, 1993 (currently in Calloporidae) may possibly belong to Pyrisinellidae. Family Quadricellariidae† Gordon, 1984 Type genus: Quadricellaria† d’Orbigny, 1851. Type species: Quadricellaria elegans† d’Orbigny, 1851. Other genera: Nellia Busk, 1852b, Nelliella Mawatari, 1974. Diagnosis: Colonies erect, quadriserial, articulated, dichotomously branching. Autozooids arranged in four longitudinal series, rectangular. Gymnocyst reduced, proximal only, smooth. Cryptocyst forming a proximal shelf extending over half or more of the frontal area. Embryos incubated internally (presumably in brood sacs) or in ovicells, subimmersed, immersed or endozooidal. Ancestrula autozooidal, erect on uncalcified radicular

support. Avicularia absent or present, adventitious or interzooidal. Kenozooids may be present. Spines absent (Fig. 11.19 C, D). Distribution: Tropical waters. Remarks: Quadricellariidae are morphologically very simple cheilostomes characterized by erect, quadrilateral and articulated colonies. The family is based on an easily recognizable and very common species from the Late Cretaceous of France. Huge gaps in the fossil record between Late Cretaceous and Recent quadricellariids and the simplicity of colonies raise the question of monophyly of this group; convergent evolution cannot be excluded. Embryonic incubation is supposedly in internal brood sacs in Quadricel­ laria. Subimmersed ovicells are reported only in one fossil genus, Cellarinidra Canu & Bassler, 1927, and endozooidal and immersed cleithral ovicells with frontal fenestra in the ectooecium are known in extant species of Nellia, while the ancestrula has been observed only in Recent material.

11.3.7.4 Superfamily Cellarioidea Lamouroux, 1816 Family Cellariidae Lamouroux, 1816 Type genus: Cellaria Ellis & Solander, 1786. Type species: Farcimia sinuosa Hassall, 1840. Other genera: Atelestozoum Harmer, 1926, Cryptostoma­ ria Canu & Bassler, 1927, Dubiocellaria d’Hondt & Schopf, 1985, Euginoma Jullien, 1882a, Formosocellaria d’Hondt, 1981, Melicerita Milne Edwards, 1836a, Melicerita (Hen­ rimilnella) d’Hondt & Gordon, 1999, Mesostomaria Canu & Bassler, 1927, Paracellaria Moyano G., 1969, Smitticellaria Gordon & Taylor, 1999, Steginocellaria David & Pouyet, 1986, Stomhypselosaria Canu & Bassler, 1927, Swanomia Hayward & Thorpe, 1989, Syringotrema Harmer, 1926. Diagnosis: Colonies erect, cylindrical, bilamellar, oligo-multiserial, articulated or non-articulated, dichotomously branching. Autozooids longitudinally rhomboidal to hexagonal. Gymnocyst absent. Cryptocyst forming a large shelf proximal to the orifice, depressed, granular, surrounded by a mural rim, encircling opesia. Opesia coincident with the orifice, subterminal, semicircular to transversely elliptical, usually with paired denticles in the proximolateral (and distolateral) corners. Spines usually absent; in at least one species, uncalcified spines originating proximally. Embryos incubated in endotoichal ovicells. Larva non-feeding. Ancestrula autozooidal. Avi­ cularia absent or present, interzooidal or vicarious, interspersed among autozooids. Kenozooids may be present, at termination of zooid rows (Fig. 11.20 A, B). Distribution: Cosmopolitan, but predominantly Southern Hemisphere.



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Fig. 11.20: (A, B) Cellariidae: Cellaria subtropicais Vieira, Gordon, Souza & Haddad, 2010. Rio de Janeiro, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, 045, holotype. (C, D) Lunulitidae: Lunulites plana d’Orbigny, 1852. Villedieu-le-Château, France, Santonian. Natural History Museum, London, D53532 (two specimens). Scale bars: A, C, 500 µm; B, 100 µm; D, 250 µm.

Remarks: Lamouroux’s “Cellariées/Cellarieæ” (1816, p. 117) was first used as a taxonomic name to group genera Cellariidae; thus, it fulfills the provisions of Article 11.7 of the International Code on Zoological Nomenclature (ICZN 1999) and this name is available with its original author and date (see also Remarks for Flustridae Lamouroux, 1816). Cellariids are characterized by cylindrical to bilamellar, erect colonies with rhomboidal or hexagonal autozooids bearing paired denticles in the proximolateral (and distolateral) corners of the orifice. They possess endotoichal brood chambers that have an additional “operculum”-like sclerite distal to the maternal orifice and closing the ovicell (Calvet 1900, Ostrovsky 2013). This ovicell type is found solely in Cellariidae and Membranicellariidae, the latter family being considered by some authors as junior synonym of Cellariidae (e.g. Hayward & Winston 2011).

Family Membranicellariidae Levinsen, 1909 Type genus: Membranicellaria Levinsen, 1909. Type species: Melicerita dubia Busk, 1884. Other genera: Cookinella d’Hondt, 1981. Diagnosis: Colonies erect, bilamellar, multiserial, non-articulated, dichotomously branching, attached to substratum by kenozooidal rhizoids. Autozooids longitudinally rhomboidal to hexagonal. Gymnocyst absent. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, depressed, granular, surrounded by a mural rim, encircling opesia. Opesia central, longitudinally elliptical. Septula in vertical and basal walls, multiporous. Embryos incubated in endotoichal ovicells. Ancestrula not reported. Avicularia vicarious, interspersed among autozooids. Kenozooids may occur. Spines absent. Distribution: Temperate to polar southern Atlantic Ocean.

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 11 Gymnolaemata, Cheilostomata

Remarks: Membranicellariids comprise four species. The family is considered a possible junior synonym of Cellariidae Lamouroux, 1821 (cf. Hayward & Winston 2011). In contrast to cellariids, the opesia is comparatively larger and centrally located. The ovicell is endotoichal as in cellariids, showing the close relationship between the two families.

11.3.7.5 Superfamily Lunulitoidea† Lagaaij, 1952 Remarks: The type family of Lunulitoidea, Lunulitidae Lagaaij, 1952, contains only genera with no Recent representatives, the type species of Lunulites Lamarck, 1816, Lunulites radiata Lamarck, 1816, being from the Miocene of France. The superfamily name was first used in Cook et  al. (2018) to group together four families of lunulitiform cheilostomes, while other authors usually included it in Microporoidea (e.g. Martha et al. 2019b). The term lunulitiform is restricted to free-living discoidal or cup-shaped colonies supported by avicularian mandibles. Family Lunulariidae Levinsen, 1909 Type genus: Lunularia Busk, 1884. Type species: Lunulites capulus Busk, 1852a. Other genera: None. Diagnosis: Colonies cup-shaped, convex frontally, concave basally, unilamellar, multiserial, free-living, motile, supported by avicularian mandibles. Basal colony surface with multiporous pore-sectors. Autozooids arranged in an alternating series radiating from the center, longitudinally rectangular to elliptical, opening on the convex surface. Gymnocyst absent. Cryptocystal shelf relatively small, steeply sloping into the opesia, granular or dentate, encircling opesia. Septula present, multi­ porous, in vertical and basal walls. Embryos incubated in immersed ovicells or interior brood sacs. Reproduction also by asexual peripheral budding and regeneration after fragmentation. Ancestrula autozooidal, located in the center of the colony. Avicularia vicarious, alternating with autozooids, bearing elongated mandibles that may trifurcate at the tips. Spines absent (Fig. 11.20 C, D). Distribution: Temperate waters off southern Australia. Remarks: The family is represented by only two Recent species off southern Australian waters, one of them also on the Kermadec Ridge (Gordon 1985). In contrast to Otionellidae Bock & Cook, 1998 and Selenariidae Busk, 1854, lunulariids lack skeletally distinct male zooids. Family Otionellidae Bock & Cook, 1998 Type genus: Otionella† Canu & Bassler, 1917. Type species: Otionella perforata† Canu & Bassler, 1917.

Other genera: Helixotionella† Cook & Chimonides, 1984, Otionellina Bock & Cook, 1998, Petasosella Bock & Cook, 1998. Diagnosis: Colonies domed or cup-shaped, convex frontally, concave or flat basally, unilamellar, multiserial, free-living, supported by avicularian mandibles. Autozooids arranged in an alternating series radiating from the center or spiralled, longitudinally hexagonal to elliptical, opening on the convex surface. Gymnocyst negligible, smooth. Cryptocyst extensive, forming a proximal shelf extending over half or more of the frontal area, granular, encircling opesia. Opesiules may occur, circular, single or paired. Septula present, multiporous, in vertical and basal walls. Embryos incubated in interior brood sacs; brooding zooids may be slightly enlarged. Reproduction also by asexual peripheral budding and regeneration after fragmentation. Ancestrula autozooidal, located in the center of the colony. Avicularia interzooidal or vicarious, with paired condyles that may fuse at the tips, bearing elongated mandibles, cryptocyst usually denticulate. Spines absent. Distribution: Tropical to temperate waters around Australia and New Zealand. Remarks: All representatives of the type genus are fossils from the Eocene to Miocene. Otionellidae combines characters of both Lunulitidae Lagaaij, 1952 and Selenariidae Busk, 1854; however, whereas female zooids may be skele­ tally distinct, males may not be so. Family Selenariidae Busk, 1854 Type genus: Selenaria Busk, 1854. Type species: Lunulites maculata Busk, 1852a. Other genera: Pseudolunularia Cadée, Chimonides & Cook, 1989. Diagnosis: Colonies cup-shaped or discoidal, convex frontally, concave or flat basally, unilamellar, multiserial, free-living, motile, supported by avicularian mandibles, partitioned into concentric astogenetic zones of zooids with differing functions; central zone of closed zooids, followed by zone of autozooids, subperipheral zone of female brooding zooids and peripheral zone of non-feeding male zooids. Basal colony surface with scattered pores. Autozooids arranged in an alternating series radiating from the center, longitudinally hexagonal to elliptical, opening on the convex surface. Gymnocyst negligible, smooth. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, granular, encircling opesia. Opesia usually longitudinally elliptical, sometimes with opesiular indentations in the proximolateral corners. Opesiules may occur, circular, as a single pair. Septula present, multiporous, inside vertical and basal walls. Embryos incubated in endozooidal



or immersed ovicells. Reproduction also by asexual peripheral budding and regeneration after fragmentation. Ancestrula autozooidal, located in the center of the colony, surrounded by six zooids. Avicularia vicarious, asymmetrical, bearing setiform mandibles, cryptocyst porous. Spines absent. Distribution: Tropical to temperate waters around Australia and New Zealand. Remarks: Selenaria is very distinctive from other lunulitiform taxa as colonies are partitioned into concentric astogenetic zones of zooids with differing functions (see Cook & Chimonides 1987, Bock & Cook 1999). Both female and male zooids are skeletally distinct and arranged in zones – males being peripheral and females subperipheral. Avicularia are large and isolated, scattered in patterns among the other zooids.

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11.3.7.6 Superfamily Microporoidea Gray, 1848 Family Microporidae Gray, 1848 Type genus: Micropora Gray, 1848. Type species: Flustra coriacea Esper, 1790. Other genera: Andreella Jullien, 1888, Coronellina Prenant & Bobin, 1966, Flustrapora Moyano G., 1970a, Metamicropora† Arakawa, 2016, Microporina Levinsen, 1909, Mollia Lamouroux, 1816, Opaeophora Brown, 1948, Otomicropora Gordon, 2014, Promicroa d’Hondt & Gordon, 1999, Rectimicropora Hayward & Winston, 2011, Reussinella Gordon, 2009b, Rosemariella Gordon, 2014, Scriblitopora Hayward & Winston, 2011, Steraechmella† Lagaaij, 1952. Diagnosis: Colonies encrusting, unilamellar, multiserial; or erect from an encrusting base, frondose, bilamellar or

Fig. 11.21: (A, B) Microporidae: Micropora angustiscapulis Winston, Vieira & Woollacott, 2014. N of Salvador, Bahia, Brazil, W Atlantic Ocean. Museum of Comparative Zoology Cambridge, USA, 137433, holotype. (C, D) Aspidostomatidae: Crateropora sp. NIWA 23297 TAN0413/171 37.4525 S, 176.9040 E, 310-410 m. Scale bars: C, 500 µm; A, D, 250 µm; B, 50 µm.

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 11 Gymnolaemata, Cheilostomata

cylindrical, articulated or non-articulated. Autozooids longitudinally pyriform to hexagonal. Gymnocyst absent or vestigial. Cryptocyst forming a proximal shelf extending usually over most of the frontal area, granular, sometimes pseudoporous, encircling opesia. Opesia usually coincident with the orifice, semicircular. Opesiules usually present, circular, as a single pair or in series. Spines in some species, oral, distolateral of orifice. Basal pore-chambers or mural septula present. Embryos incubated in hyperstomial or subimmersed ovicells or, presumably, in internal brood sacs. Ancestrula autozooidal. Avicularia may be present, interzooidal or vicarious, irregularly interspersed among autozooids. Kenozooids may be present (Fig. 11.21 A, B). Distribution: Cosmopolitan. Remarks: Microporidae is a very large and heterogeneous family. Species are characterized by encrusting or erect colonies with autozooids having an extensive cryptocystal shelf covering most of the frontal area and usually pierced by a pair or by rows of circular opesiules. Calpensiidae Canu & Bassler, 1923, which was previously regarded as a junior synonym of Microporidae, is considered valid. Family Alysidiidae Levinsen, 1909 Type genus: Alysidium Busk, 1852b. Type species: Alysidium parasiticum Busk, 1852b. Other genera: Catenicula O’Donoghue, 1924. Diagnosis: Colonies erect, uniserial, articulated; stems from small enlarged parts of a reticulate, encrusting stoloniform base. Autozooids longitudinally pyriform. Gymnocyst forming a long proximal cauda, smooth. Cryptocyst forming a proximal shelf extending usually over most of the frontal area, encircling opesia. Opesia usually coincident with the orifice, semicircular. Opesiules usually present, circular, as a single pair. Spines in one species, oral. Embryos incubated in compound brood chambers (sometimes ovicell-like). Ancestrula not reported. Avicularia may be present, interzooidal or vicarious, irregularly interspersed among autozooids. Spiniform kenozooids may also be present. Distribution: Temperate waters off the South African coast. Remarks: All three alysidiid species require reillustration with scanning electron microscope (SEM) images and the family needs proper revision. The kenozooidal origin of their ovicells has been studied by Levinsen (1909). Ostrovsky (2013) suggested their independent evolution from conventional flustrine ovicells. Family Aspidostomatidae Jullien, 1888 Type genus: Aspidostoma Hincks, 1881a. Type species: Aspidostoma crassum Hincks, 1881a (= Eschara gigantea Busk, 1854).

Other genera: Crateropora Levinsen, 1909, Lagarozoum Harmer, 1926, Larvapora Moyano G., 1970b. Diagnosis: Colonies encrusting, unilamellar, multiserial; or erect from an encrusting base, frondose, bilamellar or cylindrical, bifurcating. Autozooids longitudinally pyriform to hexagonal. Gymnocyst absent or negligible, smooth. Cryptocyst forming a proximal shelf extending over most of the frontal area, granular, encircling opesia. Opesia usually coincident with the orifice, transversely elliptical to semicircular, usually with opesiular indentations in the proximolateral corners. Basal pore-chambers may be present. Embryos incubated in hyperstomial or endozooidal ovicells. Ancestrula not reported. Avicularia may be present, adventitious or interzooidal, irregularly interspersed among autozooids. Articulated spines absent (Fig. 11.21 C, D). Distribution: Tropical and polar waters predominantly of the Southern Hemisphere. Remarks: Challenges in distinguishing between the families Aspidostomatidae and Onychocellidae Jullien, 1882b have been discussed in Gordon & Taylor (2015). They concluded that aspidostomatids are typically well calcified, have a well-defined orifice-like opesia with proximolateral indentations, non-vicarious avicularia with mandibular pivots and an ovicell with a separate opening. Onychocellids sensu stricto typically have an autozooidal opesia that is larger than the orifice, a simple avicularian opesia without any indication of mandibular pivots, and generally no ovicells. Family Calescharidae Cook & Bock, 2001 Type genus: Caleschara MacGillivray, 1880. Type species: Eschara denticulata MacGillivray, 1869. Other genera: Tretosina† Canu & Bassler, 1927. Diagnosis: Colonies encrusting, unilamellar, multiserial; or erect from an encrusting base, frondose, bilamellar or quadrilateral, bifurcating, sometimes anastomosing. Autozooids longitudinally pyriform to hexagonal. Gymnocyst absent. Cryptocyst forming a proximal shelf extending over most of the frontal area, granular, encircling opesia, developing a median process or denticle. Opesia usually coincident with the orifice, transversely elliptical to semicircular, usually with indentations in the proximolateral corners. Opesiules usually present, elongate, a single pair, sometimes with additional pair of minute, circular opesiules. Septula in vertical walls, uniporous. Embryos incubated in endozooidal ovicells. Ancestrula autozooidal. Spines and avicularia absent. Distribution: Tropical waters. Remarks: Caleschara and Tretosina have been separated from Microporidae by Cook & Bock (2001) based on the unusually large endozooidal ovicells with its brood cavity protruding into the distal zooid beneath its



cryptocyst acting as endooecium. This autozooid is therefore slightly modified. Family Calpensiidae Canu & Bassler, 1923 Type genus: Calpensia Jullien, 1888. Type species: Membranipora calpensis Busk, 1854 (= Cellepora nobilis Esper, 1792). Other genera: None. Diagnosis: Colonies encrusting, unilamellar to multilamellar, multiserial. Autozooids longitudinally rectangular to hexagonal. Gymnocyst negligible to absent, smooth. Cryptocyst forming a proximal shelf extending over most of the frontal area, granular, porous, depressed. Opesiules tubular, as a single pair, proximolateral to the opesia. Lateral walls may bear tubercles. Septula in vertical walls, multiporous. No ovicells, embryos supposedly incubated internally in brood sacs. Ancestrula autozooidal. Avicularia and spines absent. Distribution: Temperate to tropical waters of the Atlantic Ocean, the Mediterranean Sea and the Torres Strait. Remarks: This little-known family comprises only two species, Calpensia nobilis from European waters and Calpensia pulchra Harmer, 1926 from the Torres Strait between Australia and New Guinea. Comparable to Alcide d’Orbigny’s “Paléontologie française” (see Martha et  al. 2019a for publication dates of the Paléontologie française), Esper’s “Pflanzenthiere in Abbildungen nach der Natur” and “Fortsetzungen der Pflanzenthiere in Abbildungen nach der Natur” have likewise been published in several releases of text pages accompanied by a number of plates. The correct publication dates of these releases have been collated in Ott (1995). Calpensia nobilis was first described as Celle­ pora nobilis on p. 145 of the fifth release of the “Fortsetzu­ ngen der Pflanzenthiere in Abbildungen nach der Natur,” which appeared in 1796 (Esper 1796), and this is the usual publication date attributed to Cellepora nobilis (e.g. Cook et al. 2018). Nonetheless, the species was already depicted and named on a plate accompanying the tenth release of the “Pflanzenthiere in Abbildungen nach der Natur,” which appeared in 1792 (Esper 1792) (see Ott 1995, p. 12f), Celle­ pora nobilis having therefore already been available in 1792 according to Article 12.2.7 of the International Code on Zoological Nomenclature (ICZN 1999). We therefore advise that “Calpensia nobilis (Esper, 1792)” is the correct citation. Family Chlidoniidae Busk, 1884 Type genus: Chlidonia Lamouroux, 1824. Type species: Cellaria pyriformis Bertoloni, 1810. Other genera: None. Diagnosis: Colonies erect from a stolonate network of kenozooids; erect branch axes comprise jointed uniserial stems

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 353

of kenozooids, bifurcating distally in secondary kenozooids or autozooids arranged in regular serial series, with terminal spiniform kenozooids. Autozooids longitudinally pyriform. Gymnocyst extensive, smooth, forming long cauda proximally. Cryptocyst forming a proximal shelf extending over most of the frontal area, granular, depressed, encircling opesia. Opesia semicircular. Opesiules may be present, circular, as a single pair. Septula basal, uniporous. Embryos incubated in internal brood sacs of polymorphic female zooids. Ancestrula not reported. Avicularia may be present, vicarious. Spines absent (Fig. 11.22 A, B). Distribution: Tropical to temperate waters. Remarks: Chlidoniids have erect, nodal, uni- to biserial colonies that form from a stolonate network of kenozooids, not dissimilar to alysidiids. Chlidonia pyriformis is a well-known species reported from tropical to temperate waters around the world. The only other putative representative of this family was described as Cothurnicella japonica Mawatari, 1964 from the Sea of Japan; it requires restudy. The family previously also included Crepis Jullien, 1882a, which was transferred to the Calloporidae by Reverter-Gil et al. (2011) and the Cymuloporidae by Winston and Vieira (2013). Family Onychocellidae† Jullien, 1882b Type genus: Onychocella† Jullien, 1882b. Type species: Cellepora angulosa† Reuss, 1848. Other genera: Chondriovelum Hayward & Thorpe, 1987, Floridina Jullien, 1882b, Floridinella† Canu & Bassler, 1917, Rectonychocella Canu & Bassler, 1917, Smittipora Jullien, 1882b. Diagnosis: Colonies encrusting, unilamellar to multilamellar, multiserial; or erect, frondose, bilamellar or vincularian. Autozooids longitudinally pyriform to hexagonal. Gymnocyst usually absent or very narrow and smooth. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, smooth to granular, encircling opesia. Opesia ovoidal, to transversely semicircular/D-shaped or trifoliate, sometimes with indentations in the proximolateral corners and/or opesiular constrictions. Opesiules may be present, as a single pair. Articulated spines mostly lacking, if present, oral, distolateral to the orifice. Embryos incubated in endozooidal or immersed ovicells, sometimes in zooids with dimorphic opesia and strongly reduced ooecium; incubation in internal brood sac possibly present too. Ancestrula autozooidal. Avi­ cularia may be present, interzooidal or vicarious, irregularly interspersed among autozooids, often asymmetrical. Kenozooids may be present (Fig. 11.22 C, D). Distribution: Tropical to temperate waters. Remarks: Several authors have regarded Onychocella marioni Jullien, 1882b as a junior synonym of Cellepora

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 11 Gymnolaemata, Cheilostomata

Fig. 11.22: (A, B) Chlidoniidae: Chlidonia pyriformis (Bertoloni, 1810), off Seaspray, Gippsland, Victoria, Australia, unregistered. Photos: P.E. Bock (C, D) Onychocellidae: Onychocella marioni Jullien, 1882b. Marseille, France, Mediterranean Sea. Muséum National d’Histoire Naturelle, Paris, IB-2008-2686, syntype. Scale bars: A, D, 250 µm; B, 100 µm; C, 500 µm.

angulosa† Reuss, 1848 following Harmer (1926). This was, however, previously objected to by Canu and Bassler (1930a) and some subsequent authors. A comprehensive revision of the family and all its genera, fossil and Recent, has been provided by Taylor et  al. (2018). Although less common and nowadays comprising fewer species, onychocellids are one of the most widely reported groups of fossil cheilostomes, having appeared at the base of the radiation of flustrine cheilostomes (Martha & Taylor 2016). Family Poricellariidae† Harmer, 1926 Type genus: Poricellaria† d’Orbigny, 1854. Type species: Poricellaria alata† d’Orbigny, 1854. Other genera: None. Diagnosis: Colonies erect, attached to substratum by kenozooidal rhizoids, vincularian, bifurcating. Autozooids alternating, arranged in four longitudinal series,

elliptically elongate, somewhat asymmetrical. Gymnocyst extensive, smooth. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, smooth to granular, encircling opesia. Opesia usually coincident with the orifice, transversely elliptical to semicircular. Opesiules slit-like, asymmetrically placed, as single pair. Embryos incubated in immersed cleithral terminal ovicells. Avicularia adventitious, on proximal gymnocyst of each autozooid. Spines absent (Fig. 11.23 A, B). Distribution: Tropical waters of eastern Indonesia. Remarks: Harmer (1926) assigned Micropora ratoniensis Waters, 1887a (figured therein, but described in Waters, 1887b), originally from the coast of New Guinea, provisionally to Poricellaria. This is currently the only Recent representative of this monogeneric family of otherwise Palaeocene to Oligocene (or Miocene) species (see Cook et al. 2018).



11.3 Systematics of Cheilostomata 

 355

Fig. 11.23: (A, B) Poricellariidae: Poricellaria ratoniensis (Waters, 1887a). Station B 3/2, Safaga Bay, Egypt, depth 4 m, Red Sea. University of Vienna, unregistered. (C, D) Setosellidae: Setosella vulnerata (Busk, 1860a). Shetland Islands, United Kingdom, NE Atlantic Ocean. Natural History Museum, London, 1911.10.1.760, paralectotype. Scale bars: A, 250 µm; B, 50 µm; C, 500 µm; D, 100 µm.

Family Setosellidae Levinsen, 1909 Type genus: Setosella Hincks, 1877a. Type species: Membranipora vulnerata Busk, 1860a. Other genera: None. Diagnosis: Colonies encrusting, unilamellar, multiserial. Autozooids longitudinally elliptical to hexagonal. Frontal skeletal surface almost entirely a flat cryptocystal shelf, bordered by thin raised margin and narrow gymnocyst. One or two pairs of opesiules. Opesia coincident with the orifice, semicircular to transversely elliptical. Septula in vertical walls, multiporous. Embryos incubated in immersed cleithral terminal ovicells. Ancestrula autozooidal. Vibraculoid avicularia interzooidal, one distal to each autozooid, lacking pivot bar, with setiform mandible. Kenozooids may be present. Spines absent (Fig. 11.23 C, D). Distribution: Tropical to temperate waters of the North Atlantic and Mediterranean Sea.

Remarks: Comprising only six Recent species, Setosellidae is a relatively small family. Setosellids are very simple cheilostomes that are distinguished by colonies having an opesiulate zooidal cryptocyst and interzooidal vibracular polymorphs with long setae.

11.3.7.7 Superfamily Monoporelloidea Hincks, 1882b Family Monoporellidae Hincks, 1882b Type genus: Monoporella Hincks, 1881b. Type species: Haploporella nodulifera Hincks, 1881c. Other genera: None. Diagnosis: Colonies encrusting, unilamellar, multiserial; or erect, bilamellar, multiserial, bifurcating. Autozooids longitudinally elliptical to hexagonal. Gymnocyst absent or negligible, smooth. Cryptocyst forming a proximal shelf

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 11 Gymnolaemata, Cheilostomata

Fig. 11.24: (A, B) Monoporellidae: Monoporella sp. Vabbinfaru Isl., North Male Atoll, Maldive Islands, Indian Ocean. University of Vienna, unregistered. (C, D) Adeonidae: Reptadeonella bipartita (Canu & Bassler, 1928a). Bahia, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade Federal da Bahia, 0266. Scale bars: A, 1 mm; B, D, 250 µm; C, 500 µm.

extending over half or more of the frontal area, porous or granular, encircling opesia. Opesia coincident with the orifice, semicircular to transversely elliptical. Opesiules usually present, circular, a single pair. Spines oral, distolateral to the orifice. Pore-chambers basal. Several embryos incubated simultaneously in subimmersed or endozooidal cleithral ovicells with ooecium constructed from modified spines and with two lateral foramina leading to brood cavity. Ancestrula autozooidal. Kenozooids may be present. Avicularia absent (Fig. 11.24 A, B). Distribution: Tropical to polar waters predominantly of the Northern Hemisphere. Remarks: Both the genus and the type species were described word for word in Hincks (1881b) and Hincks (1881c) using the generic name Monoporella in the first account and Haploporella in the second. Although Hincks (1881c) appeared 1 month earlier, thus giving priority to the generic name Haploporella, Hincks (1881d, p. 135) clarified

his revocation of Haploporella as a junior homonym of a foraminiferan genus. Monoporellids have characteristic ovicells with ooecia constructed from spines, an arrangement considered to be ancestral (Ostrovsky & Taylor 2004, 2005). Family Macroporidae† Uttley, 1949 Type genus: Macropora† MacGillivray, 1895. Type species: Macropora centralis† MacGillivray, 1895. Other genera: None. Diagnosis: Colonies encrusting, unilamellar, unimultiserial; or erect, bilamellar, multiserial. Autozooids longitudinally elliptical to hexagonal. Gymnocyst absent or negligible, smooth. Cryptocyst forming a proximal shelf extending over half or more of the frontal area, granular, pierced by numerous opesiules, encircling opesia. Opesia coincident with the orifice, semicircular; opercula composed of calcified layer and membranous layer. Opesiules



circular, up to 60 per autozooid. Spines oral, distolateral to the orifice. Pore-chambers basal. Several embryos incubated simultaneously in hyperstomial cleithral ovicells with ooecium constructed from modified spines. Ancestrula autozooidal. Avicularium-like B-zooids in most species, vicarious, having orifice with crenulated distal rim, calcified opercula and oral spines. Kenozooids usually absent. Distribution: Tropical to temperate waters predominantly of the Southern Hemisphere. Remarks: Macroporidae comprises only one genus that has, however, many different extant and extinct species, the oldest dating back to the Eocene (Hara 2002). The type species, Macropora centralis, is from the Miocene of Victoria, Australia. Macroporidae is a very distinctive family, characterized by calcified opercula and having large, avicularium-like polymorphs with a distally crenulated orifice, spines and calcified opercula as well. The cryptocystal frontal wall in macroporids is pierced by numerous opesiules for the passage of parietal muscles (see Gordon 1984, Banta et al. 1997) and the hyperstomial ovicell is very large.

11.3.7.8 Superfamily Adeonoidea Busk, 1884 Family Adeonidae Busk, 1884 Type genus: Adeona Lamouroux, 1812. Type species: Adeona grisea Lamouroux, 1812. Other genera: Adeonella Busk, 1884, Adeonellopsis MacGillivray, 1886, Bracebridgia MacGillivray, 1886, Dimorphocella† Maplestone, 1903a, Kubaninella† Grischenko & Mawatari, 2002, Laminopora Michelin, 1842, Reptadeonella Busk, 1884. Diagnosis: Colonies encrusting or erect, uni- to multilamellar, usually multiserial, rarely uni- to oligoserial. Colony usually pigmented, yellowish, pinkish, dark red to black in color. Autozooids longitudinally rectangular to hexagonal. Frontal shield umbonuloid with frontal spiramen, or lepralioid with peristomial pseudospiramen. Primary orifice mainly sunken in the frontal shield; secondary orifice circular to transversely oblong. Oral spines absent. Condyles may be present. Avicularia variable, adventitious, interzooidal or vicarious; the latter mainly present along or near branch margins and bifurcations in erect colonies. Embryos incubated in internal brood sac of female zooids (gonozooids) that may be morphologically different or not from autozooids. Ancestrula autozooidal. Kenozooids may be present (Fig. 11.24 C, D). Distribution: Tropical to temperate waters. Remarks: Gregory (1893) previously recognized two distinct families, Adeonidae and Adeonellidae, and Cook (1973) maintained this distinction, while noting the many

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 357

obvious characters they had in common. Some genera previously assigned to Adeonidae have lepralioid frontal shields, whereas some members of Adeonellidae have an umbonuloid frontal shield. Phylogenetic relationships based on genome-skimming data sets (Orr et  al. 2019a) suggest that Adeonidae sensu lato (including taxa with either umbonuloid or lepralioid shields) is monophyletic. Vestigial ooecia were recently recorded in enlarged gonozooids in an undescribed species of Adeonellopsis. Family Inversiulidae Vigneaux, 1949 Type genus: Inversiula Jullien, 1888. Type species: Inversiula nutrix Jullien, 1888. Other genera: None. Diagnosis: Colonies encrusting, uni- to multilamellar, usually multiserial. Autozooids longitudinally oval to subhexagonal. Frontal shield umbonuloid or lepralioid, evenly perforated and tuberculate. Ascopore present, suboral. Primary orifice mainly sunken in the frontal shield, inversely D-shaped, with zooidal operculum hinged along its distal edge and opening in the reverse. Oral spines absent. Avicularia present, adventitious, lateral-oral and paired. Embryos incubated in internal brood sacs in ordinary or enlarged female zooids with dimorphic orifice. Ancestrula autozooidal. Vertical walls with basal pore-chambers. Distribution: Temperate to polar waters of the South Atlantic and South Pacific oceans. Remarks: Inversiulidae is a monogeneric family comprising five species with Southern Hemisphere distribution. The reversed orifice and distally hinged operculum, described for Inversiula inversa (Waters, 1887b), is considered unique among cheilostome bryozoans.

11.3.7.9 Superfamily Arachnopusioidea Jullien, 1888 Family Arachnopusiidae Jullien, 1888 Type genus: Arachnopusia Jullien, 1888. Type species: Lepralia monoceros Busk, 1854. Other genera: Brendella Gordon, 1989a, Briarachnia Gordon, 1984, Poricella† Canu, 1904a, Trilaminopora Moyano G., 1970b. Diagnosis: Colonies encrusting, uni- to multilamellar, multiserial; or erect, plate-like or foliaceous, uni- or bilamellar, multiserial. Autozooids sometimes with inconspicuous boundaries between autozooids. Frontal shield umbonuloid, foraminate, the number of foramina highly variable, lacking in one species. Secondary orifice variable. Oral spines ephemeral or persistent. Avicularia present, adventitious, vicarious or interzooidal. Embryos incubated in hyperstomial or subimmersed acleithral ovicells, sometimes reminiscent of endozooidal ovicell

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Fig. 11.25: (A, B) Arachnopusiidae: Poricella frigorosa Winston, Vieira, & Woollacott, 2014. Rio de Janeiro, Brazil, W Atlantic Ocean. Museum of Comparative Zoology Cambridge, USA, 137446, holotype. (C, D) Exechonellidae: Exechonella vieirai Cáceres-Chamizo, Sanner, Tilbrook, & Ostrovsky, 2017. Praia do Francês, Alagoas, Brazil, W Atlantic Ocean. Department of Palaeontology, Geozentrum, University of Vienna, 20160001-0001, holotype. Scale bars: A, C, 500 µm; B, D, 250 µm.

because of ooecium being embedded to the frontal shield of distal zooid. Ancestrula autozooidal. Vertical walls with basal pore-chambers or mural septula (Fig. 11. 25 A, B). Distribution: Cosmopolitan (predominantly southern hemisphere). Remarks: The varied mode of development of the frontal shield among the various included taxa has suggested affinities with anascan-grade, cribrimorph and umbonuloid cheilostomes. These differences have been used to highlight the inadequacies of higher classification within the Cheilostomata (Moyano G. 1970c, Hayward & Thorpe 1988a, Gordon 1989a, 2000, Cáceres-Chamizo et al. 2017). Family Exechonellidae† Harmer, 1957 Type genus: Exechonella† Canu & Bassler in Duvergier, 1924. Type species: Cyclicopora (?) grandis† Duvergier, 1920.

Other genera: Anarthropora Smitt, 1868b, Anexechona Osburn, 1950, Stephanopora Kirkpatrick, 1888, Triporula Canu & Bassler, 1927, Xynexecha Gordon & d’Hondt, 1997. Diagnosis: Colonies encrusting, uni- to multilamellar, multiserial; or erect, plate-like or foliaceous, branching. Autozooids oval or hexagonal. Frontal shield umbonuloid, smooth or pustulose, with variable number of foramina, open or covered with an external cuticular layer; marginal pores present. Primary orifice subcircular or oval. Peristome low to long and cylindrical, obscuring the primary orifice. Spines absent. Avicularia present, adventitious, sometimes associated with lateral foramina. Kenozooids sometimes present, adventitious, associated with avicularia. Embryos incubated in internal brood sacs. Ancestrula autozooidal. Vertical walls with mural septula (Fig. 11.25 C, D). Distribution: Tropical and temperate waters.



Remarks: The genus Coleopora Canu & Bassler, 1927 was considered a junior synonym of Exechonella Canu & Bassler in Duvergier, 1924, thus species previously assigned to Coleopora still require review and reassignment (Cáceres-Chamizo et  al. 2017). It is not certain if Triporula and Anarthropoma belong in this family. All exechonellids are internal brooders.

11.3.7.10 Superfamily Bifaxarioidea Busk, 1884 Family Bifaxariidae Busk, 1884 Type genus: Bifaxaria Busk, 1884. Type species: Bifaxaria submucronata Busk, 1884. Other genera: Aberrodomus Gordon, 1988, Diplonotos Canu & Bassler, 1930b, Domosclerus Gordon, 1988, Raxifabia Gordon, 1988.

11.3 Systematics of Cheilostomata 

 359

Diagnosis: Colonies erect, branching, basally rooted, narrowly biserial, with zooids arranged back to back along the stem. Frontal shield with outer porous umbonuloid layer, overlying the inner costal shield (this sometimes reduced or lacking). Primary orifice generally weakly defined, often obscured by peristome. Spines absent. Avicularia adventitious, often lateral-oral, one on either side of the orifice. Kenozooids absent. Embryos incubated in hyperstomial, subimmersed or endozooidal acleithral ovicells opening into peristome; female zooids dimorphic. Ancestrula not reported (Fig. 11.26 A, B). Distribution: Tropical and temperate deep waters predominantly of the Southern Hemisphere, rare in polar waters. Remarks: Gordon (1988) provided a detailed account of bifaxariid genera, including the differences between this family and other taxa with erect colonies and alternating

Fig. 11.26: (A, B) Bifaxariidae: Bifaxaria submucronata (Busk, 1884). RV Sonne cruise 205, Station 61, 13.174° N–3.194° N, 118.106° W–118.0953° W, Equatorial East Pacific, 3996–4007 m depth. Senckenberg Forschungsinstitute und Naturmuseen, Sektion Marine Evertebraten III (Bryozoologie), SMF 15003. Modified after Matsuyama et al. (2014). (C, D) Catenicellidae: Catenicella uberrima (Harmer, 1957). São Paulo, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, unregistered. Scale bars: A, C, 500 µm; B, 100 µm; D, 250 µm.

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 11 Gymnolaemata, Cheilostomata

zooids as some Sclerodomidae Levinsen, 1909 and Urceoliporidae Bassler, 1936. The most important external characteristic diagnostic of most Bifaxariidae is the division of the umbonuloid shield into two halves with a median suture. Family Mixtopeltidae Gordon, 1994a Type genus: Mixtopelta Gordon, 1994a. Type species: Mixtopelta indica Gordon, 1994a. Other genera: None. Diagnosis: Colonies conical, basally rooted. Frontal shield mixed: imperforate lepralioid cryptocyst proximally, with no areolar pores; suboral costal shield (spinocyst) distally, arising from a narrow gymnocyst; each costa with a large pelma. Oral spines, avicularia, and kenozooids absent. Ovicell and ancestrula not reported. Distribution: Deep sea of the Mozambique Plateau. Remarks: The conical rooted colonies and zooids with a mixed frontal shield (lepralioid proximal and costal distally) are highly diagnostic. There is no information on reproduction in Mixtopelta.

11.3.7.11 Superfamily Catenicelloidea Busk, 1852b Family Catenicellidae Busk, 1852b Type genus: Catenicella de Blainville, 1830. Type species: Catenicella savignyi de Blainville, 1830 (= Eucratea contei Audouin, 1826). Other genera: Bryosartor Gordon & Braga, 1994, Calpidium Busk, 1852b, Claviporella MacGillivray, 1887b, Cornuticella Canu & Bassler, 1927, Cornuticellina Stach, 1935a, Costati­ cella Maplestone, 1899, Cribricellina Canu & Bassler, 1927, Orthoscuticella Wass & Yoo, 1976, Paracribricellina Wass & Yoo, 1976, Plagiopora† MacGillivray, 1895, Pterocella Levinsen, 1909, Scalicella Harmer, 1957, Scuticella Levinsen, 1909, Strongylopora Maplestone, 1899, Strophipora MacGillivray, 1895, Talivittaticella Gordon & d’Hondt, 1985, Termi­ nocella Harmer, 1957, Vasignyella Gordon, 1989b. Diagnosis: Colonies erect, flexible and jointed, with each segment (internode) formed by one to three (rarely more) autozooids. Frontal skeletal wall fundamentally gymnocystal, but variably modified – with small gymnocystal pores, larger fenestrae, costae (well-developed or vestigial), or lateral pore-chambers (vittae) hugely expanded frontally in a few taxa to form a pair of cryptocystal sectors. Avicularia adventitious, single or paired at distolateral corners of zooids, sometimes dimorphic. Distal orifice transversely D-shaped or subcircular, less often distinct sinus or sinuate proximal margin; paired condyles generally present. Oral spines absent. Rhizoids may emerge from basal or lateral zooid walls. Embryos incubated in hyperstomial, subimmersed or immersed (often

terminal) or endozooidal cleithral ovicells; ectooecium partially or fully uncalcified, endooecium smooth or reticulated; maternal zooid sometimes with dimorphic orifice. Larva non-feeding (Fig. 11.26 C, D). Distribution: Tropical to temperate waters of the Southern Hemisphere. Remarks: Accounts of diversity and nomenclature used in this family have been provided by Gordon (1984) and Cook et al. (2018). This family comprises only erect taxa; thus, the relationship between other encrusting Catenicelloidea remains unknown. Ostrovsky (2013) has described the ovicells and/or reproduction in several species. Family Eurystomellidae Levinsen, 1909 Type genus: Eurystomella Levinsen, 1909. Type species: Lepralia foraminigera Hincks, 1883. Other genera: Integripelta Gordon, Mawatari & Kajihara, 2002, Selenariopsis Maplestone, 1913, Zygopalme Gordon, Mawatari & Kajihara, 2002. Diagnosis: Colonies encrusting, unilaminar, bi- to multiserial, or free-living, discoidal, anchored by basal rhizoids. Autozooids arranged in a regular series. Frontal shield comprising extensive smooth gymnocyst, perforated by large foramina or imperforate, with vestigial suboral costal field or this absent; proximal umbo sometimes present. Oral spines and avicularia absent. Embryos incubated in immersed or endozooidal cleithral ovicells; ooecium with one or more frontal fenestrae; maternal zooids dimorphic, with slightly broader orifice. Ancestrula with membranous frontal wall, without spines. Interzooidal communication via pore-chambers or mural septula (Fig. 11.27 A, B). Distribution: Tropical to temperate waters. Remarks: Revisions of taxa assigned to Eurystomellidae have been provided by Cook and Chimonides (1981a) and Gordon et al. (2002), suggesting eurystomellids could have evolved from a cribrimorph ancestor by an increase in the gymnocystal frontal shield and decrease in costal field. Australiana (Powell, 1966) is a junior synonym of Selenari­ opsis; if this distinctive genus with discoidal colonies were to be accorded its own subfamily, the subfamily name would be Australianinae Powell, 1966 (ICZN Article 40.1). Family Petalostegidae Gordon, 1984 Type genus: Petalostegus Levinsen, 1909. Type species: Catenaria bicornis Busk, 1884. Other genera: Chelidozoum Stach, 1935b. Diagnosis: Colonies erect, delicate, jointed, uniserial and branching, anchored by rhizoids. Autozooids elongate, with long tubular portion. Frontal shield formed by small to extensive flattened costal field, with pores between costae. Orifice semicircular, with distolateral



11.3 Systematics of Cheilostomata 

 361

Fig. 11.27: (A, B) Eurystomellidae: Integripelta novella Gordon, Mawatari & Kajihara, 2002. Akkeshi Bay, Hokkaido, Japan, NW Pacific, intertidal. Natural History Museum, London, 2006.2.27.53. (C, D) Savignyellidae: Savignyella lafontii (Audouin, 1826). Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, unregistered. Scale bars: A, C, 500 µm; B, 250 µm; D, 100 µm.

acute projections or with sessile avicularia. Avicularia present, adventitious, sessile, at distolateral corners of the zooid or associated with kenozooids. Kenozooids may be present, elongate and spine-like to club shaped, terminal or between internodes. Embryos incubated in hyperstomial terminal ovicells, with dimorphic maternal zooid. Distribution: Tropical to temperate waters of the Southern Hemisphere. Remarks: Petalostegidae was originally included in Buguloidea but was reassigned to the Catenicelloidea by Gordon and d’Hondt (1991), who provided an expanded morphological characterization of the family. Branch axes are composed of autozooids in Petalostegus whereas in Chelidozoum they are composed of kenozooids in all but one species. Family Savignyellidae Levinsen, 1909 Type genus: Savignyella Levinsen, 1909. Type species: Eucratea lafontii Audouin, 1826.

Other genera: Halysisis Norman, 1909. Diagnosis: Colonies erect, flexible and jointed, bifurcating, with each segment (internode) formed by one autozooid. Autozooids elongate, almost infundibuliform. Frontal skeletal wall mostly pseudoporous (extensive in Savignyella) and interior-walled (cryptocystal) with a closely applied frontal membrane; frontal pseudoporous area in Halysisis separated by narrow gymnocystal strips from wide lateral sectors, each with uniserial row of pores; caudal part of frontal wall gymnocystal. Avicularia present or absent, adventitious, single, suboral. Orifice subcircular, condyles tiny or absent. Oral spines present or absent. Rhizoids originate from the basal or lateral zooid wall. Embryos incubated in terminal hyperstomial or immersed acleithral ovicells; ectooecium in hyperstomial ovicell almost entirely membranous. Larva non-feeding. Vertical distal wall with mural septula (Fig. 11.27 C, D). Distribution: Tropical to temperate waters.

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 11 Gymnolaemata, Cheilostomata

Remarks: Savignyellidae comprises only two monospecific genera with distinct characteristics: Savignyella has avicularia, non-articulated oral spines, and its ovicells are immersed, with well-developed ooecium. Halysisis has no avicularia and oral spines, and its ovicells are immersed, with small ooecium (Waters 1913). The lateral interior-walled sectors in Halysisis appear to be broader versions of the narrow vittae (analogous to pore-chambers) seen in many catenicellids.

11.3.7.12 Superfamily Celleporoidea Johnston, 1838 Family Celleporidae Johnston, 1838 Type genus: Cellepora Linnaeus, 1767. Type species: Millepora pumicosa Pallas, 1766.

Other genera: Buffonellaria Canu & Bassler, 1927, Buskea Heller, 1867, Calvipelta Tilbrook, 2006, Celleporina Gray, 1848, Favosthimosia Hayward & Winston, 2011, Galeop­ sis Jullien in Jullien & Calvet, 1903, Lagenipora Hincks, 1877b, Lageniporina Winston, 2016, Omalosecosa Canu & Bassler, 1925a, Omanipora Berning & Ostrovsky, 2011, Orthoporidroides Moyano G., 1974, Osthimosia Jullien, 1888, Palmicellaria Alder, 1864, Pourtalesella Winston, 2005, Pseudocelleporina Mawatari, 1986, Ramicellepora Gordon, 2014, Richbunea Gordon & d’Hondt, 1997, Sin­ uporina Pouyet, 1973, Spigaleos Hayward, 1992, Tegminula Jullien, 1882a, Torquatella Tilbrook, Hayward & Gordon, 2001, Turbicellepora Ryland, 1963. Diagnosis: Colony encrusting, uni- to multilaminar, multiserial, nodular to massive; or erect, erect, arising from encrusting base, dichotomously branching. Autozooids

Fig. 11.28: Celleporidae. (A, B) Cellepora nodulosa von Lorenz, 1886. Barents Sea. Perm State National Research University, Kluge Collection, 810.16. (C, D) (former Torquatellidae) Torquatella duolamellata (Scholz, 1991). Okinawa, Japan, East China Sea, intertidal. National Museum of Nature and Science, Tsukuba, NSMT-Te 1172. Scale bars: A, C, 500 µm; B, D, 250 µm; d, 100 µm.



arranged in a regular series or chaotic in the colony. Frontal shield lepralioid, mostly imperforate, pseudoporous in a few taxa, thickened by increasing secondary calcification. Primary orifice semicircular to rounded, often with proximal sinus or broad poster. Condyles present or absent. Spines absent. Peristome well developed, raised, sometimes forming pseudospiramen or pseudosinus. Avicularia typically present; adventitious avicularia often oral, single or paired (rarely multiple); vicarious avicularia irregularly interspersed among autozooids. Embryos incubated in hyperstomial or subimmersed ovicells, mostly terminal, subcleithral and non-cleithral. Larva non-feeding. Ancestrula autozooidal (Fig. 11.28). Distribution: Cosmopolitan. Remarks: Lamouroux’s “Celléporées” (1821, p. 2) was used as a taxonomic name to group genera, but not on a family level. Thus, it does not fulfill the provisions of Article 11.7 of the International Code on Zoological Nomen­ clature (ICZN 1999), and Johnston (1838) is recognized as author of Celleporidae. Yang et  al. (2018a) placed Torquatella in Celleporidae, making Torquatellidae Tilbrook, 2006 a junior synonym of Celleporidae. Family Colatooeciidae Winston, 2005 Type genus: Colatooecia Winston, 2005. Type species: Porina serrulata Smitt, 1873. Other genera: Cigclisula Canu & Bassler 1927, Trematooe­ cia Osburn, 1940. Diagnosis: Colony encrusting, uni- to multilaminar, multiserial; or erect, bilaminar, multiserial, branched. Autozooids arranged in regular rows or chaotic in the colony. Frontal shield lepralioid, perforated by pseudopores, thickened by increasing secondary calcification as colonies age. Spiramen may be present, conspicuous in developing zooids, joining with the peristome and immersed in the frontal shield in completely calcified zooids. Primary orifice semicircular to rounded, deeply surrounded by zooidal calcification; secondary orifice level with frontal-shield surface, with or without tubercles. Condyles present or absent. Spines absent. Interzooidal and adventitious avicularia sometimes present. Ovicells hyperstomial, subcleithral, ooecium becoming embedded in secondary calcification leaving single central membranous area (in mostly calcified ectooecium) or longitudinal band of pits (supposedly corresponding to pseudopores). Ancestrula autozooidal (Fig. 11.29 A, B). Distribution: Tropical to temperate waters. Remarks: This family was erected by Winston (2005) to accommodate the single genus Colatooecia. Cigclisula and Trematooecia have previously been included in families such as Celleporidae Johnston, 1838, Hippoporidridae

11.3 Systematics of Cheilostomata 

 363

Vigneaux, 1949, and Stomatochetosellidae Canu & Bassler, 1917, based on colony form and autozooidal characters. Vieira et  al. (2010) used ovicellar characters to justify transfer of these genera into Colatooeciidae. A revision of Colatooeciidae was provided by Almeida et al. (2014). Family Hippoporidridae Vigneaux, 1949 Type genus: Hippoporidra Canu & Bassler, 1927. Type species: Cellepora edax Busk, 1859b. Other genera: Abditoporella Sosa-Yañez, Vieira & SolísMarín, 2015, Hippoporella Canu, 1917, Hippotrema Canu & Bassler, 1927, Odontoporella Héjjas, 1894, Scorpiodinipora Balavoine, 1959. Diagnosis: Colony encrusting, uni- to multilaminar, multiserial. Autozooids hexagonal, arranged in regular or irregular series. Frontal shield lepralioid, with or without pseudopores; areolar pores present. Primary orifice semicircular to bell shaped, deeply surrounded by zooidal calcification; secondary orifice level with frontal-shield surface, with or without tubercles. Condyles present. Spines absent. Kenozooids, interzooidal and adventitious avicularia sometimes present. Embryos incubated in internal brood sacs or in hyperstomial or subimmersed acleithral ovicells with ooecium becoming embedded in secondary calcification; ectooecium often with small membranous frontal area. Zooidal sexual dimorphism present (external or in polypide only). Ancestrula autozooidal or with extensive membranous frontal wall and no marginal spines (Fig. 11.29 C, D). Distribution: Cosmopolitan. Remarks: Hippoporidra had been included in Cleidochasmatidae Cheetham & Sandberg, 1964. Based on similarities in colony morphology between Hippoporidra and celleporids, Pouyet (1973) raised the subfamily Hippoporidrinae Vigneaux, 1949 to family rank. Hippoporidrid colonies have been frequently found encrusting gastropod shells inhabited by pagurids and dead sea-urchin tests. Family Phidoloporidae† Gabb & Horn, 1862 Type genus: Phidolopora† Gabb & Horn, 1862. Type species: Phidolopora labiata† Gabb & Horn, 1862. Other genera: Bryorachis Gordon & Arnold, 1998, Chevron Gordon, 1989c, Crenulatella Sokolover, Taylor & Ilan, 2016, Dentiporella Barroso, 1926, Dictyochasma Hayward, 1999, Fodinella Tilbrook, Hayward & Gordon, 2001, Hippello­ zoon Canu & Bassler, 1917, Iodictyum Harmer, 1933, Mal­ leatia Jullien in Jullien & Calvet, 1903, Metacleidochasma Soule, Soule & Chaney, 1991b, Plesiocleidochasma Soule, Soule & Chaney, 1991b, Pleuromucrum† Vigneaux, 1949,

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Fig. 11.29: (A, B) Colatooeciidae: Colatooecia serrulata (Smitt, 1873). Caribbean Sea, unregistered. Photos: J. E. Winston. (C, D) Hippoporidridae: Abditoporella dimorpha Sosa-Yañez, Vieira & Solís-Marín, 2015. Mexico, E Pacific Ocean. Instituto de Ciencias del Mar y Limnología, Universidad Nacional Autónoma de (see Sosa-Yañez et al. 2015, fig. 1B, D). Scale bars: A, 500 µm; B, D, 100 µm; C, 250 µm.

Psammocleidochasma Winston & Vieira, 2013, Psileschara Busk, 1861b, Reteporella Busk, 1884, Reteporellina Harmer, 1933, Rhynchozoon Hincks, 1895, Schizoretepora Gregory, 1893, Schizotheca Hincks, 1877a, Sparsiporina† d’Orbigny, 1852, Stephanollona Duvergier, 1920, Triphyllozoon Canu & Bassler, 1917. Diagnosis: Colony encrusting, or erect and dichotomously branching or reticulate, forming a lace-like mesh with zooids opening on one side of the branches. Autozooidal frontal shield typically smooth and porcellanous, although it may sometimes be mamillated, dimpled or otherwise textured; with a few small marginal areolarseptular pores. Primary orifice generally with toothed or beaded distal rim. Proximal border straight, concave or distinctly sinuate with condyles present. When oral spines present, they may have a series of nested joints. Avicularia

generally present and polymorphic, being adventitious and/or vicarious. Embryos incubated in hyperstomial non-cleithral ovicells with ooecium often becoming immersed in secondary calcification. Ectooecium may have a membranous area as an arch or narrow fissure or a broad window, oval or triangular. Larva non-feeding (Fig. 11.30 A, B). Distribution: Cosmopolitan. Remarks: Phidoloporidae is a very speciose bryozoan family. Some 25 genera, three of them extinct, are currently recognized as belonging to this family, comprising some 370 living species (Cook et al. 2018). Authors historically had used the names Reteporidae and Sertellidae to refer to Phidoloporidae; nevertheless, Gordon (1989c) showed that the family taxon Phidoloporidae should take precedence over the other family names.



11.3 Systematics of Cheilostomata 

 365

Fig. 11.30: (A, B) Phidoloporidae: Phidolopora elongata (Smitt, 1868b). Akkeshi Bay, Hokkaido, Japan, NW Pacific, intertidal. Natural History Museum, London, 2006.2.27.107. (C, D) Conescharellinidae: Conescharellina cookae Vieira, Gordon, Souza & Haddad, 2010. São Paulo, Brazil, REVIZEE station 6693, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, 0430–453, paratype. Scale bars: A, 500 µm; B, D, 250 µm; C, 1 mm.

11.3.7.13 Superfamily Chlidoniopsoidea Harmer, 1957 Family Chlidoniopsidae Harmer, 1957 Type genus: Chlidoniopsis Harmer, 1957. Type species: Chlidoniopsis inflata Harmer, 1957. Other genera: None. Diagnosis: Colony erect, uniserial, jointed, with internodes comprising one to three autozooids. Autozooid with umbonuloid frontal shield, smooth, imperforate, except for the presence of one or more rows of marginal septular pores. Autozooids narrower proximally, to claviform with long tubular cauda. Peristomial orifice terminal, primary orifice concealed. Oral spines and avicularia absent. Embryos incubated in hyperstomial terminal ovicells opening into peristome.

Distribution: Halmahera Sea. Remarks: The family, revised in Zágoršek et  al. (2015), contains only one Recent representative, Chlidoniopsis inflata, endemic to the Halmahera Sea.

11.3.7.14  Superfamily Conescharellinoidea Levinsen, 1909 Family Conescharellinidae Levinsen, 1909 Type genus: Conescharellina d’Orbigny, 1852. Type species: Conescharellina angustata d’Orbigny, 1852. Other genera: Bipora Whitelegge, 1887, Crucescharellina Silén, 1947, Flabellopora d’Orbigny, 1851, Ptoboroa Gordon & d’Hondt, 1997, Trochosodon Canu & Bassler, 1927, Zeug­ lopora Maplestone, 1909a.

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 11 Gymnolaemata, Cheilostomata

Diagnosis: Colony free-living, conical to discoidal, rarely suberect or branching, attached to small particles by cuticular roots from kenozooidal pores. Autozooids deep-bodied with reversed frontal budding. Frontal shield fundamentally lepralioid, concealed in most zooids by the next generation of zooids and with only a small part showing around the orifice, but fully seen antapically if radially arranged frontal shields form much of the surface, otherwise antapical surface covered by small kenozooids (cancelli) of central core, often accompanied by avicularia. Primary orifice subcircular, often with sinus and paired condyles. Oral spines and lyrula absent. Avicularia present or absent, adventitious or interzooidal, sometimes regularly dispersed in the colony. Antapical surface of colony may comprise kenozooids. Embryos incubated in hyperstomial non-cleithral ovicells with partially membranous ectooecium. Ancestrular area adapical, often with root pores and avicularia (Fig. 11.30 C, D). Distribution: Tropical to temperate waters. Remarks: Although widely distributed in tropical and temperate regions around the world, the family is particularly diverse in the Indo-West Pacific and Australia. The genera assigned to Conescharellinidae are mainly distinguished by orientation of the zooids in the colony and colony shape (Cook et al. 2018). The orientation of the primary orifice is with the apparent distal border directed toward to the apical ancestrular region. The family was revised by Bock and Cook (2004), who suggested that reversed frontal budding in Batoporidae Neviani, 1901, Lekythoporidae Levinsen, 1909, and Orbituliporidae Canu & Bassler, 1923 may not necessarily reflect close systematic relationships among these groups. Nevertheless, Cook et al. (2018) used reversed frontal budding to include all these families in the superfamily Conescharellinoidea Levinsen, 1909, a classification that we follow here. Ovicells are known in relatively few species. Family Batoporidae† Neviani, 1901 Type genus: Batopora† Reuss, 1867. Type species: Batopora stolickzai† Reuss, 1867. Other genera: Lacrimula Cook, 1966. Diagnosis: Colony free-living, globular, conical or stellate, with rootlets arising from an apical pit surrounded by kenozooids or from a single central kenozooid. Autozooids with reversed frontal budding. Autozooids elongate, alternate. Primary orifice subcircular. Oral spines, sinus, and lyrula absent. Avicularia present or absent, interzooidal or suboral, with a paired condyles or complete crossbar. Apical kenozooid may be associated with avicularia. Embryos incubated in peristomial ovicells. Distribution: Tropical Indo-Pacific. Remarks: Bock and Cook (2004) suggested maintaining Batoporidae for both Batopora and Lacrimula,

characterized by orificial and ovicellar characters that differ from both Conescharellinidae and Orbituliporidae Canu & Bassler, 1923. Species of Batoporidae are widely reported as fossils in Europe, while Recent species are known from the Indo-Pacific region and in deep waters of Australia. Family Lekythoporidae Levinsen, 1909 Type genus: Lekythopora MacGillivray, 1883a. Type species: Lekythopora hystrix MacGillivray, 1883a. Other genera: Aulopocella† Maplestone, 1903b, Catadysis Canu & Bassler, 1927, Harpagozoon Gordon, 2009b, Jug­ escharellina Gordon, 1989c, Orthoporidra Canu & Bassler, 1927, Poecilopora MacGillivray, 1886, Turritigera Busk, 1884. Diagnosis: Colony erect, cylindrical, unbranched to irregularly branched, sometimes laterally expansive or subglobular, arising from an encrusting base. Autozooids with reversed frontal budding. Autozooids elongate, bi- to oligoserial. Frontal shield lepralioid, smooth, with marginal septular pores. Primary orifice subcircular or with distinct sinus. Peristome often developed, sometimes with one or more avicularia. Oral spines and lyrula absent. Avicularia present or absent, adventitious or interzooidal, with complete crossbar. Embryos incubated in peristomial ovicells. Ectooecium with large membranous window. Distribution: Tropical to polar waters of the Southern Hemisphere. Remarks: Cook and Hayward (1983) and Bock and Cook (2000) provided detailed reviews of this family. Like other families assigned to Conescharelloidea, Lekythopora is characterized by having colonies with reversed frontal budding, but this placement still requires further study (Cook et al. 2018). Family Orbituliporidae† Canu & Bassler, 1923 Type genus: Orbitulipora† Stoliczka, 1862. Type species: Orbitulipora haidingeri† Stoliczka, 1862. Other genera: Sphaerulobryozoon d’Hondt, 1981. Diagnosis: Colony conical to discoidal, anchored by rhizoids from an apical or radial kenozooid. Autozooids budding from the frontal shield, radially arranged in conical colonies, with the orifice organized at periphery of the colony. Primary orifice subcircular, with or without condyles. Oral spines, sinus, and lyrula absent. Peristome may be present, elongate. Avicularia present or absent, interzooidal, with paired condyles. Embryos incubated in hyperstomial ovicells. Distribution: Tropical Atlantic Ocean. Remarks: Gordon (1989c) considered the presence of apical or radial tubes associated with the anchoring rhizoids the only distinction between orbituliporids (present) and conescharellinids (absent). Batoporidae Neviani, 1901 has been considered a senior synonym of Orbituliporidae,



but Orbituliporidae, for only Orbitulipora, was maintained as distinct family by having “discoidal” colonies (Bock & Cook 2004). The type of Orbitulipora Stoliczka, 1862 Orbitulipora haidingeri has orbicular, bilamellar, discoidal (in the vertical plane, not horizontal) colonies with a perforated peduncle (Zágoršek & Gordon 2014), distinct from globular or conical colonies found in other genera currently assigned to the Orbituliporidae. Currently, this family comprises nine genera, of which Sphaerulobryo­ zoon pedunculatum d’Hondt, 1981, type species of Sphaer­ ulobryozoon, is the only Recent representative.

11.3.7.15 Superfamily Cribrilinoidea Hincks, 1879 Family Cribrilinidae Hincks, 1879 Type genus: Cribrilina Gray, 1848.

11.3 Systematics of Cheilostomata 

 367

Type species: Lepralia punctata Hassall, 1841. Other genera: Anaskopora† Wass, 1975, Braikovia Gontar, 2012, Callistopora Winston, 2005, Cinclidia Denisenko, 2018, Collarina Jullien, 1886, Corbulipora† MacGillivray, 1895, Corbuliporina Vieira, Gordon, Souza & Haddad, 2010, Cribralaria Silén, 1941, Cribrilaria Canu & Bassler, 1929b, Dendroperistoma Moyano G., 1985, Distansescharella† d’Orbigny, 1853, Figularia Jullien, 1886, Filaguria Moyano G., 1991, Gephyrotes Norman, 1903b, Glabrilaria Bishop & Househam, 1987, Harmelinius Rosso in Rosso et  al., 2018, Hayamiellina Grischenko & Gordon in Grischenko et al., 2004, Inferusia Kukliński & Barnes, 2009, Inversis­ caphos Hayward & Cook, 1979, Jolietina Jullien, 1886, Jul­ lienula Bassler, 1953, Cribrilina (Juxtacribrilina) Yang, Seo, Min, Grischenko & Gordon, 2018b, Khulisa BoonzaaierDavids, Florence & Gibbons, 2020, Klugerella Moyano G., 1991, Membraniporella Smitt, 1873, Parafigularia Moyano G.,

Fig. 11.31: (A, B) Cribrilinidae: Cribrilina punctata (Hassall, 1841). Moray Firth, Scotland, United Kingdom, North Sea. Natural History Museum, London, 1985.11.20.1, neotype. (C, D) Didymosellidae: Didymosella pluma Cook & Chimonides, 1981b. Bahia, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade Federal da Bahia, 0710. Scale bars: A, 1 mm; B, 100 µm; C, 500 µm; D, 250 µm.

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 11 Gymnolaemata, Cheilostomata

1984, Puellina Jullien, 1886, Reginella Jullien, 1886, Reginelloides Soule, Soule & Chaney, 1995, Rosulapelta Winston & Vieira, 2013, Spiniflabellum Di Martino & Rosso, 2015, Teresaspis Rosso in Rosso et al., 2018. Diagnosis: Colonies encrusting or erect, uni- to multilamellar, usually multiserial, rarely uni- to oligoserial. Autozooids longitudinally rectangular to hexagonal or elliptical. Gymnocyst extensive or not visible frontally, surrounding the whole frontal shield or proximally only, smooth. Frontal shield spinocystal, costae varying in size, shape, number, contiguity, and porosity. Primary orifice uncalcified; secondary orifice often simulating a primary orifice, with discrete rim, operculum, and condyle-like structures. Oral spines may be present. Embryos incubated in hyperstomial, subimmersed, immersed or endozooidal ovicells, acleithral, cleithral or semicleithral. Ooecium costate, bilobate or complete. Ectooecium with pseudopores or membranous windows. Larva non-feeding. Avicularia present or absent, adventitious, interzooidal or vicarious, sometimes of more than one type. Porechambers basal. Ancestrula tatiform or costate. Kenozooids may be present (Fig. 11.31 A, B). Distribution: Cosmopolitan. Remarks: Currently, most cheilostome species having a spinocystal shield are grouped into one family, Cribrilinidae. However, the cribrilinid frontal shield appears to have evolved from calloporids with circumopesial spines and the transition from articulated periopesial spines to unjointed costae may have evolved several times in Earth’s history (see e.g. Voigt 1999, Dick et al. 2009), the family being therefore polyphyletic. Concepts presented in Lang (1916a, b, 1921, 1922) to divide cheilostome genera with a spinocystal shield into several families were done for fossil species from the Late Cretaceous and the Palaeocene. These families are not accepted nowadays pending a proper revision of all cribrimorph genera. The morphological characteristics of some Recent cribrimorphs, e.g. Figularia and Parafigularia, indicate that they could be related to other cheilostome families, such as Euthyroididae, Euthyriselloidea, Polliciporidae, and Vitrimurellidae. Family Euthyroididae Levinsen, 1909 Type genus: Euthyroides Harmer, 1902. Type species: Carbasea episcopalis Busk, 1852b. Other genera: None. Diagnosis: Colonies erect, unilamellar, multiserial, branching dichotomously, flexible, may form broad flabellate fronds, anchored by rhizoids. Autozooids weakly calcified, longer than wide. Gymnocystal frontal shield, smooth, imperforate. Small costal field present, proximal to the orifice, sometimes with pelmatidia. Primary orifice

almost semicircular, with concave proximal border. Oral spines absent. Avicularia may be present, vicarious, large. Embryos incubated in hyperstomial cleithral ovicells; ectooecium with median suture line and large paired membranous area. Maternal zooids often with slightly wider orifice and distinct costae area with pelmatidia. Kenozooids present on margins of the branches. Distribution: Temperate waters of the Tasman Sea. Remarks: This monogeneric family was introduced by Levinsen (1909) to include two species from Pacific waters. Family Polliciporidae Moyano G., 2000 Type genus: Pollicipora Moyano G., 2000. Type species: Pollicipora fucata Moyano G., 2000. Other genera: None. Diagnosis: Colonies erect, ramified, dichotomously branching, flexible, lightly calcified, comprising alternate back-to-back pairs of zooids. Autozooids elongate, slightly wider distally. Frontal shield with extensive gymnocystal part, smooth, with reduced suboral costal field. Primary orifice almost semicircular, with straight proximal border formed by fused costae. Oral spines absent. Avicularia present, adventitious, distal. Embryos incubated in hyperstomial cleithral ovicells. Kenozooids present on margins of the branch margins. Distribution: Polar Pacific Ocean off the southern coast of Tierra del Fuego. Remarks: The general morphology of the colony in Pol­ licipora resembles that of Malakosaria Goldstein, 1882, which is distinguished by having a suboral ascopore and pore-chambers around the orifice but no costal field or avicularia.

11.3.7.16 Superfamily Didymoselloidea Brown, 1952 Family Didymosellidae Brown, 1952 Type genus: Didymosella Canu & Bassler, 1917. Type species: Lepralia larvalis MacGillivray, 1869. Other genera: Tubiporella Levinsen, 1909. Diagnosis: Colonies encrusting, multiserial, or erect from an encrusting base, bilaminar. Autozooids irregularly hexagonal. Frontal shield lepralioid with frontal pseudopores and marginal septular pores. Primary orifice almost semicircular, obscured by well-developed secondary orifice with long peristome. Spiramen present, single or paired. Avicularia present, adventitious, uni- or bilateral, with complete crossbar. Embryos incubated in peristomial ovicells (Fig. 11.31 C, D). Distribution: Tropical waters. Remarks: Didymosellidae includes two genera with roughly 16 species that date from the Eocene to the Recent.



11.3.7.17 Superfamily Euthyriselloidea Bassler, 1953 Family Euthyrisellidae Bassler, 1953 Type genus: Euthyrisella Bassler, 1936. Type species: Euthyris obtecta Hincks, 1882a. Other genera: Pleurotoichus Levinsen, 1909, Tropidozoum Harmer, 1957. Diagnosis: Colonies erect, flexible and branching; or cellariiform, jointed. Interior-walled frontal shield with lepralioid ascus. Subfrontal hypostegal coelom comprising an extension of the frontal hypostegal coelom (or part of it) beneath the calcified frontal shield. Small pores in the distal part of the shield and mural septular pores may be present in frontal wall. Areolar pores absent. Avicularia absent. No interzooidal pores. Embryos incubated in internal brood sacs in dimorphic zooids. Distribution: Tropical waters of the Indo-Pacific (from Indonesia to Australia). Remarks: Based on frontal wall characteristics Euthyrisellidae was recently redefined by Gordon & Sanner (2020), who reassigned Pseudoplatyglena Gordon & d’Hondt, 1997 to Clathrolunulidae Gordon & Sanner, 2020. Family Clathrolunulidae Gordon & Sanner, 2020 Type genus: Clathrolunula Gordon & Sanner, 2020. Type species: Schizorthosecos radiatum Canu & Bassler, 1920. Other genera: Pseudoplatyglena Gordon & d’Hondt, 1997. Diagnosis: Colonies discoidal and cupuliform, with exterior basal walls, or erect, cellariiform. Orifice with condyles; proximal rim forms suboral radiate bar. Hypostegal coelom of suboral area discontinuous with rest of frontal shield and with subhypostegal extension. Frontal shield with marginal pores, communicating with visceral coelom in discoidal colonies but with axial coelom in cellariiform colonies. Spines absent. Heterozooids may be present, distolateral to the orifice. Embryonic incubation uncertain. Ancestrula not reported. Distribution: Tropical waters of the Indo-Pacific (New Caledonia). Remarks: Clathrolunulidae was erected to accommodate two species, Schizorthosecos radiatum Canu & Bassler, 1920 described from Gosport Sand, Middle Eocene of Alabama, USA, and designated as type species of Clath­ rolunula Gordon & Sanner, 2020 and Pseudoplatyglena mirabilis Gordon & d’Hondt, 1997, type species of Pseu­ doplatyglena, endemic to New Caledonia. This family is distinguished from Euthyrisellidae by having a radiate suboral bar with hypostegal coelom disconnected from the hypostegal coelom of the remainder of the frontal shield and by having interzooidal pores.

11.3 Systematics of Cheilostomata 

 369

Family Neoeuthyrididae Gordon & Sanner, 2020 Type genus: Neoeuthyris Bretnall, 1921. Type species: Euthyris woosteri MacGillivray, 1891. Other genera: None. Diagnosis: Colonies encrusting, basal wall centrally uncalcified, closely adherent to substratum without intervening coelom. Autozooids with hyaline frontal shield, thinly calcified, sometimes with protuberances. No visceral-to-hypostegal communication via marginal areolar septular pores in frontal shield; instead, hypostegal coelom, which is proximally deep, communicates with proximal zooid via multiporous septulum in transverse wall. Subfrontal hypostegal coelom lacking. Spines absent. Avicularia may be present, adventitious, latero-oral. Ovicells endozooidal, cleithral; female zooids with dimorphic orifice. Ancestrula not reported. Distribution: Western and Eastern Australia. Remarks: The dimorphic zooids and visceral-to-hypostegal communication via the proximal transverse wall support inclusion of Neoeuthyris (thus Neoeuthyrididae) in the Euthyriselloidea.

11.3.7.18 Superfamily Hippothooidea Busk, 1859b Family Hippothoidae Busk, 1859b Type genus: Hippothoa Lamouroux, 1821. Type species: Hippothoa divaricata Lamouroux, 1821. Other genera: Antarctothoa Moyano G., 1987, Austrothoa Moyano G., 1987, Celleporella Gray, 1848, Haplota Marcus, 1940, Jessethoa Gordon, 2020, Neothoa Moyano G., 1987, Plesiothoa Gordon & Hastings, 1979. Diagnosis: Colony encrusting or erect, with morphologies very variable between genera; from crustose sheet-like colonies, unilaminar or multilaminar to highly branching uniserial chains of zooids or erect forms with uniserial colonies to robust multiserial colonies. Frontal shield gymnocystal, thin and characteristically non-pseudoporous (one partial exception) with primary orifice schizoporelloid, without spines. Hypertrophied, vicarious avicularia rare. Frontal or basal pore-chambers present, small, within the wall, or tubular. Zooidal polymorphism distinct, with feeding zooids, female zooids, male zooids, and in some cases dwarf zooids (zooeciules) of unknown function. Embryos incubated in hyperstomial terminal cleithral ovicells. Ovicellate zooids dimorphic. Larva non-feeding (Fig. 11.32 A, B). Distribution: Cosmopolitan. Remarks: Representatives of Hippothoidae exist from the Late Cretaceous to the present day (Voigt 1979b, Voigt & Hillmer 1983). They are one of the best examples of a fully gymnocystal frontal shield (Gordon 2000). There are

370 

 11 Gymnolaemata, Cheilostomata

Fig. 11.32: (A, B) Hippothoidae: Hippothoa calcicola Winston & Vieira, 2013. Ilhabela, São Paulo, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, 734, paratype. (C, D) Chorizoporidae: Chorizopora brongniartii (Audouin, 1826). Gorringe seamount, Portugal NE Atlantic. University of Vienna, unregistered. Scale bars: A, 100 µm; B, D, 50 µm; C, 250 µm.

currently eight Recent hippothoid genera, with about 78 Recent species. Family Chorizoporidae Vigneaux, 1949 Type genus: Chorizopora Hincks, 1879. Type species: Flustra brongniartii Audouin, 1826. Other genera: Costulostega Tilbrook, 2006. Diagnosis: Colony encrusting, unilaminar, formed by disjunct zooids, interconnected by small tubes and even some interzooidal kenozooids. Tubes as extensions of the basal pore-chambers. Thin frontal shield gymnocystal, generally smooth or with costa-like processes. Autozooids with semicircular primary orifice, without spines. Avicularia interzooidal, interdispersed among connecting tubes or associated with ooecium. Embryos incubated in

hyperstomial terminal cleithral ovicells. Ovicellate zooids sometimes with dimorphic orifice. Larva non-feeding (Fig. 11.32 C, D). Distribution: Tropical to temperate waters. Remarks: Only two genera are included in this small family. There are 10 species worldwide, which occur mainly in tropical and temperate waters, all characterized by the disjunct zooids and the semicircular primary orifice. Chorizoporids may be unrelated to other gymnocystal-shielded ascus-bearing taxa (Powell 1967, Gordon 2000). Family Haplopomidae Gordon in De Blauwe, 2009 Type genus: Haplopoma Levinsen, 1909. Type species: Flustra impressa Audouin, 1826. Other genera: None.



Diagnosis: Colonies encrusting, unilaminar, pluriserial. Autozooids suboval or polygonal (rhomboid or hexagonal), separated by deep grooves or slightly raised walls. Gymnocystal frontal shield with variable number of marginal and frontal pseudopores; pseudopores single or cribrate, sometimes absent. Ascopore present, sometimes on a disto-median umbo. Orifice monomorphic or dimorphic, almost semicircular, without sinus except in ancestrular or periancestrular area; small condyles may be present. Spines absent. Avicularia absent. Embryos incubated in hyperstomial terminal cleithral ovicells; female zooid with dimorphic orifice; ooecium pseudoporous. Larva non-feeding. Ancestrula zooidal, single, smaller than autozooids, with sinuate orifice and without ascopore. Kenozooids absent. Interzooidal connections via well-developed basal pore-chambers.

11.3 Systematics of Cheilostomata 

 371

Distribution: Temperate waters of the European Atlantic Ocean, Mediterranean Sea and Red Sea. Remarks: Haplopoma was previously considered related to Microporellidae (Hayward & Ryland 1998). Today, this monogeneric family is known to have a gymnocystal frontal shield like other hippothooidean families, but with a distinct ascopore. A gymnocystal frontal shield with ascopore is also reported in the fossil Dysnoetoporidae Voigt, 1971, which lacks frontal pseudopores. Family Pasytheidae Davis, 1934 (1881) Type genus: Pasythea Lamouroux, 1812. Type species: Cellaria tulipifera Ellis & Solander, 1786. Other genera: Baudina Gordon, 2009a, Eutaleola Vieira & Gordon, 2010, Gemellipora Smitt, 1873, Tecatia Morris, 1980.

Fig. 11.33: (A, B) Pasytheidae: Pasythea tulipifera (Ellis & Solander, 1786). Maceió, Alagoas State, Brazil, W Atlantic Ocean. Universidade Federal de Pernambuco, unregistered. (C, D) Trypostegidae: Trypostega ilhabelae Winston & Vieira, 2013. Ilhabela, São Paulo, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo, 0735, holotype. Scale bars: A, D, 250 µm; B, 100 µm; C, 500 µm.

372 

 11 Gymnolaemata, Cheilostomata

Diagnosis: Colonies encrusting, unilaminar, uni- to pluriserial, ramifying; or erect from basal kenozooids, jointed or unjointed, branched laterally or distally; jointed colonies with bi- to multizooidal internodes, sometimes with kenozooidal internodes. Autozooids with gymnocystal frontal shield with many pseudopores. Orifice with or without condyles. Articulated oral spines absent. Avicularia absent. Horn-like latero-oral processes rarely present. Brooding zooids with dimorphic orifice. Embryos presumably incubated in internal brood sacs. Ancestrula zooidal in encrusting species or kenozooidal in erect species (Fig. 11.33 A, B). Distribution: Tropical waters. Remarks: This family is characterized by perforate gymnocystal frontal walls and the complete absence of ovicells and polymorphs like articulated oral spines and avicularia. At least three generic names are considered junior synonyms of Pasythea: Epicaulidium Hincks, 1881a, Lirizoa Lamouroux, 1816, and Tuliparia de Blainville, 1834. Davis (1934) erected the family Pasytheidae, but there are two older family-group names available – Epicaulidiidae Hincks, 1881a, and Liriozoidae MacGillivray, 1895. Despite the use of Liriozoidae by Canu and Bassler (1920), Pasyth­ eidae is almost universally used and it is to be maintained according to Article 40.2 of the International Code on Zoological Nomenclature (ICZN 1999). Family Trypostegidae Gordon, Tilbrook, & Winston in Winston, 2005 Type genus: Trypostega Levinsen, 1909. Type species: Lepralia venusta Norman, 1864. Other genera: Pulpeirina Reverter-Gil & Souto, 2015. Diagnosis: Colonies encrusting; frontal shield gymnocystal, perforated by numerous tiny tubular pseudopores. Orifice subterminal with distinctive sinus. Suboral vestigial costae fused in the proximal rim in a few taxa. Embryos incubated in subimmersed terminal cleithral ovicells. Ovicellate zooids sometimes with dimorphic orifice. Dwarf zooids or zooeciules, with operculum, located between zooids and on distal ends of ovicells; their function unknown. Vicarious avicularia present in some taxa (Fig. 11.33 C, D). Distribution: Tropical to temperate Atlantic and Pacific oceans. Remarks: Only two living genera are actually known, but 11 fossil genera belong to this family, many of which have previously been included in Hippothoidae (e.g. Voigt & Hillmer 1983). Trypostegids are fairly widespread in warmer waters and have not been found in polar waters. Hyperstomial ovicells are known in fossil species.

Family Vitrimurellidae Winston, Vieira, & Woollacott, 2014 Type genus: Vitrimurella Winston, Vieira & Woollacott, 2014. Type species: Gemellipora lata Smitt, 1873. Other genera: None. Diagnosis: Colonies encrusting, unilamellar, multiserial. Autozooids rectangular to irregularly polygonal, arranged in regular rows. Gymnocystal frontal shield with large pseudopores. Zooid with shallow sinus delimited by two to four fused vestigial costae, comprising highly reduced suboral spinocyst. Condyles present. Oral spines absent. Avicularia present or absent. Ovicells hyperstomial or endozooidal, cleithral, with pseudoporous ooecium, large or small. Ovicellate zooids with dimorphic orifice (Fig. 11.34 A, B). Distribution: Tropical waters. Remarks: This monogeneric family was erected to include some living species previously inadvertently assigned to the lepralioid fossil genus Tremoschizodina Duvergier, 1920 and to two Indo-Pacific species previously assigned to Figularia Jullien, 1886. There are some similarities between vitrimurellids and trypostegid fossils, including the gymnocystal frontal shield and the proximal sinus delimited by fused vestigial costae, but a close relationship between these families is unlikely since Trypostega is thought to have a pliophloeine ancestor (Gordon 2000). The presence of a medial suture on the proximal part of ooecia in some species of Vitrimur­ ella species may indicate a relationship with some Figu­ laria species. Thus, Vitrimurellidae seems to be a highly derived cribrilinid clade.

11.3.7.19 Superfamily Lepralielloidea Vigneaux, 1949 Family Lepraliellidae Vigneaux, 1949 Type genus: Lepraliella Levinsen, 1916. Type species: Cellepora ramulosa forma contigua Smitt, 1868b. Other genera: Acanthophragma Hayward, 1993, Cellepo­ raria Lamouroux, 1821, Drepanophora Harmer, 1957, Kladap­ heles Gordon, 1993, Schizocoryne Hayward & Winston, 2011, Sinuporaria Pouyet, 1973, Sphaeropora Haswell, 1880. Diagnosis: Multilayered colonies, encrusting and/or erect, branched, zooids often irregularly disposed when frontal budding occurs. Autozooids with umbonuloid frontal shield, sinuate or nearly semicircular, and bordered by marginal pores; often well-developed lateral condyles. Oral spines present or absent. Peristome variably developed. Umbo or avicularia present, usually associated with the orifice. Adventitious avicularia small, crossbar often with ligula. Vicarious avicularia lingulate



11.3 Systematics of Cheilostomata 

 373

Fig. 11.34: (A, B) Vitrimurellidae: Vitrimurella fungens (Marcus, 1955). Brazil, W Atlantic Ocean. Museum of Comparative Zoology Cambridge, USA, 137444, Photo: J. E. Winston. (C, D) Lepraliellidae: Celleporaria atlantica (Busk, 1884). Bahia, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade Federal da Bahia, 1320. Scale bars: A, 1 mm; B, D 250 µm; C, 500 µm.

or spatulate. Ovicells hyperstomial, non-cleithral; ooecia often covered by secondary calcification. Larva non-feeding. Mural or basal pore-chambers present (Fig. 11.34 C, D). Distribution: Cosmopolitan. Remarks: Seven Recent genera belong to this family, comprising about 125 species. Some of the genera are taxonomically very difficult, and the real number of extant species is uncertain. For example, the genus Celleporaria comprises around 103 described species, many of which were not adequately characterized (or even illustrated) when first described. Family Atlantisinidae Berning, Harmelin, & Bader, 2017 Type genus: Atlantisina Berning, Harmelin & Bader, 2017. Type species: Atlantisina atlantis Berning, Harmelin & Bader, 2017.

Other genera: Bathycyclopora Berning, Harmelin & Bader, 2017, Calvetopora Berning, Harmelin & Bader, 2017. Diagnosis: Colonies encrusting. Autozooidal frontal shield umbonuloid, imperforate or pseudoporous. Orifice with condyles and oral spines. Avicularia adventitious and/or interzooidal, present only in some taxa. Interzooidal communications via single or a few septular pores in basal pore-chambers. Ovicells hyperstomial, often terminal, acleithral with tubular opening. Ectooecium partially calcified. Ancestrula tatiform, without cryptocyst (Fig. 11.35 A, B). Distribution: Temperate waters of the northeastern Atlantic Ocean and deep-water in the Kuril-Kamchatka Trench, northwestern Pacific. Remarks: The family was erected by Berning et al. (2017) to include new taxa in three new genera. Atlantisinid

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 11 Gymnolaemata, Cheilostomata

Fig. 11.35: (A, B) Atlantisinidae: Atlantisina tricornis Berning, Harmelin & Bader, 2017. Galicia Bank, Spain, NE Atlantic. University of Vienna, unregistered. (C, D) Bryocryptellidae: Bryocryptella torquata (Jullien in Jullien & Calvet, 1903). Hirondelle, Station 58, 43° 39′ 59.9976′′ N; 8° 54′ 59.9976′′ E, Bay of Biscay, NE Atlantic Ocean off the Spanish coast. Muséum National d’Histoire Naturelle, Paris, IB-2008-4166, syntype. Scale bars: A, C, 500 µm; B, D, 250 µm.

species share characters with the Romancheinidae Jullien, 1888 (formerly including Escharellidae Levinsen, 1909 and Exochellidae Bassler, 1935). Family Bryocryptellidae Vigneaux, 1949 Type genus: Bryocryptella Cossmann, 1906. Type species: Cryptella torquata Jullien in Jullien & Calvet, 1903. Other genera: Bryobuchneria nom. nov., Cystisella Canu & Bassler, 1917, Marguetta Jullien in Jullien & Calvet, 1903, Palmiskenea Bishop & Hayward, 1989, Porella Gray, 1848, Porelloides Hayward, 1979, Rhamphosmittina Hayward & Thorpe, 1988b, Simibryocryptella Álvarez, 1991. Diagnosis: Colony encrusting, or fixed-erect with branching, rigid and heavily calcified bilamellar expansions. Autozooids

hexagonal, oval or pyriform, arranged in quincunx or straight rows. Frontal shield umbonuloid, convex, imperforate centrally, bordered by areolar-septular pores. Primary orifice immersed, with or without lyrula; condyles variably developed. Secondary orifice peristomial, mostly cormidial, formed by flanking lateral lappets or fully encircling primary orifice, often incorporating avicularian cystid or/and spiramen sinus proximally and connecting proximolateral corners of ooecium. Articulated oral spines present or absent. Suboral avicularium mainly present. Additional adventitious and vicarious avicularia may be present. Embryos incubated in hyperstomial acleithral ovicells, with ooecium rapidly overgrown by secondary calcification; ectooecium sometimes with pseudopores, often concealed by secondary calcification. Larva non-feeding (Fig. 11.35 C, D).



Distribution: Cosmopolitan. Remarks: Bryocryptellidae has much in common with Romancheinidae. In general, whereas all bryocryptellids have ovicells with the ectooecium either wholly calcified, or pseudoporous, it is largely membranous in typical romancheinids (see Ostrovsky 2013). There can be exceptions in ooecium structure among the families Lepraliellidae, Romancheinidae, Bryocryptellidae, and Umbonulidae, however, and the disposition of genera needs to be restudied, based on ovicellar and other morphological characters, as well as on genetic data. Bryobuchneria nom. nov. is here proposed as a replacement name for Buchne­ ria Harmer, 1957, which is a secondary homonym of the diplopod Buchneria Verhoeff, 1941. Family Dhondtiscidae Gordon, 1989a Type genus: Dhondtiscus Gordon, 1989a. Type species: Dhondtiscus sphericus Gordon, 1989a. Other genera: None. Diagnosis: Colony initially subconical, becoming pinecone shaped or spherical when mature, ancestrular apex facing the water column. Autozooids arranged in a subregular longitudinal series around the colony axis, which is open or closed. Zooids budding antapically. Frontal shield umbonuloid, granular, convex with marginal areolar-septular pores. Vertical walls deep. Orifice with broad shallow poster or this barely differentiated. No condyles. Diminutive oral-spine bases present or absent. Avicularia present, adventitious, oral. Ovicells hyperstomial, acleithral, with membranous ectooecium and thick calcified endooecium. Ancestrula kenozooidal, surrounded by six autozooids. Rhizoids lacking or, if axial cavity present, long and slender and produced from zooidal basal walls lining cavity. Distribution: Temperate deep sea off the east coast of New Zealand, southwestern Pacific Ocean. Remarks: This monogeneric family includes two Zealandian species. There are zooidal similarities between dhondtiscids and the lepraliellid genera Celleporaria Lamouroux, 1821 and Sphaeropora Haswell, 1880, including frontal shield (imperforate with marginal areolae), orifice (lacking sinus and with suboral avicularium) and ovicell (not closed by the operculum). New data (recorded here based on the discovery of more specimens) allow the discrimination of Dhondtiscidae from Sphaeropora by its colony form, having an apical ancestrula surrounded by six autozooids, and thin rootlets that emerge from basal walls of zooids lining the axial cavity when this is present. Family Hincksiporidae Powell, 1968 Type genus: Hincksipora Osburn, 1952. Type species: Mucronella spinulifera Hincks, 1889.

11.3 Systematics of Cheilostomata 

 375

Other genera: None. Diagnosis: Colony encrusting. Autozooids large, arranged in quincunx. Frontal shield umbonuloid, strongly thickened, coarsely granulated to dimpled, with a row of small marginal pores. Orifice subquadrate to transversely oval, proximal margin straightened, with small central denticle. Operculum heavily cuticularized. Embryos incubated in subimmersed cleithral ovicells with membranous ectooecium and calcified endooecium. Spines and avicularia lacking. Multiporous septula present in lateral and distal wall. Distribution: Temperate to polar waters of the Northwestern Pacific Ocean and the Arctic Ocean. Remarks: This family is monotypic for Hincksipora established by Osburn (1952) for Mucronella spinulifera Hincks, 1889 and a senior objective synonym of Escharelloides Kluge, 1962. Historically, this species has been attributed to Discoporidae Smitt, 1868c (Smitt 1872), Escharidae Johnston 1838 (Hincks 1889, Whiteaves 1901), Smittinidae Levinsen, 1909 (Osburn 1933), Hippothoidae Busk, 1859b (Osburn 1952, Bassler 1953), Escharellidae Levinsen, 1909 (Kluge 1962), and Hincksiporidae (Powell 1968). In the diagnosis for Hincksipora, Osburn (1952 p. 282) described the operculum of H. spinulifera as simple and firmly attached to the putative floor of the compensation sac. Powell (1968) concluded that Hincksipora possessed an umbonuloid frontal shield and established a new family. The umbonuloid nature of the frontal wall in H. spinulifera was subsequently confirmed by Gordon (1993). Family Jaculinidae Zabala, 1986 Type genus: Jaculina Jullien in Jullien & Calvet, 1903. Type species: Jaculina blanchardi Jullien in Jullien & Calvet, 1903. Other genera: None. Diagnosis: Colony erect and branching. Branches formed by uniserial or biserial autozooids. Autozooids large, frontal shield smooth or with faint texturing and pores. Primary orifice, circular, with sinus. Oral spines present or absent. Avicularia monomorphic or polymorphic, at least one situated proximal to the orifice. Rhizoids present, each growing from oval pore in the distal margin of the abfrontal wall. Embryos incubated in hyperstomial acleithral ovicells with mainly membranous ectooecium (Fig. 11.36). Distribution: Temperate waters of the Northeastern Atlantic Ocean and the Mediterranean Sea. Remarks: Only one Recent and one fossil genus are included in this family, with only four Recent species occurring in the bathyal Eastern Atlantic and Mediterranean Sea. Jaculinidae was not originally included in any superfamily (Zabala 1986), but later assigned to

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 11 Gymnolaemata, Cheilostomata

Fig. 11.36: (A, B) Jaculinidae: Jaculina blanchardi Jullien in Jullien & Calvet, 1903. Hirondelle, Station 247, 30° 24′ 0′′ N; 30° 21′ 23.9976′′ E, N Atlantic Ocean, 318 m depth. Muséum National d’Histoire Naturelle, Paris, IB-2008-2253, syntype. Scale bars: A, 500 µm; B, 100 µm.

Schizoporelloidea. Recently, Zágoršek et  al. (2014) suggested the inclusion of the Jaculinidae in the superfamily Lepralielloidea based on ovicell type and umbonuloid components in the frontal shield of the fossil genus Pirabasoporella Zágoršek, Ramalho, Berning, & Araújo Távora, 2014. Family Metrarabdotosidae† Vigneaux, 1949 Type genus: Metrarabdotos† Canu, 1914. Type species: Eschara monilifera† Milne Edwards, 1836c. Other genera: Aequilumina Gontar, 2002, Polirhabdotos Hayward & Thorpe, 1987. Diagnosis: Colonies encrusting, unilamellar, oligo- to multiserial; or erect from an encrusting base, frondose, bilamellar or cylindrical, branching. Autozooids arranged in regular rows, longitudinally hexagonal. Frontal shield umbonuloid, smooth, with conspicuous marginal pores. Primary orifice almost circular and obscured by peristome, sometimes with pseudosinus, sinus, denticles or ridges on its inner surface. Embryos incubated in very large immersed cleithral ovicells of dimorphic zooids; ooecium with membranous ectooecium and calcified endooecium with complex relief and, often, numerous pseudopores. Avicularia present, adventitious, lateral oral. Spines absent. Larva non-feeding. Ancestrula autozooidal (Fig. 11.37 A, B). Distribution: Tropical to polar waters of the Atlantic and Southern oceans. Remarks: None. Family Romancheinidae Jullien, 1888 Type genus: Romancheina Jullien, 1888. Type species: Romancheina martiali Jullien, 1888.

Other genera: Allerescha Gordon, 1989c, Antarcticaetos Hayward & Thorpe, 1988b, Arctonula Gordon & Grischenko, 1994, Bioptica Gordon, 2014, Bostrychopora Hayward & Thorpe, 1988b, Cheilonella† Koschinsky, 1885, Elleschara† Gordon, 1984, Escharella Gray, 1848, Escharoides Milne Edwards, 1836b, Exochella Jullien, 1888, Hellerasca Gordon, 1989a, Hemicyclopora Norman, 1894, Hippopleu­ rifera† Canu & Bassler, 1925b, Lageneschara Hayward & Thorpe, 1988b, Metrarabdotomorpha d’Hondt, 1983, Neola­ genipora Vigneaux, 1949, Perigastrella Canu & Bassler, 1917, Pseudosclerodomus d’Hondt & Schopf, 1985, Psilopsella Canu & Bassler, 1927, Ragionula Canu & Bassler, 1925a, Temacheilonella d’Hondt, 2017, Temachia Jullien, 1882a. Diagnosis: Colonies encrusting, uni- to multilamellar, uni to multiserial; or erect from an encrusting base, bilamellar or cylindrical, branching. Autozooids longitudinally rectangular or hexagonal. Frontal shield umbonuloid, smooth, with conspicuous marginal pores. Primary orifice almost circular, sometimes obscured by well-developed peristome that may have lyrula, condyles, or ridges on its inner surface. Oral spines present or absent. Avicularia present, adventitious, single or paired. Embryos incubated in internal brood sacs or ovicells, hyperstomial or subimmersed, acleithral, with membranous ectooecium and calcified endooecium. Larva non-feeding. Basal pore-chambers present. Ancestrula tatiform (Figs. 11.37 C, D & 11.38). Distribution: Cosmopolitan. Remarks: We follow Cook et  al. (2018) in including the families Escharellidae Levinsen, 1909 and Exochellidae Bassler, 1935 in the concept of the Romancheinidae, in contradistinction to Hayward and Ryland (1999), who retained Escharellidae.



11.3 Systematics of Cheilostomata 

 377

Fig. 11.37: (A, B) Metrarabdotosidae: Metrarabdotos jani Winston, Vieira & Woollacott, 2014. Rio de Janeiro, Brazil, W Atlantic Ocean. Museum of Comparative Zoology Cambridge, USA, 137457, holotype. (C, D) Romancheinidae: Temachia opulenta Jullien, 1882a. Travailleur, Station DR02(bis), 41° 43′ 0.0156′′ N; 9° 19′ 0.012′′ E, NE Atlantic Ocean off the Portuguese coast, 1068 m depth. Muséum National d’Histoire Naturelle, Paris, IB-2008-2979, paralectotype. Scale bars: A, 2.5 mm; B, D, 250 µm; C, 500 µm.

Family Sclerodomidae Levinsen, 1909 Type genus: Sclerodomus Levinsen, 1909. Type species: Bifaxaria denticulata Busk, 1884. Other genera: Cellarinella Waters, 1904, Cellarinelloides Moyano G., 1970b, Systenopora Waters, 1904. Diagnosis: Colonies erect, rigid, well calcified, bilamellar, irregularly branched, anchored by chitinous or calcified kenozooids (rootlets or other polymorphs). Autozooids almost longitudinally hexagonal, growing in quincunx. Frontal shield umbonuloid, with conspicuous areolar pores. Primary orifice obscured by well-developed peristome. Oral spines absent. Avicularia present, adventitious, present near or within secondary orifice. Ovicells hyperstomial with ooecium sometimes totally embedded in secondary calcification and reminiscent of endozooidal

ovicells, acleithral, opening into peristome; ectooecium without membranous areas. Distribution: Polar waters of the Southern Hemisphere. Remarks. Gordon (1988) included Cellarinella, Cellarinel­ loides, and Systenopora, in the Sclerodomidae, which is a senior synonym of Cellarinellidae Moyano G., 1970b. Family Tessaradomidae Jullien in Jullien & Calvet, 1903 Type genus: Tessaradoma Norman, 1869. Type species: Onchopora borealis Busk, 1860b. Other genera: Smithsonius Gordon, 1988. Diagnosis: Erect colonies with rather small basal encrusting portion. Primary orifice in deep, tubular peristome. Frontal shield with a tubiform spiramen and marginal pores. Ovicells with ooecia totally embedded in secondary

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 11 Gymnolaemata, Cheilostomata

Fig. 11.38: Romancheinidae. (A, B) (Former Escharellidae): Escharella ventricosa (Hassall, 1842). Galician coast, Spain, NE Atlantic. Museo de Historia Natural of the University of Santiago de Compostela, MNHNUSC-Bry-177. (C, D) (Former Exochellidae): Exochella frigidula Winston, Vieira & Woollacott, 2014. Rio de Janeiro, Brazil, W Atlantic Ocean. Museum of Comparative Zoology Cambridge, USA, 137456, holotype. Scale bars: A, 500 µm; B, 100 µm; C, D, 250 µm.

calcification, opening inside of peristome; ooecium small, without cuticular areas (Fig. 11.39 A, B). Distribution: Tropical to temperate waters. Remarks: Only two Recent genera and seven species belong to the family. Some specimens of Tessaradoma have been recorded from shallow waters (50 m depth), but most representatives of the family are found in deep water, reaching 5265 m in the case of Smithsonius quadra­ tus (Grischenko et al., 2019). Family Umbonulidae Canu, 1904b Type genus: Umbonula Hincks 1880a. Type species: Cellepora verrucosa Esper, 1790 (= Umbonula ovicellata Hastings, 1944). Other genera: Astochoporella Hayward & Thorpe, 1988b, Desmacystis Osburn, 1950, Escharopsis Verrill, 1879c,

Oshurkovia Grischenko & Mawatari, 2005, Posterula Jullien in Jullien & Calvet, 1903, Rhamphostomella von Lorenz, 1886. Diagnosis: Colonies encrusting, multiserial, unilaminar; or erect, arising from encrusting base and forming extensive bilamellar expansions; deeply pigmented, with bright embryos. Autozooids hexagonal, oval to pyriform, arranged in quincunx. Frontal shield umbonuloid, lacking central pseudopores, with a well-developed series of marginal areolae, and strongly reduced to a moderately exposed area of visible frontal membrane. Primary orifice without or with condyles and lyrula. Peristome present or absent. Secondary orifice cormidial. Spines usually absent, if present: oral, articulated. Avicularia commonly suboral or/and adventitious, frontal, or absent; if present, they are oral and articulated. Embryos incubated



11.3 Systematics of Cheilostomata 

 379

Fig. 11.39: (A, B) Tessaradomidae: Tessaradoma boreale (Busk, 1860b). BANGAL 0811, Station V01, NE Atlantic Ocean off the Spanish coast, 867 m depth. Museo Nacional de Ciencias Naturales, Madrid, 25.03/3958. (C, D) Umbonulidae: C, Rhamphostomella bilaminata (Hincks, 1877), Barents Sea, Kluge Collection, PSU 810.22. D, Rhamphostomella ussowi (Kluge, 1908), Barents Sea, Kluge Collection, PSU 810.24. Scale bars: A, C, 500 µm; B, D, 250 µm.

in internal brood sacs, or in immersed or hyperstomial ovicells, cleithral, semicleithral, or subcleithral; ooecium often embedded in secondary calcification; ectooecium uni- to multipseudoporous or with enlarged foramen. Larva non-feeding. Basal pore-chambers or mural septula present. Basal wall fully calcified, sometimes with protuberances. Ancestrula tatiform in genera where observed (Fig. 11.39 C, D). Distribution: Boreal-Arctic waters, predominantly North Pacific, and Antarctic. Remarks: Although only 10 (3 fossil and 7 Recent) genera are included in Umbonulidae, it remains a rather heterogeneous and morphologically diverse group. The combination of characters intrinsic to the type species of Umbonula, U. ovicellata, includes the following: 1) umbonuloid frontal shield with well-developed marginal areolar-septular pores, 2) median suboral avicularium,

and 3) hyperstomial ovicell with pseudoporous ooecium. At the same time, some umbonulid taxa possess more limited or, conversely, an extended set of taxonomic criteria relative to those in the type genus: 1) lack of ovicells and avicularia (Oshurkovia); 2) immersed ovicells with vestigial ooecium comprising a small, transversely elongate hood suspended in the distal curvature of the aperture (Desmacystis); 3) ectooecium with only a single pseudopore or enlarged foramen (Escharopsis and Asto­ choporella); and 4) permanent presence of hollow articulated oral spines (some species of Rhamphostomella). In addition to this, the presence of frontal adventitious avicularia is restricted to certain species belonging to only three Recent genera (Umbonula patens (Smitt, 1868b), Astochoporella cassidula Hayward & Thorpe, 1988b, and a few species of Rhamphostomella). Although the status of Umbonulidae was discussed by Cheetham (1968)

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 11 Gymnolaemata, Cheilostomata

and Gordon (1984), the family might still need a deeper revision. According to Taylor (1993), the earliest fossil umbonulids were documented from the Late Eocene of France.

11.3.7.20 Superfamily Mamilloporoidea Canu & Bassler, 1927 Family Mamilloporidae Canu & Bassler, 1927 Type genus: Mamillopora Smitt, 1873. Type species: Mamillopora cupula Smitt, 1873. Other genera: Anoteropora Canu & Bassler, 1927. Diagnosis: Colonies cup-shaped, convex frontally, concave basally, unilamellar, multiserial, supported above the substratum by rhizoids emanating from the

basal surface. Autozooids arranged in regular rows radiating from the centrally located ancestrula, hexagonal, opening on the convex surface. Frontal shield with occasional circular areolar marginal pores. Orifice opening centrally on frontal shield, circular to elliptical, with lateral, proximally directed condyles. Avicularia present, adventitious, oral. Septula basal, multiporous. Ovicell hyperstomial or subimmersed, cleithral, often terminal, orifice of fertile zooid dimorphic; ectooecium membranous except at ovicell opening, endooecium calcified with numerous small pseudopores. Ancestrula autozooidal, located in center of colony, surrounded by six zooids. Kenozooids may be present, usually on the basal side. Spines absent (Fig. 11.40 A, B). Distribution: Tropical waters of the Atlantic and Indian oceans.

Fig. 11.40: (A, B) Mamilloporidae: Anoteropora latirostris Silén, 1947. Murray, Station 75, Gulf of Oman, NW Indian Ocean, 115 fms depth. Natural History Museum, London, 1963.8.10.18 (two specimens). (C, D) Ascosiidae: Ascosia pandora Jullien, 1882a. Travailleur, Station DR01(bis), 43° 1′ 0.012′′ N; 9° 37′ 0.012′′ E, NE Atlantic Ocean off the Spanish coast, 2018 m depth. Muséum National d’Histoire Naturelle, Paris, IB-2008-2505, syntype. Scale bars: A, 1 mm; B, 250 µm; C, 500 µm; D, 100 µm.



Remarks: Mamilloporids have colonies comparable to those of cupuladriids and the lunulitoid families. However, these other families lack a frontal shield and ascus. Furthermore, the zoarium of mamilloporids is anchored to the substratum by rhizoids emanating from the basal surface. The oldest known mamilloporids are from the Eocene (Cook & Chimonides 1994b). The latter authors also provided a revision of Mamilloporidae and the genus Anoteropora. In their view, Ascosiidae should be included in Mamilloporidae. Family Ascosiidae Jullien, 1882a Type genus: Ascosia Jullien, 1882a. Type species: Ascosia pandora Jullien, 1882a. Other genera: Fedora Jullien, 1882a. Diagnosis: Colonies discoidal to conical, convex frontally, concave basally, unilamellar, multiserial, supported above the substratum by rhizoids. Autozooids arranged in regular rows radiating from the centrally located ancestrula, recumbent to suberect, tubular, opening on the convex surface. Frontal shield smooth, imperforate. Orifice subterminal, circular to longitudinally elliptical, with lateral, proximally directed condyles. Septula may be present, basal, multiporous. Ovicells absent or hyperstomial, terminal, cleithral. Avicularia absent or present, adventitious, oral. Ancestrula autozooidal, located in the center of the colony, surrounded by about six zooids. Spines absent (Fig. 11.40 C, D). Distribution: Tropical to temperate waters of the Atlantic Ocean of the Northern Hemisphere and tropical waters of the Pacific Ocean off New Caledonia. Remarks: Ascosiidae was a rarely used family name until Gordon and d’Hondt (1997) endorsed its validity. Ascosiid taxa had previously been included into the Mamilloporidae (e.g. Cook & Chimonides 1994b). The two extant genera included in this family differ in many respects; for example, in Fedora, ovicells are lacking, autozooids are recumbent and basal pore-chambers are present, while in Ascosia, ovicells are hyperstomial, autozooids are suberect, and basal pore-chambers are lacking. The family concept of Ascosiidae may therefore be in need of revision. Family Cleidochasmatidae Cheetham & Sandberg, 1964 Type genus: Cleidochasma Harmer, 1957 (= Characodoma Maplestone, 1900). Type species: Gemellipora protrusa Thornely, 1905. Other genera: Anchicleidochasma Soule, Soule & Chaney, 1991b, Calyptooecia Winston, 1984b, Cleidochas­ midra Ünsal & d’Hondt, 1979, Fedorella Silén, 1947, Gemel­ liporina Bassler, 1936, Yrbozoon Gordon, 1989a.

11.3 Systematics of Cheilostomata 

 381

Diagnosis: Colonies encrusting, unilamellar to multi­ lamellar, multiserial; or erect from an encrusting base, branching, multiserial, supported above the substratum by rhizoids. Autozooids arranged in regular rows, hexagonal, sexually dimorphic in one genus. Frontal shield granular to tuberculate, with a single row of circular areolar pores at the margin. Orifice terminal, keyhole shaped, with lateral, proximally directed condyles. Spines usually absent; if present oral, non-articulated. Septula basal, multiporous. Embryos incubated internally (presumably in brood sacs) or in hyperstomial non-cleithral ovicells, ooecium often covered by or almost totally embedded in secondary calcification; ectooecium membranous or with pseudopores. Avicularia present, single or paired, adventitious, oral. Ancestrula autozooidal. Kenozooids may be present (Fig. 11.41 A, B). Distribution: Tropical and temperate waters. Remarks: Although containing relatively few genera, Cleidochasmatidae is a rather heterogeneous family that is currently mainly defined by the appearance of the ovicell and the frontal shield. The name-bearing genus of the family, Cleidochasma Harmer, 1957, with type species Gemellipora protrusa Thornely, 1905, is considered a junior synonym of Characodoma Maplestone, 1900 (Cook & Bock 1996). Nonetheless, Gemellipora protrusa Thornely, 1905 remains the nominal type species of Cleidochasmatidae according to Article 40.1 of the International Code on Zoological Nomenclature (ICZN 1999). The placement of Yrbozoon Gordon, 1989a, which initially was included by Gordon (1989a) in Celleporidae, into Cleidochasmatidae was based on the ovicell (Gordon & d’Hondt 1997). It is currently the only mamilloporoid genus having calcified, non-articulated, oral spines. The affinity of Yrbozoon to Cleidochasmatidae remains somewhat questionable. Calyptooecia shows zooidal dimorphism, with female zooids internally brooding embryos (presumably in brood sacs) and having smaller orifices. Family Crepidacanthidae Levinsen, 1909 Type genus: Crepidacantha Levinsen, 1909. Type species: Crepidacantha poissoni crinispina Levinsen, 1909. Other genera: None. Diagnosis: Colonies encrusting, unilamellar, multiserial. Autozooids arranged in regular rows, hexagonal. Frontal shield smooth to granular, with a single row of elliptical to slit-like areolar pores at the margin, each separated by a recumbent spine that lies flat against the substratum. Orifice subterminal, trifoliate with orificial indentations at the proximolateral margins, with lateral, proximally directed condyles. Spines present in early ontogeny of autozooids, oral, usually not calcified and

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Fig. 11.41: (A, B) Cleidochasmatidae: Characodoma strangulatum (Calvet, 1906). BALGIM, Station CP92, 34° 24′ 17.9964′′ N; 7° 30’ 18′′ E, NE Atlantic Ocean off the Moroccan coast, 1182 m depth. Muséum National d’Histoire Naturelle, Paris, IB-2008-19804, syntype. (C, D) Crepidacanthidae: Crepidacantha longiseta Canu & Bassler, 1928b. Bahia, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade Federal da Bahia, 3161. Scale bars: A, C, 500 µm; B, D, 100 µm.

getting lost during later ontogeny. Septula basal, multiporous. Embryos incubated in terminal subimmersed or hyperstomial cleithral ovicells; ectooecium with arch-like or oval membranous area and calcified bars of endooecium underneath. Avicularia present, paired, adventitious, adjacent to or proximolateral of orifice, derived from frontal septular pores, bearing setiform mandibles. Ancestrula tatiform (Fig. 11.41 C, D). Distribution: Tropical to temperate waters. Remarks: Crepidacanthidae is a monogeneric family comprising about 15 species and with worldwide distribution in tropical to temperate waters. The orifice of all crepidacanthids has a characteristic trifoliate shape with orificial indentations at the proximolateral corners. Weakly calcified spinous processes develop around the periphery of the zooid.

11.3.7.21 Superfamily Pseudolepralioidea Silén, 1941 Family Pseudolepraliidae Silén, 1941 Type genus: Pseudolepralia Silén, 1941. Type species: Pseudolepralia ellisinae Silén, 1941. Other genera: None. Diagnosis: Colonies encrusting. Autozooids arranged quincuncially, each zooid surrounded by six zooids and six interzooidal avicularia. Frontal shield umbonuloid arching over and tucking in behind the zooidal rim; areola absent. Spines absent. Opercular flap broad. Avicularia interzooidal. Ovicells hyperstomial; ectooecium membranous, endooecium calcified with relief similar to the frontal shield. Ancestrula not reported. Interzooidal communication via uniporous and multiporous septula.



Distribution: Tropical waters of the Pacific Ocean at the Bonin Islands, Japan. Remarks: This monospecific family has a distinct umbonuloid frontal shield with no areolae. The systematics of this curious family have been discussed by Silén (1942), Sandberg (1977, pp. 164–165), and Gordon (1993).

11.3.7.22 Superfamily Schizoporelloidea Jullien, 1882a Family Schizoporellidae Jullien, 1882a Type genus: Schizoporella Hincks, 1877a. Type species: Lepralia unicornis Johnston, 1847. Other genera: Fovoporella Gordon, 2014, Gemelliporidra Canu & Bassler, 1927, Schizobrachiella Canu & Bassler, 1920, Stylopoma Levinsen, 1909.

11.3 Systematics of Cheilostomata 

 383

Diagnosis: Colonies encrusting, uni- to multilaminar, multiserial; or erect from an encrusting base, cylindrical, branching, non-articulated, or uni- to multilaminar. Autozooids primarily arranged in a regular series, quadrangular to hexagonal. Lepralioid frontal shield, with pseudopores, sometimes with distinct areolar pores. Orifice with subcircular anter and distinct median U-shaped or V-shaped sinus; condyles present. Spines absent. Embryos incubated in hyperstomial ovicells, acleithral (with proximal labellum = sclerite) or non-cleithral; ectooecium membranous, endooecium calcified, pseudoporous. Larva non-feeding. Avicularia often present, adventitious or vicarious. Ancestrula autozooidal (Fig. 11.42 A, B). Distribution: Cosmopolitan. Remarks: Historically, this family has included species with a sinuate orifice and variable morphology of frontal

Fig. 11.42: (A, B) Schizoporellidae: Schizoporella errata (Waters, 1878). Santa Catarina, Brazil, W Atlantic Ocean. Universidade Federal de Pernambuco, unregistered. (C, D) Actisecidae: Actisecos regularis Canu & Bassler, 1927. Philippines, W Pacific Ocean. Smithsonian National Museum of Natural History, Washington, DC, USNM 8325, syntype. © Smithsonian National Museum of Natural History. Scale bars: A, C, 500 µm; B, D, 250 µm.

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 11 Gymnolaemata, Cheilostomata

shield and ovicells. Currently, schizoporellids are characterized by a sinuate orifice, pseudoporous frontal wall, and an endooecial surface resembling the zooidal frontal shield. Many genera previously assigned to the Schizoporellidae have been assigned to different families, including Bitectiporidae MacGillivray, 1895, Buffonellidae Jullien, 1888, Eminooeciidae Hayward & Thorpe, 1988c, Escharinidae Tilbrook, 2006, and Lacernidae Jullien, 1888. Family Acoraniidae López‐Fé, 2006 Type genus: Acorania López‐Fé, 2006. Type species: Acorania enmediensis López‐Fé, 2006. Other genera: None. Diagnosis: Colonies erect, branching. Autozooids elongate, forming alternating series on one side of the branch. Frontal shield lepralioid with pseudopores, except for an imperforate area proximal to the orifice. Orifice subcircular, with paired condyles. Spines absent. Avicularia may be present, adventitious. Ovicells hyperstomial, not closed by zooidal operculum; ectooecium membranous except for a basal rim, endooecium calcified and imperforate. Ancestrula autozooidal. Ancestrula erect, tatiform. Interzooidal communication by uniporous septula. Distribution: Temperate waters of the Atlantic Ocean off the Canary Islands, Spain. Remarks: Acoraniidae is monospecific. The type species is only known from its original locality, therefore more studies are needed to investigate the relationship between this family and other Schizoporelloidea. Family Actisecidae Harmer, 1957 Type genus: Actisecos Canu & Bassler, 1927. Type species: Actisecos regularis Canu & Bassler, 1927. Other genera: None. Diagnosis: Colonies free-living, discoidal, unilamellar, multiserial, convex frontally. Zooids elongated, arranged in regular rows. Frontal shield centrally smooth or with sporadic tubercles that are commoner on the periphery, with foramina. Basal pore-chambers present. Marginal pores absent. Basal autozooidal walls covered by flat kenozooids with a large membranous area. Primary orifice subcircular. Peristome long and cylindrical, swollen at base. Spines and avicularia absent. Ovicells prominent, peristomial, terminal, developing exclusively in peripheral zooids; ectooecium membranous, endooecium pustulose with numerous pseudopores. Ancestrula autozooidal (Fig. 11.42 C, D). Distribution: Tropical waters of the Pacific Ocean off the Philippines and Indonesia. Remarks: Actisecidae, which includes a single extant genus, externally closely resembles some species of Exechonellidae. According to Cáceres-Chamizo et al. (2017),

the monospecific fossil genus Oviexechonella Di Martino & Taylor, 2015 from the Miocene of Indonesia, previously assigned to Exechonellidae, may better be placed in Actisecidae owing to the presence of peristomial ovicells. Family Buffonellidae Jullien, 1888 Type genus: Buffonella Jullien, 1888. Type species: Buffonella rimosa Jullien, 1888. Other genera: Aimulosia Jullien, 1888, Hippadenella Canu & Bassler, 1917, Ipsibuffonella Gordon & d’Hondt, 1997, Julianca Gordon, 1989c, Kymella Canu & Bassler, 1917, Maiabuffonella Gordon & d’Hondt, 1997, Xenogma Gordon, 2014. Diagnosis: Colony encrusting, uni- to multilamellar, multiserial. Zooids arranged in regular series. Frontal shield lepralioid, smooth, with marginal areolar pores. Primary orifice with median sinus or lyrula; condyles present. Proximal peristome may form umbo. Oral spines sometimes present. Avicularia adventitious, single, suboral or frontal. Ovicells hyperstomial, subimmersed or peristomial, cleithral and (possibly) acleithral; ectooecium membranous or with oval window, endooecium calcified. Ancestrula autozooidal, smoothly calcified (Fig. 11.43 A, B). Distribution: Cosmopolitan (predominantly Southern Hemisphere). Remarks: Buffonellodidae was introduced to replace the Buffonellidae Jullien, 1888, based on Buffonella Jullien, 1888, The latter name, however, was erroneously considered preoccupied by Bufonella Girard, 1853, a now invalid name of terrestrial frogs from Australia, in Strand (1928). Strand (1928) also mistakenly attributed the authorship to Keferstein (1868), who misspelled Bufonella to Buffonella, but already indicated that Bufonella may be a junior synonym of another frog genus. Neither Bufonella nor Buffonella was accepted by subsequent herpetologists. Buffonellids are considered related to Lacernidae Jullien, 1888, but distinguished from the latter in having zooids lacking pseudopores on the frontal wall. Family Calwelliidae MacGillivray, 1887 Type genus: Calwellia Thomson, 1858. Type species: Calwellia bicornis Thomson, 1858. Other genera: Ichthyaria Busk, 1884, Ijimaiellia Gordon, 2009b, Malakosaria Goldstein, 1882, Onchoporella Busk, 1884, Onchoporoides Ortmann, 1890, Wrigiana Gordon & d’Hondt, 1997. Diagnosis: Colonies erect or semierect, anchored by rhizoides, flexible, unilaminar and unbranched, or dichotomously branching, or foliaceous. Autozooids arranged in pairs (alternating back-to-back), biserial or multiserial. Primary orifice almost subcircular, sometimes with spinous processes, perioral excavations or pore-chambers.



11.3 Systematics of Cheilostomata 

 385

Fig. 11.43: (A, B) Buffonellidae: Buffonella rimosa (Jullien, 1888). La Romanche, Tierra del Fuego, Southern Ocean. Muséum National d’Histoire Naturelle, Paris, IB-2008-45, syntype. © Muséum National d’Histoire Naturelle. (C, D) Calwelliidae: Malakosaria atlantica Vieira, Gordon, Souza & Haddad, 2010. Brazil, W Atlantic Ocean, 25° 36.99′ S; 45° 13.57′ W. Museu de Zoologia, Universidade de São Paulo, 0312, holotype. Scale bars: A, 100 µm; B, D, 50 µm; C, 500 µm.

Frontal shield smooth with distinct ascopore. Avicularia absent. Embryos incubated in hyperstomial acleithral and cleithral ovicells; ectooecium mostly membranous, endooecium calcified (Fig. 11.43 C, D). Distribution: Tropical to temperate waters. Remarks: Onchoporella buskii Harmer, 1923 was proposed as a new replacement name for Carba­ sea bombycina Busk, 1852b (non Flustra bombycina Ellis & Solander, 1786), and Harmer (1923) suggested this species should be assigned as type species of Onchoporella Busk, 1884. This was recently followed by Florence et al. (2007). Thus, in accordance with Article 69.2.3 of the International Code on Zoological Nomen­ clature (ICZN 1999), Flustra bombycina Ellis & Solander, 1786 sensu Busk, 1852b (= Onchoporella buskii Harmer, 1923) is regarded to be type species of Onchoporella.

However, Busk (1884) expressly employed the name Onchoporella bombycina in the sense of a misidentification, thus is deemed to denote a new nominal species, and the specific name Onchoporella bomby­ cina Busk, 1884 needs to be considered available in accordance with Article 11.10 (ICZN 1999), Onchoporella bombycina Busk, 1884 then being a senior synonym of Onchoporella buskii. Orr et al. (2019b) noted many similarities between zooids of Fenestrulinidae Jullien, 1888 and Calwelliidae, suggesting the two families could be related. Colony form and the absence of avicularia are the principal characters separating calwelliids from fenestrulinids. Calwellidae is characterized by having erect to semierect colonies and weakly calcified shields with distinct ascopore (similar to those of Fenestrulinidae) and often large ovicells.

386 

 11 Gymnolaemata, Cheilostomata

Family Cheiloporinidae Bassler, 1936 Type genus: Cheiloporina Canu & Bassler, 1923. Type species: Hippoporina circumcincta Neviani, 1896a. Other genera: Cheilopora Levinsen, 1909, Cyttaridium Harmer, 1957, Hagiosynodos Bishop & Hayward, 1989, Retelepralia Gordon & Arnold, 1998. Diagnosis: Colonies encrusting, unilaminar, multiserial, sheet-like; or erect, bilaminar, or selenariiform, dome shaped. Autozooids almost polygonal, separated by distinct walls or furrows. Frontal shield lepralioid, pseudoporous, except for an imperforate area around the orifice. Primary orifice campanulate to trifoliate, with condyles; peristome imperforate, sometimes slightly raised and distinct. Spines absent. Kenozooids and adventitious avicularia present in some species. Kenozooids present in some species. Embryos incubated in internal brood sacs, immersed, subimmersed or hyperstomial cleithral ovicells; calcified endooecium pseudoporous or complete,

ectooecium membranous; maternal zooids with dimorphic orifice. Ancestrula autozooidal (Fig. 11.44 A, B). Distribution: Tropical to temperate waters. Remarks: Cheiloporinidae was introduced for some genera with endozooidal ovicells that were previously assigned to Hippopodinidae Levinsen, 1909, whose type species Hip­ popodina feegeensis Busk, 1884, has hyperstomial ovicells. Currently, Cheiloporinidae is considered morphologically diverse, with variable colonial, orificial, avicularian and ovicellar morphology. Differences between the species assigned to different genera, including both Cheilopora and Cheiloporina and their type species, indicate that this family requires proper review. Among cheiloporinids, Cyttaridium is distinguished by having dome-shaped colonies. Based mainly on the ovicell, Gordon and Arnold (1998) suggested a possible affinity between Retelepralia and Cheiloporinidae. Retelepralia is characterized by the presence of tubular interzooidal connections and a median gymnocystal strip.

Fig. 11.44: (A, B) Cheiloporinidae: Cheilopora sincera (Smitt, 1868b). Akkeshi Bay, Hokkaido, Japan, NW Pacific, intertidal. Natural History Museum, London, 2006.2.27.77 and 2006.2.27.78. (C, D) Cryptosulidae: Cryptosula zavjalovensis Kubanin, 1976. Akkeshi Bay, Hokkaido, Japan, NW Pacific, intertidal. Natural History Museum, London, 2006.2.27.61 and 2006.2.27.60. Scale bars: A–C, 500 µm; D, 250 µm.



Family Cryptosulidae Vigneaux, 1949 Type genus: Cryptosula Canu & Bassler, 1925a. Type species: Eschara pallasiana von Moll, 1803. Other genera: Harmeria Norman, 1903b. Diagnosis: Colonies encrusting, uni- to multilaminar, multiserial. Autozooids separated by distinct marginal walls. Frontal shield lepralioid, convex, pseudoporous, except by an imperforate area below the orifice; proximal umbo often present. Primary orifice semicircular to bell shaped, with lateral condyles. Spines absent. Avicularia may be present, adventitious, suboral. Kenozooids may be present, with the entire frontal surface calcified and with pseudopores, lacking orifice. Embryos incubated in internal brood sacs, sometimes in dimorphic (dwarf) maternal zooids. Larva non-feeding. Vertical wall with multiporous septula. Ancestrula autozooidal or with membranous frontal wall and without spines (Fig. 11.44 C, D). Distribution: Cosmopolitan. Remarks: Canu and Bassler (1920) cited Eschara pallasi­ ana type species of Hippodiplosia Canu, 1916. However, according to Article 67.2 of the International Code on Zoo­ logical Nomenclature (ICZN 1999), Eschara pallasiana is not eligible to be fixed as the type species of Hippo­ diplosia since this species was not originally included as nominal species in the original description of the genus. Hastings (1929) subsequently designated Hippodiplosia verrucosa Canu, 1916 from the Aquitanian of Southwestern France as the type species of Hippodiplosia. The genus is now regarded as a junior synonym of the fossil genus Reussia Neviani, 1896b (family Bryocryptellidae Vigneaux, 1949). Family Cyclicoporidae Hincks, 1884b Type genus: Cyclicopora Hincks, 1884b. Type species: Lepralia longipora MacGillivray, 1883b. Other genera: None. Diagnosis: Colonies encrusting, unilaminar, multiserial. Autozooids separated by distinct thin lines of calcification. Frontal shield lepralioid, pseudoporous, except for an imperforate area around proximal half of orifice. Primary orifice almost circular, without condyles. Spines absent. Avicularia present in some species. Kenozooids absent. Embryos incubated in hyperstomial cleithral ovicells; ooecium with pseudoporous endooecium and membranous ectooecium; maternal zooid with dimorphic orifice. Ancestrula not reported (Fig. 11.45 A, B). Distribution: Temperate waters off Victoria, Australia. Remarks: The genus Cyclicopora requires revision since there are some clear morphological differences between the type species and fossil species (described by Canu &

11.3 Systematics of Cheilostomata 

 387

Bassler 1920). These have avicularia and the morphology of the frontal shield and ovicells does not conform. Family Echinovadomidae Tilbrook, Hayward, & Gordon, 2001 Type genus: Echinovadoma Tilbrook, Hayward & Gordon, 2001. Type species: Echinovadoma anceps Tilbrook, Hayward & Gordon, 2001. Other genera: None. Diagnosis: Colonies encrusting, unilaminar, multiserial. Autozooids separated by distinct lateral walls. Frontal shield lepralioid, pseudoporous. Primary orifice subcircular with paired condyles, often with well-developed peristome. Spines absent. Avicularia and kenozooids absent. Ovicells hyperstomial, non-cleithral; ooecium with partially calcified spinous endooecium with oval foramina or triangular median slit, ectooecium membranous. Ancestrula autozooidal (Fig. 11.45 C, D). Distribution: Tropical waters of the western Pacific Ocean. Remarks: Echinovadoma was inadvertently published by Tilbrook (2000), but the type species indicated therein was not described. The name must therefore be considered as a nomen nudum. The first valid mention of the generic name is Tilbrook et al. (2001). These authors cited the characters of the ovicell and frontal shield as the basis of the family. It comprised three species in one genus that have obvious similarities to some Cleidochasmatidae. Family Eminooeciidae Hayward & Thorpe, 1988c Type genus: Eminooecia Hayward & Thorpe, 1988c. Type species: Hippadenella carsonae Rogick, 1957. Other genera: Isoschizoporella Rogick, 1960, Macrocam­ era Gordon & d’Hondt, 1997. Diagnosis: Colonies erect from an encrusting base, rigid, cylindrical and narrow, dichotomously branching and anastomosing, or bilaminar to foliaceous. Autozooids regular or irregularly arranged in the colony, longitudinally rectangular to hexagonal. Frontal shield lepralioid, with few marginal areolar pores. Orifice with proximal sinus and condyles; primary orifice often obscured by raised peristome; pseudospiramen may be present. Oral spines absent. Avicularium adventitious, suboral or frontal. Kenozooids may be present. Embryos incubated in hyperstomial ovicells, acleithral or non-cleithral; endooecium calcified, ectooecium membranous or partially calcified with membranous window. Distribution: Temperate to polar waters of the Southern Hemisphere. Remarks: The morphological characters of this family are quite heterogeneous, suggesting that it requires review.

388 

 11 Gymnolaemata, Cheilostomata

Fig. 11.45: (A, B) Cyclicoporidae: A, Cyclicopora spongiopsis (De Gregorio, 1890). Monroeville, Alabama, USA, Vicksburgian. Natural History Museum, London, D30145. B, Cyclicopora longipora (MacGillivray, 1882) South of Eucla, Western Australia. Photo: P.E. Bock. (C, D) Echinovadomidae: Echinovadoma anceps Tilbrook, Hayward, & Gordon, 2001. Okinawa, Japan, East China Sea, intertidal. National Museum of Nature and Science, Tsukuba, NSMT-Te 1148. Scale bars: A, C, 500 µm; B, 200 µm; D, 100 µm.

Family Escharinidae Tilbrook, 2006 Type genus: Escharina Milne Edwards, 1836b. Type species: Eschara vulgaris von Moll, 1803. Other genera: Allotherenia Winston & Vieira, 2013, Bry­ opesanser Tilbrook, 2006, Chiastosella Canu & Bassler in Bassler, 1934, Herentia Gray, 1848, Hippomenella Canu & Bassler, 1917, Phaeostachys Hayward, 1979, Taylorus Pérez, López Gappa, Vieira & Gordon 2020, Therenia David & Pouyet, 1978, Toretocheilum Rogick, 1960. Diagnosis: Colonies encrusting, unilamellar, multiserial; or erect from an encrusting base, cylindrical, foliaceous, or bilamellar. Autozooids arranged in regular rows or chaotic, rhomboid, hexagonal, or polygonal in shape, separated by marginal walls or deep grooves. Frontal shield lepralioid, with or without frontal pseudopores; distinct marginal areolar pores present. Primary orifice with distinct

proximal sinus and paired condyles; raised peristome may occur. Oral spines present or absent. Avicularia may be present, adventitious and mainly latero-oral, or interzooidal. Kenozooids may be present. Ovicells endozooidal to hyperstomial, sometimes terminal, cleithral; ectooecium entirely or partially membranous; maternal zooid often with dimorphic orifice. Larva non-feeding. Ancestrula tatiform, kenozooidal or autozooidal (Fig. 11.46 A, B). Distribution: Cosmopolitan. Remarks: The family was erected to accommodate schizoporelloids species with imperforate ooecia and tatiform ancestrula (Tilbrook, 2006). At least two species, Herentia hyndamanni (Johnston, 1847) and Allotherenia sabuosa Winston & Vieira, 2013, have different ancestrulae, i.e. kenozooidal and autozooidal, respectively (see Berning et al. 2008, Winston & Vieira, 2013).



Family Fatkullinidae Grischenko, Gordon, & Morozov, 2018 Type genus: Fatkullina Grischenko, Gordon, & Taylor, 1998. Type species: Fatkullina paradoxa Grischenko, Gordon, & Taylor, 1998. Other genera: Lepralioides Kluge, 1962, Pachyegis Osburn, 1952, Stomacrustula Winston & Hayward, 2012. Diagnosis: Colony encrusting, unilamellar, darkly pigmented. Zooids with or without reversed-polarity budding. Frontal shield lepralioid, evenly pseudoporous. Primary orifice with median sinus and weakly developed condylar ridges. Oral spines and avicularia absent. Embryos incubated internally or in endozooidal or hyperstomial cleithral ovicells, ooecium sometimes covered

11.3 Systematics of Cheilostomata 

 389

by secondary calcification; ectooecium membranous, endooecium calcified, sometimes with small central fenestra. Mural septula uniporous, set in buttressed recesses. Ancestrula externally resembling postancestrular zooids, but internally with reduced umbonuloid area suborally that also includes possible vestigial costal elements. Immediate periancestrular zooids part of an ancestrular complex in type genus (Fig. 11.46 C, D). Distribution: Temperate to polar waters of the Northern Hemisphere. Remarks: The family Fatkullinidae was recently erected by Grischenko et  al. (2018) to accommodate Fatkullina (the type genus) and three other genera, Stomacrustula,

Fig. 11.46: (A, B) Escharinidae: Allotherenia sabulosa Winston & Vieira, 2013. Caraguatatuba, São Paulo, Brazil, W Atlantic Ocean. Museu de Zoologia, Universidade de São Paulo. (C, D) Fatkullinidae: Fatkullina imitata Grischenko, Gordon & Morozov, 2018. Continental slope of western Kamchatka Peninsula, Sea of Okhotsk, 58.03833° N, 155.72028° E, depth 290 m. Zoological Institute, Russian Academy of Science, Saint Petersburg, ZIRAS 1/50661. Scale bars: A, D, 500 µm; B, 250 µm; C, 1 mm.

390 

 11 Gymnolaemata, Cheilostomata

Lepralioides, and Pachyegis, which earlier belonged to the family Stomachetosellidae Canu & Bassler, 1917. As Zágoršek & Gordon (2013) noted, Stomachetosellidae is very heterogeneous. It is based on the type genus and species Stomachetosella crassicollis Canu & Bassler, 1917 (described from Early Oligocene, Mississippi, USA), which has an erect bilamellar to flabellate colony form in which the zooids open on both sides, and a regularly perforated pseudoporous lepralioid frontal shield. The orifice has a tapering rounded poster and condyles appear to be lacking. Ooecia are evenly pseudoporous and, according to Canu & Bassler (1920), an inconspicuous avicularium is located near the orifice in some zooids. After Winston and Hayward (2012) compared the characters of S. cras­ sicollis to those of living species attributed to Stomache­ tosella, they concluded that the latter are unrelated to the former. They established a new genus, Stomacrustula, for all “the Recent species formerly attributed to Stoma­ chetosella,” making new combinations for three of them, Stomacrustula cruenta (Busk, 1854) (the designated type species), S. sinuosa (Busk, 1860a), and S. hincksi (Powell, 1968), but they did not make new combinations for the 10 other Recent species. Stomacrustula was not attributed to a family but left incertae sedis. In sharing such characters as 1) encrusting colony form, 2) deep-colored pigmentation, 3) thickly calcified pseudoporous frontal shields, 4) no oral spines, 5) mural septula set in buttressed recesses, 6) avicularia are lacking, and 7) ooecia are either lacking or, if present, without pseudopores or with single non-calcified fenestra, the genera of Fatkullinidae have little in common with the fossil type species of Stomachetosella, which had erect colonies, avicularia, and pseudoporous ooecia. Family Fenestrulinidae Jullien, 1888 Type genus: Fenestrulina Jullien, 1888. Type species: Cellepora malusii Audouin, 1826. Other genera: Adelascopora Hayward & Thorpe, 1988b, Tenthrenulina Gordon, 1984 (see Orr et al. 2019b, p. 196). Diagnosis: Colonies encrusting, multiserial, uni- to multilamellar, or erect, foliaceous, uni- to bilamellar. Autozooids subrectangular to hexagonal, convex, separated by raised walls. Gymnocyst developed laterally and proximally. Lepralioid frontal shield with suboral ascopore; frontal wall with sparse or fully distributed complex (rarely simple) pseudopores; marginal areolar pores present. Orifice more or less semicircular, sometimes with indentations, denticles or condyles. Oral spines mostly present. Avicularia very rare, adventitious. Ovicells hyperstomial, cleithral; ooecium with membranous ectooecium and calcified endooecium. Larva non-feeding. Abfrontal pore-chambers and radicles may be present. Ancestrula autozooidal or tatiform (Fig. 11.47 A, B).

Distribution: Cosmopolitan. Remarks: Despite the characters advanced by Soule et al. (1995) to justify the genus Fenestruloides, including the presence of avicularia (absent in Fenestrulina) and having a fully pseudoporous frontal shield (often only peripheral in Fenestrulina), Tilbrook (2006) suggested that Fenestru­ loides should be included in Fenestrulina. Gene sequencing shows that Fenestrulina and Microporella Hincks, 1877a are not confamilial (Orr et al. 2019b). These authors also hypothesized that Fenestrulinidae should also include Adelascopora and Tenthrenulina. Family Gigantoporidae Bassler, 1935 Type genus: Gigantopora Ridley, 1881. Type species: Gigantopora lyncoides Ridley, 1881. Other genera: Barbadiopsis Winston & Woollacott, 2009, Cosciniopsis Canu & Bassler, 1927, Gephyrophora Busk, 1884, Stenopsella Bassler, 1952. Diagnosis: Colonies encrusting, uni- to multilaminar, multiserial; or erect, bilamellar arising from the encrusting base. Autozooids typically large, longitudinally oval, quadrangular to hexagonal. Lepralioid frontal shield pseudoporous; marginal areolar pores present. Orifice subcircular, with wide proximal sinus; small lateral condyles present. Secondary orifice variable, sometimes with elevated paired avicularia forming a bridge and spiramen. Oral spines absent. Ovicell peristomial; ectooecium membranous, endooecium pseudoporous. Avicularia adventitious, lateral oral, single or paired. Ancestrula autozooidal (Fig. 11.47 C, D). Distribution: Predominantly tropical waters. Remarks: There is some ongoing discussion on the status of some of the genera previously assigned to Gigantoporidae (e.g. Brown 1952, Gordon 1984, Cook 1985, Cook et al. 2018). The family comprises one fossil and five extant genera. Family Hippaliosinidae Winston, 2005 Type genus: Hippaliosina Canu, 1919. Type species: Escharella rostrigera Smitt, 1873. Other genera: None. Diagnosis: Colonies encrusting, uni- to multilaminar, multiserial; rarely erect, multiserial, unilamellar. Autozooids longitudinally quadrangular to hexagonal. Lepralioid frontal shield without frontal pseudopores; distinct marginal areolar pores present. Orifice clithridiate, with wide V-shaped sinus; lateral condyles present. Oral spines absent. Embryos incubated internally (presumably in brood sacs) in female zooids with larger dimorphic orifice. Avicularia present, adventitious, elongate or with setiform mandible, lateral-oral, single or paired, sometimes dimorphic. Vicarious avicularia rare. Ancestrula autozooidal (Fig. 11.48 A, B).



11.3 Systematics of Cheilostomata 

 391

Fig. 11.47: (A, B) Fenestrulinidae: Fenestrulina constellata Winston, Vieira & Woollacott, 2014. Off Cabo Frio, Rio de Janeiro, Brazil, W Atlantic Ocean. Museum of Comparative Zoology Cambridge, USA, 137464, holotype. (C, D) Gigantoporidae: Cosciniopsis violacea (Canu & Bassler, 1928a). Brazil, W Atlantic Ocean. Smithsonian National Museum of Natural History, Washington, DC, USNM 8556, syntype. Scale bars: A, C, 500 µm; B, D, 250 µm.

Distribution: Tropical to temperate waters. Remarks: Hippaliosinidae is monogeneric and distinguished from other schizoporelloid genera in having a non-pseudoporous frontal shield, clithridiate orifice, and dimorphic female zooids with larger orifice than in autozooids. Family Hippopodinidae Levinsen, 1909 Type genus: Hippopodina Levinsen, 1909. Type species: Lepralia feegeensis Busk, 1884. Other genera: Saevitella† Bobies, 1956, Thornelya Harmer, 1957, Trilochites Hayward, 1991. Diagnosis: Colonies encrusting, uni- to multilaminar, multiserial, rarely erect, multiserial, uni- to bilamellar. Autozooids longitudinally quadrangular to hexagonal. Lepralioid frontal shield, convex, uniformly pseudoporous;

marginal areolar pores present, small. Orifice subcircular to subquadrangular, with broad shallow sinus; lateral condyles and spines may be present. Embryos incubated in hyperstomial or subimmersed cleithral ovicells; ectooecium membranous, endooecium calcified, pseudoporous; maternal zooid sometimes with dimorphic orifice. Larva non-feeding. Avicularia present, adventitious, often lateral-oral. Ancestrula autozooidal (Fig. 11.48 C, D). Distribution: Cosmopolitan. Remarks: The composition of the family is problematic. There are some differences between Thornelya and other hippopodinid genera, including the type genus Hippopo­ dina, such as the presence of oral spines, two different types of avicularia and a cormidial orifice. The characteristics of avicularia, cormidial orifice and ovicell structure of Thornelya resemble those found in some species

392 

 11 Gymnolaemata, Cheilostomata

Fig. 11.48: (A, B) Hippaliosinidae: Hippaliosina imperfecta (Canu & Bassler, 1928a). Brazil, W Atlantic Ocean. Smithsonian National Museum of Natural History, Washington, DC, USNM 8563, syntype. (C, D) Hippopodinidae: Hippopodina adunca Tilbrook, 2006. Okinawa, Japan. East China Sea, intertidal. National Science Museum of Nature and Science Tsukuba, Japan, NSMT-Te 1137. Scale bars: A, D, 500 µm; B, 100 µm; C, 1 mm.

assigned to the lanceoporid genus Calyptotheca Harmer, 1957 (e.g. Calyptotheca ornatissima (Canu & Bassler, 1928a); see Almeida et al. 2017, 2018). Family Lacernidae Jullien, 1888 Type genus: Lacerna Jullien, 1888. Type species: Lacerna hosteensis Jullien, 1888. Other genera: Arthropoma Levinsen, 1909, Cheilonellopsis Gordon, 2014, Cribellopora Gautier, 1957, Cylindroporella Hincks, 1877a, Ministiaphila De Blauwe & Gordon, 2014, Nimba Jullien in Jullien & Calvet, 1903, Nimbella Jullien in Jullien & Calvet, 1903, Phonicosia Jullien, 1888, Ralepria Hayward, 1991, Rogicka Uttley & Bullivant, 1972, Schizo­ coryne Hayward & Winston, 2011, Vitrius Parker & Gordon, 1992, Woosukia Min, Seo, Grischenko, Lee & Gordon, 2017.

Diagnosis: Colonies encrusting, uni- to multilaminar, uni- to multiserial; rarely erect from an encrusting base, cylindrical, branching, non-articulated, with alternating rows of zooids around axis. Autozooids longitudinally pyriform to hexagonal. Lepralioid frontal shield, smooth, pseudopores lacking, sparse or abundant, these simple, stellate or occluded; areolar pores present. Orifice semicircular with wide poster and sinus, sometimes with condyles. Oral spines may be present. Embryos incubated in hyperstomial or peristomial ovicells, cleithral or (possibly) acleithral; ectooecium membranous, endooecium calcified, sometimes pseudoporous; maternal zooid with dimorphic orifice. Avicularia rarely present, adventitious or vicarious. Ancestrula autozooidal or with membranous frontal wall (Fig. 11.49 A, B).



11.3 Systematics of Cheilostomata 

 393

Fig. 11.49: (A, B) Lacernidae: Arthropoma harmelini Dick & Grischenko, 2017. Okinawa, Japan, East China Sea, intertidal. National Science Museum of Nature and Science, Tsukuba, Japan, NSMT-Te 1160. (C, D) Marcusadoreidae: Marcusadorea jamaicensis Vieira, Migotto & Winston, 2010. Rio Bueno, Jamaica, Carribbean Sea. Virginia Museum of Natural History, USA, 13359, holotype. Scale bars: A, B, D, 250 µm; C, 500 µm.

Distribution: Cosmopolitan. Remarks: Parker and Gordon (1992) distinguished Lacernidae from the Schizoporellidae based on the ovicell and frontal-shield characteristics. The ovicells mainly have a complete endooecium in Lacernidae, but a pseudoporous one (like those of Schizoporellidae) may be found in some species of Arthropoma, Rogicka, Vitrius, and Woosukia. There are also different frontal-shield characters in Lacernidae (smooth and pseudoporous) that need to be reassessed in the family. Family Marcusadoreidae Winston, Vieira, & Woollacott 2014 Type genus: Marcusadorea Vieira, Migotto, & Winston, 2010b. Type species: Marcusadorea jamaicensis Vieira, Migotto & Winston, 2010b.

Other genera: None. Diagnosis: Colony encrusting, uni- to multilamellar, multiserial, sometimes erect with tubular branches. Autozooids large, longitudinally rectangular to hexagonal; frontal shield lepralioid, convex, irregularly pseudoporous and with marginal pores. Primary orifice with semicircular anter and well-developed sinus, obscured by well-developed peristome; condyles present. Spines, lyrula, or sinus absent. Peristomial avicularium occasional, other avicularia absent. Ovicells subimmersed, hyperstomial or peristomial, with calcified pseudoporous endooecium and membranous ectooecium Ancestrula autozooidal. Kenozooids absent (Fig. 11.49 C, D). Distribution: Tropical waters of the western Atlantic Ocean and the southwestern Pacific Ocean.

394 

 11 Gymnolaemata, Cheilostomata

Remarks: Marcusadoreidae is monogeneric. Some morphological characteristics are shared with Marcusadoreidae and both Schizoporelloidea and Smittinoidea (Vieira et al. 2010c), but further molecular work is needed to elucidate the phylogenetic relationships among these taxa. Family Margarettidae Harmer, 1957 Type genus: Margaretta Gray, 1843. Type species: Cellaria barbata Lamarck, 1816. Other genera: None. Diagnosis: Colony erect, cellariform, jointed, elongate, branched, arising from an erect ancestrula; anchored by numerous rhizoids. Internodes cylindrical, sometimes

curved, composed of alternating series of tubular zooids forming whorls along internodes. Autozooids with lepralioid frontal shield, densely pseudoporous, with ascopore opening proximally to the secondary orifice. Primary orifice subcircular, obscured by long tubular peristome. Oral spines absent. No avicularia. A pair of long cuticular “bristles” (possibly kenozooidal) flanks the ascopore in one species. Ovicells peristomial, associated with prominent curved peristome (Fig. 11.50 A, B). Distribution: Tropical to temperate waters. Remarks: Margarettidae includes only Margaretta and Tubucella Canu & Bassler, 1917, the latter wholly fossil. Fertile zooids of Margaretta have a functional polypide smaller than other zooids (Ostrovsky 2013).

Fig. 11.50: (A, B) Margarettidae: Margaretta tenuis (Harmer, 1957). Al-Lith, Saudi Arabia, coral reef flat, Red Sea. Senckenberg Forschungsinstitute und Naturmuseen, Sektion Marine Evertebraten III (Bryozoologie), SMF 40068. Images provided by Christoph Neu and Joachim Scholz. (C, D) Microporellidae: Microporella trigonellata Suwa & Mawatari, 1998. Akkeshi Bay, Hokkaido, Japan, NW Pacific, intertidal. Natural History Museum, London, 2006.2.27.96 and 2006.2.27.95. Scale bars: A, 1 mm; B, C, D, 250 µm.



Family Mawatariidae Gordon, 1990 Type genus: Mawatarius Gordon, 1990. Type species: Prostomaria inexpectabilis Gordon, 1985. Other genera: None. Diagnosis: Colony erect, uniserial, dichotomously branching, unjointed. Autozooids with lepralioid frontal shield, pseudopores lacking, sparse or moderate. Primary orifice sunken, with secondary orifice forming a distinct proximal pseudosinus. Lyrula well developed or just a small denticle. Oral spines absent. Avicularia absent. Abfrontal (basal) wall with sparse areolar-septular pores. Ovicells peristomial, ectooecium membranous, endooecium calcified, non-pseudoporous. Ancestrula autozooidal, erect, anchored by a short cuticular portion. Distribution: Tropical to temperate waters of the southwestern Atlantic Ocean and the southwestern Pacific Ocean. Remarks: This family includes a single genus, Mawatarius, including three Recent species from deep waters. Family Microporellidae Hincks, 1879 Type genus: Microporella Hincks, 1877a. Type species: Eschara ciliata Pallas, 1766. Other genera: Calloporina† Neviani, 1896b, Chronocer­ astes† Gordon, 1989c, Diporula Hincks, 1879, Flustramor­ pha Gray, 1872. Diagnosis: Colonies encrusting, multiserial, uni- to multilamellar, or erect, foliaceous, uni- to bilamellar. Autozooids subrectangular to hexagonal, convex, separated by raised walls. Lepralioid frontal shield with suboral ascopore, with or without pseudopores; marginal areolar pores present. Orifice semicircular, sometimes with indentations, denticles or condyles; mucro may be present. Oral spines present. Avicularia adventitious, mainly placed near the orifice, sometimes with long mandible (vibracula). Ovicells hyperstomial acleithral or endozooidal cleithral, ectooecium totally or partially membranous, endooecium calcified, usually pseudoporous. Larva non-feeding. Ancestrula autozooidal or tatiform (Fig. 11.50 C, D). Distribution: Cosmopolitan. Remarks: Fenestrulina and Microporella both have an ascopore, but molecular-genetic analysis gives evidence that they are unrelated and thus no longer confamilial. For this reason Orr et al. (2019b) resurrected Fenestrulinidae Jullien, 1888 to accommodate Fenestrulina and related genera. Di Martino et al. (2020) note that Diporula and Flustramorpha have so many characters overlapping with Microporella that they are probably synonymous. Confirmation requires molecular-genetic analysis. Other genera attributed to Microporellidae require similar analysis.

11.3 Systematics of Cheilostomata 

 395

Family Myriaporidae Gray, 1841 Type genus: Myriapora de Blainville, 1830. Type species: Millepora truncata Pallas, 1766. Other genera: Leieschara Sars, 1863, Myriozoella Levinsen, 1909. Diagnosis: Colony encrusting, unilamellar to multilamellar, multiserial; or erect, cylindrical, branching, unjointed. Autozooids polygonal often with indistinct boundaries; frontal shield lepralioid, pseudoporous with indistinct marginal areolae. Primary orifice may be sunken, with distinct sinus. Oral spines absent. Avicularia adventitious. Ovicells endozooidal, cleithral, with membranous or calcified ectooecium covered by secondary calcification, and dimorphic orifice of fertile zooid. Ancestrula zooidal (twinned) or tatiform. Kenozooids absent (Fig. 11.51 A, B). Distribution: Temperate to polar waters. Remarks: Truncularia Wiegmann & Ruthe, 1832 is regarded as an objective synonym of Myriapora (Truncu­ laria teres Wiegmann & Ruthe, 1832 = Millepora truncata Pallas, 1766). The genus Leieschara may be a subjective synonym of Myriapora. Family Pacificincolidae Liu & Liu, 1999 Type genus: Pacificincola Liu & Liu, 1999. Type species: Mucronella perforata Okada & Mawatari, 1937. Other genera: Burdwoodipora López Gappa, Liuzzi & Zelaya, 2017, Primavelans De Blauwe, 2006. Diagnosis: Colonies encrusting, unilamellar, sometimes with erect bilamellar lobes. Frontal shield lepralioid, pseudoporous. Orifice subcircular, with wide shallow sinus and proximo-lateral condyles; oral spines absent. Peristome may form a complete raised rim, with raised umbo proximally and avicularium or kenozooid placed between orifice and umbo. Avicularia adventitious, lateral-oral or placed on suboral umbo. Ovicells hyperstomial, cleithral, ectooecium membranous, endooecium calcified, non-pseudoporous. Ancestrula autozooidal or tatiform (Fig. 11.51 C, D). Distribution: Cosmopolitan. Remarks: Pacificincolidae has been revised by De Blauwe (2006), who added a new genus to the family. Another new genus was recently added by López Gappa et al. (2017). Family Petraliidae Levinsen, 1909 Type genus: Petralia MacGillivray, 1869. Type species: Petralia undata MacGillivray, 1869. Other genera: Mobunula Gordon, 1989c, Mucropetraliella Stach, 1936, Petraliella Canu & Bassler, 1927, Riscodopa Gordon, 1989c, Sinupetraliella Stach, 1936, Utinga Marcus, 1949.

396 

 11 Gymnolaemata, Cheilostomata

Fig. 11.51: (A, B) Myriaporidae: Myriapora subgracilis (d’Orbigny, 1852). Svalbard, Norway, Arctic Sea. Perm State National Research University, Kluge Collection, 810.28. (C, D) Pacificincolidae: Pacificincola insculpta (Hincks, 1882b). Ketchikan, Alaska, USA, NE Pacific, intertidal. Scale bars: A, 500 µm; B, C, D, 250 µm.

Diagnosis: Colonies encrusting, unilamellar to multilamellar, to semierect or tubular, occasionally discoidal. Autozooids with pseudoporous frontal shield and distinct marginal areolar pores. Orifice wide, sometimes with indentations, denticles, condyles or proximal sinus; mucro may be present, sometimes aviculiferous. All avicularia adventitious, variable in position and shape. Basal walls usually with numerous septular pores, or radicular chambers, sometimes in groups, or single, often very large, occupying much of the basal wall, producing anchoring rootlets. Ovicells hyperstomial or subimmersed, sometimes reminiscent of endozooidal ovicell, acleithral, cleithral or non-cleithral; endooecium pseudoporous, ectooecium membranous (Fig. 11.52 A, B). Distribution: Tropical to temperate waters. Remarks: Pending formal revision, the morphology of Utinga suggests it might be a junior synonym of Petraliella.

Family Phoceanidae Vigneaux, 1949 Type genus: Phoceana Jullien in Jullien & Calvet, 1903. Type species: Phoceana columnaris Jullien in Jullien & Calvet, 1903. Other genera: Sertulipora Harmelin & d’Hondt, 1992. Diagnosis: Colonies encrusting, uniserial, dichotomously branching, or erect, with longitudinal rows of autozooids. Autozooids with pseudoporous or smooth frontal wall, with distinct marginal pores. Orifice with semicircular anter, wide poster and paired condyles; secondary orifice well developed, tubular. Avicularia absent. Ovicells peristomial, not closed by the operculum, imperforate. Ancestrula autozooidal (Fig. 11.52 C, D). Distribution: Tropical to temperate waters of the North Atlantic Ocean. Remarks: Phoceanidae includes only two genera with four species that are all found in the Mediterranean and North Atlantic realm.



11.3 Systematics of Cheilostomata 

 397

Fig. 11.52: (A, B) Petraliidae: Utinga castanea (Busk, 1884). Saint Peter and Saint Paul Archipelago, Brazil, W Atlantic Ocean. Universidade Federal de Pernambuco, unregistered. (C, D) Phoceanidae: Phoceana tubulifera (Heller, 1867). Anafi, Greece, Aegean Sea. Senckenberg Forschungsinstitute und Naturmuseen, Sektion Marine Evertebraten III (Bryozoologie), unregistered. Scale bars: A, 500 µm; B, D, 250 µm; C, 1 mm.

Family Phorioppniidae Gordon & d’Hondt, 1997 Type genus: Phorioppnia Gordon & d’Hondt, 1997. Type species: Phorioppnia cookae Gordon & d’Hondt, 1997. Other genera: Oppiphorina Gordon & d’Hondt, 1997, Quadriscutella Bock & Cook, 1993. Diagnosis: Colonies erect from an encrusting base, branching, or jointed and basally rooted. Autozooids with pseudoporous lepralioid frontal wall, separated by a raised marginal edge. Orifice almost circular, lateral condyles may be present. Oral spines absent. Avicularia absent. Embryos incubated in subimmersed or endozooidal cleithral ovicells; ectooecium membranous, endooecium calcified, pseudoporous; maternal zooids with dimorphic orifice. Distribution: Tropical to temperate waters of the southwestern Pacific Ocean and the southeastern Indian Ocean. Remarks: The family is known from New Caledonia, New Zealand, and Australia.

Family Porinidae d’Orbigny, 1852 Type genus: Porina d’Orbigny, 1852. Type species: Eschara gracilis Lamarck, 1816. Other genera: Haswelliporina Gordon & d’Hondt, 1997, Mosaicoporina Gordon & d’Hondt, 1997, Semihaswellia Canu & Bassler, 1917. Diagnosis: Colonies erect, from a small encrusting base or erect ancestrula, rod shaped, flattened or branching; rarely repent, uniserial and supported just above the substratum by short wall extensions. Autozooidal boundaries indistinct unless uniserial; frontal shield pseudoporous, often with secondary calcification. Orifice with subcircular anter and broad or narrow sinus, obscured by secondary orifice; lateral condyles may be present. Oral spines absent. Secondary orifice forming a long tube around the whole orifice. Tubular spiramen present. Avicularia adventitious, variable in shape and position. Ovicells

398 

 11 Gymnolaemata, Cheilostomata

Fig. 11.53: (A, B) Porinidae: Porina gracilis (Lamarck, 1816). Encounter Bay, South Australia, Australia, Indian Ocean. National Institute of Water & Atmospheric Research, Wellington, unregistered. © http://bryozoa.net. (C, D) Robertsonidridae: Robertsonidra sp. Vema seamount, SE Atlantic. University of Vienna, unregistered. Scale bars: A, C, 500 µm; B, D, 250 µm.

endozooidal, non-cleithral; ooecium embedded in pseudoporous frontal shield of distal zooid (Fig. 11.53 A, B). Distribution: Tropical to temperate waters. Remarks: The family is widely reported in the fossil record, but all Cretaceous taxa attributed to the family belong elsewhere (Gordon 2002). Some Recent representatives are known from deep water. The type genus, Porina d’Orbigny, 1852 remained without a designated type species until Lang (1917) selected Eschara gracilis Lamarck, 1816. Detailed comments on the nomenclatorial and historical changes in the definition of the family were provided by Gordon and d’Hondt (1997) and Gordon (2002). Family Robertsonidridae Rosso in Rosso et al., 2010 Type genus: Robertsonidra Osburn, 1952. Type species: Schizoporella oligopus Robertson, 1908.

Other genera: Bertorsonidra Rosso, Sciuto & Sinagra, 2010, Jodoella Yang, Seo & Gordon, 2018a. Diagnosis: Colonies encrusting, multilamellar. Autozooids with pseudoporous or smooth frontal shield, with distinct marginal pores. Orifice with semicircular anter and proximal sinus; lateral condyles present or absent; secondary orifice slightly raised, sometimes inconspicuous; oral spines present or absent. Avicularia adventitious, variable in shape. Embryos incubated in hyperstomial or subimmersed (presumably cleithral) ovicells with membranous ectooecium and pseudoporous endooecium; female zooids with dimorphic orifice. Vertical walls with uniporous or multiporous septula (Fig. 11.53 C, D). Distribution: Tropical to temperate waters. Remarks: Jodoella is distinguished from the other genera of this family by having dimorphic orifices.



Family Stomachetosellidae† Canu & Bassler, 1917 Type genus: Stomachetosella† Canu & Bassler, 1917. Type species: Stomachetosella crassicollis† Canu & Bassler, 1917. Other genera: Junerossia Dick, Tilbrook & Mawatari, 2006, Tremoschizodina Duvergier, 1920. Diagnosis: Colonies encrusting, multiserial. Autozooids with a lepralioid frontal shield with some pseudopores. Orifice with or without condyles; oral spines absent. Primary orifice mostly surrounded by peristome. Avicularia present in some species. Ovicells hyperstomial, subimmersed, endozooidal or immersed (then cleithral), maternal zooids often with dimorphic orifice; ectooecium calcified, with or without pseudopores, often covered by secondary calcification, opens into a secondary orifice; a collar surrounds the orifice in ovicellate zooids. Ancestrula autozooidal (Fig. 11.54 A, B).

11.3 Systematics of Cheilostomata 

 399

Distribution: Tropical to polar waters of the Northern Hemisphere. Remarks: It is possible that Stomachetosellidae has no Recent representatives. The sole species of Junerossia has an umbonuloid component in the frontal shield near the orifice and may be unrelated, while Tremoschizodina crassa Canu & Bassler, 1929 from the Philippines has dimorphic orifices and large hyperstomial ovicells, in contradistinction to the type species with endozooidal ovicells. Family Tetraplariidae Harmer, 1957 Type genus: Tetraplaria Tenison-Woods, 1879. Type species: Tetraplaria australis Tenison-Woods, 1879. Other genera: None. Diagnosis: Colonies erect from a small encrusting base, branching, jointed, with quadriserial branches.

Fig. 11.54: (A, B) Stomachetosellidae: Stomachetosella decorata Grischenko, Dick & Mawatari, 2007. Akkeshi Bay, Hokkaido, Japan, NW Pacific, intertidal. Natural History Museum, London, 2006.2.27.89. (C, D) Teuchoporidae: Lagenicella sp. Mexico. unregistered. Scale bars: A, B, D, 250 µm; C, 500 µm.

400 

 11 Gymnolaemata, Cheilostomata

Autozooids paired back-to-back in four longitudinal series along the internode. Autozooids with pseudoporous frontal shield. Orifice subcircular to oval, with or without sinus; condyles may be present; oral spines absent. Suboral avicularia may be present. Embryos incubated internally (presumably in brood sacs) in enlarged dimorphic zooids or in endozooidal cleithral ovicells; ectooecium membranous, endooecium with few pseudopores. Distribution: Tropical waters. Remarks: Tetraplaria is a junior synonym of Onchopora Busk, 1855, considered a nomen oblitum (Gordon 1989c). According to Harmer (1957), another four genera were synonymized with Tetraplaria: Arborella Osburn, 1914, Bige­ mellaria MacGillivray, 1895, Diploecium Kirkpatrick, 1888, and Pollaploecium Maplestone, 1909b. These genera, however, require review since there are ranges of reproductive patterns (e.g. hyperstomial ovicells and internal brooding) suggesting that the genus Tetraplaria needs splitting (Cook et al. 2018). Family Teuchoporidae† Neviani, 1896b Type genus: Teuchopora† Neviani, 1896b. Type species: Alecto castrocarensis† Manzoni, 1875. Other genera: Lagenicella Cheetham & Sandberg, 1964. Diagnosis: Colonies encrusting. Autozooids with well-calcified lepralioid frontal shield, evenly pseudoporous. Primary orifice obscured by peristome, with broad sinus, condyles present or absent; no oral spines. Secondary orifice well developed; one or more adventitious avicularia may be present in the peristome. No other avicularia. Ovicells peristomial, ectooecium membranous entirely or partially, ectooecium pseudoporous. Ancestrula autozooidal. Vertical walls with basal pore-chambers or multiporous septula (Fig. 11.54 C, D). Distribution: Tropical to temperate waters. Remarks: The above diagnosis is based on both included genera. Soule et  al. (1995) argued against the validity of the family name Teuchoporidae, preferring Phylactellidae Canu & Bassler, 1917 instead. Lepralia labrosa Busk, 1854, type species of Phylactella Hincks, 1879, has a lyrula; thus, this genus is currently better assigned to Smittinidae Levinsen, 1909 (Rosso 2004). Using light microscopy and a drawing, Poluzzi (1977) illustrated the lectotype and topotypic material of T. cas­ trocarensis from the Pliocene of northern Italy and its characters provide the proper basis for characterizing the family – the colony is bi- to pluriserial, zooids are evenly pseudoporous, the smooth-walled peristome has a denticle on its inner proximal face, and the primary orifice is almost transversely elliptical and lacks condyles. The ooecium appears to have a smooth surface (Poluzzi 1977, pl. 1, fig. 7) and opens into the peristome. The line drawing

that shows putative basal pore-chambers suggests they may simply comprise slightly buttressed recesses. Poluzzi’s (1977) material would benefit from SEM study. Family Vicidae Gordon, 1988 Type genus: Vix Gordon, 1988. Type species: Bifaxaria vagans Thornely, 1912. Other genera: Cyclostomella Ortmann, 1890. Diagnosis: Colonies erect, basally rooted, branching, quadriserial; joints may be present. Zooids with wellcalcified lepralioid frontal shield, with some pseudopores. Primary orifice obscured by peristome, with broad poster and small proximal condyles; oral spines absent. Secondary orifice well developed and continuous with frontal zooidal surface, with oral avicularium. Oral avicularia without crossbar. Other avicularia absent. Ovicells hyperstomial or missing (Fig. 11.55 A, B). Distribution: Tropical waters of the Indian and Pacific oceans. Remarks: Unfortunately, there are no recent descriptions of Cyclostomella Ortmann, 1890, but the genus was tentatively assigned to Vicidae owing to morphological similarities with V. vagans (see Harmer 1957, Gordon 1988). No ovicells were described for Vix vagans, while ovicells are hyperstomial with possibly pseudoporous ooecium in fossil material from the Miocene assigned to Vix (Di Martino et al. 2017).

11.3.7.23 Superfamily Siphonicytaroidea Harmer, 1957 Family Siphonicytaridae Harmer, 1957 Type genus: Siphonicytara Busk, 1884. Type species: Siphonicytara serrulata Busk, 1884. Other genera: None. Diagnosis: Colonies erect, cylindrical to flattened, branching, often bilamellar, anchored by few to many rootlets. Zooids with a lepralioid frontal shield that is characteristically divided into sectors by low, thin ridges; areolar-septular pores few or many, small or considerably expanded by merging of pores through secondary calcification. Primary orifice subcircular to oval, becoming deeply concealed at the bottom of a long peristomial shaft; no lyrula or condyles; no oral spines. Ascopore present, typically some distance from the peristomial orifice, opening into zooidal interior by long narrow tube. Avicularia adventitious, arising from marginal septular pores, with complete pivot bar. Ovicells presumably peristomial. Distribution: Tropical waters of the eastern Indian Ocean and the western Pacific Ocean. Remarks: The type species of Siphonicytara was redescribed by Harmer (1957) and Bock and Cook (2001). The latter



11.3 Systematics of Cheilostomata 

 401

Fig. 11.55: (A, B) Vicidae: Vix vagans (Thornely, 1912). North Male Atoll, Maldive Islands, Indian Ocean. University of Vienna, unregistered. (C, D) Smittinidae: Smittina nitidissima (Hincks, 1880a). Okinawa, Japan, East China Sea, intertidal. National Museum of Nature and Science, Tsukuba, NSMT-Te 1119 (MIN-2). Scale bars: A, C, 500 µm; B, 50 µm; D, 250 µm.

authors discussed the fossil genus Tubitrabecularia Bassler, 1934, attributed to Siphonicytaridae, and considered it unrecognizable. Gordon and Taylor (2015) described an Early Eocene species of Siphonicytara from New Zealand that had a small umbonuloid component in the frontal shield, inviting comparison with the Cretaceous–Oligocene tessaradomid genus Beisselina Canu, 1913, which is characterized by an umbonuloid frontal shield with a spiramen. Structurally, the zooids of Siphonicytara are fairly similar to those of Beis­ selina, and it is possible that Beisselina, or something very similar, included the ancestor to Siphonicytara.

11.3.7.24 Superfamily Smittinoidea Levinsen, 1909 Family Smittinidae Levinsen, 1909 Type genus: Smittina Norman, 1903b. Type species: Lepralia landsborovii Johnston, 1847.

Other genera: Amynaskolia Figuerola, Gordon & Cristobo, 2018, Aspericreta Hayward & Thorpe, 1990, Breo­ ganipora Souto, Berning & Ostrovsky, 2016, Dakariella Moyano G., 1966, Dengordonia Soule, Soule & Chaney, 1995, Dittomesia Gordon, 1989c, Hemismittoidea Soule & Soule, 1973, Parasmittina Osburn, 1952, Pemmatoporella Hayward & Taylor, 1984, Phylactella Hincks, 1879, Platychelyna Hayward & Thorpe, 1990, Pleurocodonellina Soule & Soule, 1973, Prenantia Gautier, 1962, Pseudoflus­ tra Bidenkap, 1897, Raymondcia Soule, Soule & Chaney, 1995, Smittinella Canu & Bassler in Bassler, 1934, Smit­ toidea Osburn, 1952, Thrypticocirrus Hayward & Thorpe, 1988a, Tracheloptyx Hayward, 1993 (Fig. 11.55 C, D). Diagnosis: Colonies encrusting, uni- to multilamellar, or erect. Zooids with lepralioid or umbonuloid frontal shield, with or without pseudopores. Zooids may have secondary calcification in later ontogeny. Orifice subcircular to oval, often with proximal lyrula or narrow sinus; condyles

402 

 11 Gymnolaemata, Cheilostomata

typically present and oral spines often present, especially in young zooids at colony margins. Avicularia present or absent, variable in shape, adventitious, vicarious or interzooidal. Embryos incubated in hyperstomial or subimmersed, acleithral, cleithral, or subcleithral ovicells, ooecium often overgrown and deeply embedded in secondary calcification, reminiscent of endozooidal ovicells; both ooecial walls calcified, ectooecium pseudoporous. Larva non-feeding. Distribution: Cosmopolitan. Remarks: All smittinids have ooecia with two calcified walls with pseudoporous ectooecium. Genera are differentiated by orifice characters (such as presence/absence of lyrula, sinus and condyles), the morphology of the frontal shield (umbonuloid or lepralioid, i.e. smooth or pseudoporous), and avicularia origins. Gordon (1984) questioned the validity of some genera, such as Hemi­ smittoidea, highlighting the use of some features to distinguish genera may be questionable owing to inconsistency of expression. Farias et al. (2020) have suggested redefinition of four genera – Smittina, Smittoidea, Parasmittina, and Hemismittoidea – resulting in some new combinations for different taxa. Further integrative studies should clarify the assignment of taxa to this family. Family Bitectiporidae† MacGillivray, 1895 Type genus: Bitectipora† MacGillivray, 1895. Type species: Bitectipora lineata† MacGillivray, 1895. Other genera: Caesiopora Gordon, 2014, Schizomavella (Calvetomavella) Reverter-Gil, Berning & Souto, 2015, Cri­ bella Jullien in Jullien & Calvet, 1903, Galiciapora Souto, Berning & Ostrovsky, 2016, Hippomonavella Canu & Bassler in Bassler, 1934, Hippoporina Neviani, 1895, Hippothyris Osburn, 1952, Kermadecazoon Tilbrook, 2006, Metroperiella Canu & Bassler, 1917, Neodakaria Soule, Soule & Chaney, 1995, Nigrapercula Tilbrook, 2006, Parkermavella Gordon & d’Hondt, 1997, Pentapora Fischer von Waldheim, 1807, Plac­ idoporella Souto, Berning & Ostrovsky, 2016, Schizomavella Canu & Bassler, 1917, Schizosmittina Vigneaux, 1949, Suhius Min, Seo, Grischenko & Gordon, 2017. Diagnosis: Colonies encrusting, multiserial, uni- to multilamellar, or erect, foliaceous, uni- or bilamellar. Zooids with lepralioid frontal shield with or without pseudopores. Zooids may have secondary calcification in later ontogeny. Orifice with subcircular to oval anter and broad to narrow poster or sinus, but no lyrula; condyles and oral spines often present. Avicularia present or absent, variable in shape, adventitious, vicarious or interzooidal. Ovicells hyperstomial to subimmersed, acleithral, cleithral and, possibly, subcleithral, ooecium sometimes overgrown by secondary calcification; both ooecial walls calcified, ectooecium pseudoporous. Larva non-feeding (Fig. 11.56 A, B).

Distribution: Cosmopolitan. Remarks: Overall, bitectiporids can be said to resemble smittinids but lack an orificial lyrula. The family is based on a species from the Miocene of Victoria, Australia. In a revision of the type material, Gordon (1994b) resurrected the family and found that many extant taxa may be accommodated into this family. Gordon (1994b) also found that Bitectiporidae is a senior synonym of Hippoporinidae Brown, 1952. Family Lanceoporidae Harmer, 1957 (1927) Type genus: Lanceopora d’Orbigny, 1852. Type species: Lanceopora elegans d’Orbigny, 1852. Other genera: Calyptotheca Harmer, 1957, Emballotheca Levinsen, 1909, Stephanotheca Reverter-Gil, Souto & Fernández-Pulpeiro, 2012. Diagnosis: Colonies variable in shape, encrusting, multiserial, uni- to multilamellar, or erect, foliaceous, uni- or bilamellar, stalked or discoidal. Zooidal frontal shield lepralioid, pseudoporous, with marginal areolar-septular pores generally resembling pseudopores. Zooids often with secondary calcification in later ontogeny. Orifice with subcircular to oval anter and either a broad poster or smaller sinus; condyles present. Spines absent. Avicularia present, variable in shape, adventitious, vicarious or interzooidal. Embryos brooded in hyperstomial or subimmersed ovicells, cleithral or subcleithral; ooecia covered by secondary calcification with non-calcified areas corresponding to pseudopores in ectooecium (Fig. 11.56 C, D). Distribution: Cosmopolitan. Remarks: The family is speciose in tropical and subtropical waters, mostly in the Indo-Pacific and Mediterranean. Cook et al. (2018) indicated that the name Lanceoporidae may be maintained under prevailing usage over Parmulariidae Canu & Bassler, 1927. However, based on Recommendation 40A (ICZN 1999) the original author and date should be cited followed by the date of its priority enclosed in parentheses, thus Lanceoporidae Harmer, 1957 (1927) is the correct citation. Family Powellithecidae Di Martino, Taylor, Gordon, & Liow, 2016 Type genus: Powellitheca Di Martino, Taylor, Gordon & Liow, 2016. Type species: Powellitheca terranovae Di Martino, Taylor, Gordon & Liow, 2016. Other genera: None. Diagnosis: Colonies encrusting, multiserial, with zooids in well-defined longitudinal rows. Autozooids with convex, lepralioid frontal shield, regularly and evenly perforated. Vertical walls with multiporous septula. Orifice dimorphic, wider in maternal zooids. Primary



11.3 Systematics of Cheilostomata 

 403

Fig. 11.56: (A, B) Bitectiporidae: Hippoporina lacrimosa Cook, 1964. Maceió, Alagoas, Brazil, W Atlantic Ocean. Universidade Federal de Pernambuco, unregistered. (C, D) Lanceoporidae: Lanceopora elegans d’Orbigny, 1851. Strait of Malacca, Malaysia, Indian Ocean. Muséum National d’Histoire Naturelle, Paris, F.R64250, syntype. Scale bars: A, C, 500 µm; B, 100 µm; D, 250 µm.

orifice with weakly convex proximal rim; condyles small. Suboral umbo and lyrula present or absent; oral spines lacking. Avicularia present or absent. Ovicells hyperstomial, cleithral; ectooecium membranous, endooecium calcified, pseudoporous; female zooids with dimorphic orifice. Ancestrula minimally calcified, with a rim of granular cryptocystal calcification. Distribution: Southwestern Pacific Ocean off New Zealand. Remarks: This family is monogeneric and contains only three species, one of which is from the Pliocene–Pleistocene of New Zealand. Powellithecids differ from other families with similar ovicell structure in having dimorphic orifices lacking a sinus and in having no ascopore. Family Watersiporidae Vigneaux, 1949 Type genus: Watersipora Neviani, 1896b. Type species: Lepralia cucullata Busk, 1854.

Other genera: Terwasipora Reverter-Gil & Souto, 2019, Uscia Banta, 1969b, Veleroa Osburn, 1952. Diagnosis: Colonies encrusting, multiserial, uni- to multilamellar, or erect, foliaceous, uni- or bilamellar. Colony mostly with pigmented epitheca. Autozooids subrectangular to hexagonal, separated by raised walls. Lepralioid frontal shield with numerous rounded pseudopores; lateral oral intrazooidal septula may be present proximolateral to the orifice; intrazooidal septula sometimes present at proximal corners of frontal shield. Orifice monomorphic or dimorphic, subcircular to oval; poster sometimes with well-defined proximal sinus; condyles present. Operculum dark colored, often with a central band demarcated by sclerites; lucidae often present. Spines absent. Avicularia absent. Embryos incubated in internal brood sacs, possibly in dimorphic zooids in Uscia. Multiporous mural pore-plates in distolateral and transverse distal walls. Larva non-feeding. Ancestrula schizoporelloid,

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single, smaller than autozooids, often obscured in later astogeny (Fig. 11.57 A, B). Distribution: Tropical to temperate waters. Remarks: Partly owing to misidentification of the type species of Watersipora Neviani, 1896b, as well as the paucity of morphological characters and consequent similarity of species, there has been much confusion in the literature surrounding taxon identities. Vieira et al. (2014a) clarified and stabilized the type and most other species of Watersipora, and Reverter-Gil & Souto, 2019 created a new genus, Terwasipora, for one other species previously attributed to Watersipora. The monotypic genus Veleroa Osburn, 1952 still requires evaluation; it resembles the lacernid genus Woosukia Min, Seo, Grischenko, Lee, & Gordon, 2017, but ovicells have yet to be discovered in Veleroa, which would clarify the situation. Ovicells are absent in all known Recent species of Watersiporidae, but one fossil species with immersed ovicells and dimorphic orifices has been attributed to Watersipora (Berning 2006).

11.3.7.25 Superfamily Urceoliporoidea Bassler, 1936 Family Urceoliporidae Bassler, 1936 Type genus: Urceolipora MacGillivray, 1881. Type species: Urceolipora nana MacGillivray, 1881. Other genera: Reciprocus Gordon, 1988. Diagnosis: Colonies erect, branching, biserial, segments jointed or unjointed, basally rooted. Zooids arranged back to back, with lepralioid frontal shield that can be divided by a thin ridge into two sectors; frontally imperforate or with only a cluster of pores near the orifice. Orifice monomorphic or dimorphic, with a sinus. No peristome, but oral processes may be present. Spines and avicularia absent. Embryos incubated in endozooidal cleithral ovicells; ooecium with membranous ectooecium and pseudoporous endooecium, or in internal brood sacs in swollen dimorphic zooids (Reciprocus). Distribution: Australia and New Zealand. Remarks: Urceoliporidae is a small Australasian family comprising only three Recent and one fossil species, the latter, Cureolipora miocenica Gordon, 2000, occurring in the Early Miocene of New Zealand. All living colonies are erect with biserial internodes in which each zooid communicates dorsally with three other zooids. Family Prostomariidae† MacGillivray, 1895 Type genus: Prostomaria† MacGillivray, 1895. Type species: Prostomaria gibbericollis† MacGillivray, 1895. Other genera: None.

Diagnosis: Colonies erect, biserial, segmented, basally rooted, with zooids arranged back to back. Autozooids with lepralioid frontal shield, evenly perforate. Orifice monomorphic, without sinus. Peristome well developed, obscuring the orifice. Spines and avicularia absent. Ovicells unknown. Distribution: Tropical of the Atlantic Ocean and Australia. Remarks: In a revision of Prostomariidae, Gordon (1990) argued the family to be monospecific, the only representative being the type species from the Miocene of Victoria, Australia. A putative living species, Prostomaria inexpect­ abilis Gordon, 1985, was later reassigned to Mawatarius Gordon, 1990. Prostomaria cyclostomata d’Hondt & Schopf, 1985, is also highly unlikely to be related to Pros­ tomaria; based on its external morphology, Gordon (1990) suggested that the frontal shield may be umbonuloid rather than lepralioid. The species may require a new genus; its family relationships also are presently uncertain. It is currently the only nominally extant prostomariid.

11.3.8 Suborder Skylonina Viskova in Viskova & Morozova, 1988 Remarks: “Skyloniida” was originally proposed and described by Viskova in Viskova and Morozova (1988) as a new order. Viskova (1992) assigned the authorship of the suborder Skylonina to Sandberg (1963), which is unjustified as Sandberg (1963) proposed only the family-group name. Later authors have not used Skylonina and the name is reintroduced here to incorporate the problematic putative cheilostome families Skyloniidae Sandberg, 1963, Bicorniferidae Keij, 1977 and Fusicellariidae Canu, 1900. 11.3.8.1 Superfamily Skylonoidea Sandberg, 1963 Family Bicorniferidae† Keij, 1977 Type genus: Bicornifera† Lindenberg, 1965. Type species: Bicornifera alpina† Lindenberg, 1965. Other genera: Voorthuyseniella† Szczechura, 1969. Diagnosis: Colonies erect, fusiform; or encrusting, unilamellar and multiserial. Autozooids arranged in up to seven regular series, quadrate to longitudinally hexagonal. Frontal aperture roundish, elliptical or slit-like, in a central position. Frontal wall calcification smooth. Kenozooid-like polymorphs may be present at the top and at the bottom of a colony. Ancestrula not reported. Spines, avicularia and ovicells unknown (Fig. 11.57 C, D). Distribution: Tropical waters of the South China Sea and the Gulf of Mexico.

Literature 

 405

Fig. 11.57: (A, B) Watersiporidae: Watersipora cucullata (Busk, 1854). Greece, Aegean Sea. Natural History Museum, London, 2012.6.30.1, paralectotype. (C, D) Bicorniferidae: Skylonia bermudezi Keij, 1973. Jabaco, Cuba, Eocene. Natural History Museum, London, D51856, paratype. Scale bars: A, C, 500 µm; B, 250 µm; D, 100 µm.

Remarks: Keij (1970) described two Recent species of Voorthuyseniella Szczechura, 1969 from the Gulf of Mexico and the South China Sea. Although the attribution of V. borneensis Keij, 1970 and V. occidentalis Keij, 1970 to Bicorniferidae is not challenged here, based on the figures given in Keij (1970), both require re-examination and careful study. No other Recent bicorniferids have since been reported, which is striking considering the large number of colonies of V. occidentalis reported by Keij (1970). The occurrence of Recent bicorniferids is therefore doubtful. Unplaced genera Acanthodesiomorpha d’Hondt, 1981, Bountyella Gordon, 2014, Brettia Dyster, 1858, Ogivalia Jullien, 1882a, Para­ stichopora Cook & Chimonides, 1981a

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Winston, J.E., Vieira, L.M. & Woollacott, R.M. (2014): Scientific results of the Hassler Expedition. Bryozoa. No. 2. Brazil. Bull Mus Comp Zool 161(5): 139–239. DOI: 10.3099/MCZ14.1. Wolff, T. (1959): The hadal community, an introduction. Deep Sea Res 6: 95–124. Wyer, D.W. & King, P. E. (1973). Relationships between some British littoral and sublittoral bryozoans and pycnogonids. In: Larwood. G.P. (Ed.). Living and Fossil Bryozoa. Academic Press, London: 199–207. Yang, H.J., Seo, J.E. & Gordon, D.P. (2018a): Sixteen new generic records of Korean Bryozoa from southern coastal waters and Jeju Island, East China Sea: evidence of tropical affinities. Zootaxa 4422(4): 493–518. DOI: 10.11646/zootaxa. 4422.4.3. Yang, H.J., Seo, J.E., Min, B.S., Grischenko, A.V. & Gordon, D.P. (2018b): Cribrilinidae (Bryozoa: Cheilostomata) of Korea. Zootaxa 4377(2): 216–234. DOI: 10.11646/zootaxa.4377.2.4. Zabala, M. (1986): Fauna dels briozous dels Països Catalans. Institut d’Estudis Catalans, Barcelona. Zágoršek, K. & Gordon, D.P. (2013): Late Tortonian bryozoans from Mut Basin, Central Anatolian Plateau, southern Turkey. Acta Palaeontol Polonica 58(3): 595–607. DOI: 10.4202/ app.2011.0100. Zágoršek, K. & Gordon, D.P. (2014): Revision of the Oligocene bryozoan taxa described by Stoliczka (1862), with the description of a new genus of Bryocryptellidae. Geodiversitas 36(4): 541–564. DOI: 10.5252/g2014n4a3. Zágoršek, K., Gordon, D.P. & Vávra, N. (2015): Revision of Chlidoniopsidae Harmer (Bryozoa: Cheilostomata) including a description of Celiopsis vici gen. and sp. nov. J Paleontol 89(01): 140–147. DOI: 10.1017/jpa.2014.12. Zágoršek, K., Ramalho, L.V., Berning, B. & de Araújo Távora, V. (2014): A new genus of the family Jaculinidae (Cheilostomata, Bryozoa) from the Miocene of the tropical western Atlantic. Zootaxa 3838(1): 98–112. DOI: 10.11646/ zootaxa.3838.1.5.

Appendix Glossary of common terms in Cheilostomata Term Abfrontal Acanthostegal brood chamber

Acleithral Adeoniform Adnate Adoral tubercle Adventitious avicularium Adventitious kenozooid Anastomosing Ancestrula, pl. ancestrulae Ancestrular complex Androzooid Anter Apertural bar Aperture Apex Areola, pl. areolae Areolar pore Articulated Articulated spine Ascopore Ascus Astogeny Autozooid Autozooidal polymorph Avicularium, pl. avicularia Basal pore-chamber Bilamellar Bilaminar Bilobate non-kenozooidal ooecium Bimineralic Bird‘s head avicularium Biserial https://doi.org/10.1515/9783110586312-012

– Explanation – See dorsal – Incubation chamber from periopesial spines with incubation cavity situated between the zooidal frontal membrane and tilted contiguous periopesial spines – Ovicell in which the opening is not closed by zooidal operculum and plugged by ooecial vesicle – See palmate – See encrusting – Tubercle-like gymnocystal structure in an oral position – Avicularium positioned on the frontal, lateral, or basal wall of a single zooid – Kenozooid positioned on the surface of a zooid – Erect colony with network of diverging and rejoining branches – First zooid(s) of the colony, formed by metamorphosis of a larva – Two to six zooids simultaneously formed from larva during metamorphosis – Incidentally used term for male zooid, modified or non-modified, containing spermatogenic tissue – Part of the orifice distal of the condyles – Pair of fused costae proximal of the orifice in cribrilinids – Opening of skeletal peristome – Peak of cup- or dome-shaped colonies – Shallow depression surrounding areolar pore in some umbonulomorphs – Pore on the periphery of umbonuloid/lepralioid frontal shield leading to areolar canal connecting hypostegal and perivisceral coeloms – Colonies or spines having flexible, chitinous joints – Skeletal spine with cuticular ring (joint) at its base – Median pore in lepralioid frontal shield serving as the inlet to the ascus – Sac-like hydrostatic structure (also termed compensation sac) underneath the frontal shield in ascophoran cheilostomes – Formation of bryozoan colony by zooidal budding – Feeding zooid – Term embracing all zooidal categories with autozooid-like cystid and protrusible lophophore with either functional or non-functional gut – Heterozooid with strongly modified operculum (mandible) and vestigial polypide – Pore-chamber situated at junction between lateral and basal or transverse and basal zooidal walls – Erect colony consisting of two layers of back-to-back zooids and with their fused basal walls – See bilamellar – Incubation chamber of fused hollow outgrowths of the frontal wall of a maternal zooid – Calcified cystid wall composed of aragonite and calcite layers – Stalked avicularium with the shape of a bird‘s head – Colony branch of two series of zooids

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 Appendix

Brood chamber Bud B-zooid Cauda Cellariiform Cleithral Closure plate Colony Compound ancestrula Compound kenozooidal ooecium Condyles Conventional ovicell Conventional ovicell with complete ooecium Conventional ovicell with spinose ooecium Cormidial orifice Coronate larva Coscinopleures Costa, pl. costae Costal field Costate shield Crossbar Cryptocyst Cup-shaped Cuticle Cystid Cyphonautes larva Dichotomous Disc-shaped (discoidal) Distal Dorsal Ectocyst Ectooecium Encrusting Endocyst Endooecium Endotoichal ovicell Endozooidal ovicell

– Chamber for larval incubation, including ovicells and ovicell-like structures, acanthostegal brood chambers and external and internal brood sacs – Newly developing zooid – Enlarged autozooidal polymorph that simulates an avicularium; frequently containing a polypide, e.g. in Steginoporella – Narrow elongated proximal portion of a club-like autozooid – Erect colony with multiple articulations – Ovicell with opening closed by zooidal operculum – Calcified plate sealing the opesia or orifice of a zooid – Functional unit of all asexually budded zooids derived from one larva or colony fragment – See ancestrular complex – Incubation chamber made of costae-like or plate-like kenozooids in Scrupariidae and Alysidiidae – Pair of oppositely placed calcified protuberances of skeletal orifices on which the operculum pivots – Brood chamber with spherical or semispherical protective two-walled outfold (ooecium), variously projected above the colony surface – Ovicell with complete (non-spinose) ooecium – Ancestral conventional ovicell with ooecium made of hollow non-jointed spines (costae) – Orifice that looks compound since it is surrounded by the skeletal shields of two to four neighbor zooids – Short-living non-feeding bryozoan larva – Asymmetrical, avicularia-like polymorphs with a porous cryptocyst – Modified, non-articulated periopesial spine overarching the frontal membrane – Area of suboral costae – Frontal shield formed from series of fused or contiguous costae – See pivot bar – Calcified interior wall underlying frontal membrane – Conical colony shaped like a cup – Organic outer layer of body wall – Zooidal body wall, receptacle of polypide – Long-living planktotrophic bryozoan larva with bivalved cuticular shell – Branching pattern dividing branch on two – Conical colony shape with flat base – Within the growth direction, i.e. away from the ancestrula – Side of bryozoan colony opposite the frontal side bearing the zooidal apertures, often synonymous to basal – Outer extracellular secreted layer (skeletonized in most bryozoans) of the cystid wall – Outer (external) wall of ooecium, calcified, with cuticular pseudopores or fenestrae or totally membranous – Colony creeping over substrate, uni- or multilamellar, attached by basal zooidal walls over a large part of its area – Epithelial lining of ectocyst – Also termed entooecium – inner (internal) wall of ooecium, totally calcified, but sometimes with pseudopores or cuticular areas – Ovicell in Cellariidae, strongly modified endozooidal type – Ovicell with its brooding cavity enclosed in the proximal part of the distal zooid

Glossary of common terms in Cheilostomata 



Ephebic Epitheca Erect Eschariform External brood sac Fenestr(ul)a Fertile zooid Foramen, pl. foramina

– – – – – – – –

Fragmentation Free-living Frondose Frontal Frontal budding

– – – – –

Frontal membrane Frontal shield

– –

Fungiform Fusion of costae Gymnocyst Gynozooid

– – – –

Heterozooid

–

Hydrostatic system

–

Hyperstomial ovicell

–

Hypostegal coelom

–

Immersed ovicell Intercostal lacuna, pl. lacunae Internal brood sac

– – –

Internode Interzooidal Intracolony overgrowth Intracuticular skeleton Intramural budding

– – – – –

Kenozooid Longitudinal (section) Lophophore Lunulitiform

– – – –

Lyrula

–

Mandible

–

 427

Descriptive term for fully-formed zooids in zone of astogenetic repetition Cuticular exterior layer of a skeletal frontal wall Upright colony attached to substratum either by rhizoids or encrusting base See frondose External membranous sac for embryonic incubation Open space in reticulate colonies Egg-producing zooid Openings in the frontal shield connecting external environment to the space between the calcified frontal shield and the frontal membrane in arachnopusiids and exechonellids Asexual type of reproduction via breakage of colony into smaller pieces Discoidal, conical, or irregular colony non-attached to substratum Colony composed of broad erect sheets (also eschariform) Side of bryozoan colony bearing the zooidal apertures Budding from frontal colony surface to produce adventitious polymorphs or multilamellar colonies Frontal wall of flexible cuticular layer lined by epithelium in anascans Protective skeletal structure developing on the frontal side of autozooid in cribrimorphs and ‘ascophorans’ Mushroom-shaped or umbrelliform colony Lateral and distal fusion of costae Simple exterior calcified wall (frontal, lateral, basal) with cuticular outer layer Old-fashioned term for female zooid, modified or non-modified, that is used for larval incubation Specialized non-feeding zooid with modified cystid and no polypide or its vestige System for protrusion of the lophophore, consisting of the flexible frontal membrane or ascus and attached parietal muscles Synonym for prominent ovicell in which half or more of the brood cavity is above the colony surface Slit-like coelom between membranous frontal wall and underlying calcification in cheilostomes with cryptocyst and umbonuloid and lepralioid frontal shields Ovicell with its brooding cavity enclosed in the distal part of the maternal zooid True pore between costae in cribrimorphs Deep invagination of non-calcified distal wall of the maternal zooid serving for embryonic incubation Part/fragment of articulated colony between nodes (joints) Positioned between zooids, e.g. avicularia or kenozooids Overgrowth of encrusting zooids on the colony surface Skeletal parts sandwiched between cuticular walls/layers Zooidal bud formed within cystid of an existing zooid as a result of zooidal damage followed by regeneration Heterozooidal polymorph lacking a polypide and operculum See sagittal Сiliated tentacle crown Disc- to cup-shaped free-living colonies, and frequent, regularly dispersed vibracula; slightly raised above the sea-floor by the marginal setae Supraopercular median tooth on proximal side of the skeletal orifice, often anvil-shaped Motile cuticular part of an avicularium, comprising a modified zooidal operculum

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 Appendix

Medial

– Along the midline

Metamorphosis Mucro (pl. mucrones) Multilamellar

– Transformation of a larva to produce ancestrula or ancestrular complex – Spinous elevation of the proximal margin of the skeletal orifice – Colony consisting of two or more laminae of zooids

Multilaminar Multiserial Mural pore-chamber

– See multilamellar – Colony or colony branch of more than 10 series of zooids – Pore-chamber situated between the adjacent lateral walls of two neighboring zooids and consisting of an enclosed space between a uni- or mutiporous septulum and a single pore in the opposing zooidal wall – Descriptive term for early-formed zooids near growing edge of colony – Place of articulation, cuticular tube-like joint – Pair of skeletal plates or ridges adjacent to distolateral zooidal walls; the intervening cavity accommodates opercular occlusor muscle insertions – Muscles closing the operculum – Non-calcified contractile outfold of the distal zooidal wall plugging an ovicell opening – Colony or colony branch of 3 to 10 rows of zooids – All events during the development of one zooid – Protective part of the ovicell, well-developed or vestigial, outfold with totally or partially calcified walls and interviening coelom – Trace of operculum preserved in a closure plate – Cuticular lid for closure of the orifice in cheilostomes – In “anascans” the opening below the frontal membrane restricted by skeletal walls, gymnocystal or cryptocystal – Lateral indentations in distal margin of cryptocyst (proximal margin of opesia) for the passage of parietal muscles – Small opening or pore in the cryptocyst proximal to the opesia for passage of parietal muscles or their ligament; pairwise or in series – Situated around the skeletal orifice, e.g. spines – Opening in the frontal wall (membranous or skeletal) closed by operculum, through which the lophophore is protruded – Incubation chamber, consisting of brood cavity, protective calcified outfold (ooecium) and (often) ooecial vesicle, plugging (ovicell) opening to the brood cavity – Horizontal membranous wall of the rostrum on which the mandible lies when closed, corresponds to flattened vestibulum of autozooid – Colony with flattened branches (also adeoniform) – Bilaterally paired muscles traversing the body cavity and attached to the frontal membrane or ascus and serving for polypide protrusion – Elevated on stalk – Respectively small or large pseudopores in spinocystal costae

Neanic Node Occlusor lamina Occlusor muscles Ooecial vesicle Oligoserial Ontogeny/ontogenetic Ooecium Opercular scar Operculum Opesia (pl. opesiae) Opesiular indentations Opesiule Oral Orifice Ovicell

Palate Palmate Parietal muscles Pedunculate Pelmatidium/pelma (pl. pelmatidia/pelmata) Periopesial Periorificial teeth Peripheral caverns Peristome Peristomial ovicell Petraliiform

– – – – – –

Surrounding the opesia, e.g. spines Tooth-like skeletal projections around the primary orifice Hollows or grooves of the cryptocyst Skeletal tube surrounding the primary orifice and operculum at inner end Ovicell with the ooecium enclosed in peristomial tube Encrusting, unilamellar colony loosely attached to substratum by basal walls or basally budded kenozooids



Pivot bar

Glossary of common terms in Cheilostomata 

 429

– Complete skeletal crossbar in some avicularia on which the mandible is pivoted Pointed rostrum – Rostrum of avicularia having a sharp (acute) or tapered tip Polymorphism – Discontinuous variation in morphology of zooids in colony Polypide – Internal organs including tentacle crown (lophophore), gut and associated musculature and nerve elements Pore-chamber – Interzooidal communication organ – small space developing between (or at junction of) two zooidal walls and enclosed between uni- or multiporous septula from one side and a single opening in the opposite zooidal wall Pore-plate – Round or oval thinned area in transverse zooidal wall perforated by one or several communication pores; pore-plates in pore-chambers are termed septula Poster – Part of the skeletal orifice proximal of the condyles, leading to the ascus Primary orifice – In cheilostomes with a peristome, zooidal opening at its proximal end, i.e. original zooidal orifice Proximal – Against the growth direction, i.e. toward the ancestrula Proximally attached ancestrula – Erect ancestrula Pseudopore – Non-calcified round or irregular area with thickened cuticle in the calcified wall Pseudosinus – Slit (or notch) at proximal edge of the secondary orifice formed at the distal end of the peristome Pyriform – Elongate pear-shaped Quincunxial – Arrangement of five zooids with one being located in the center and the other four placed at the corners Regeneration – Restoration of the damaged zooid or colony part (see also intramural budding) Reteporiform – See reticulate Reticulate – Colony formed like a reticulate network Retractor muscles – Paired muscles serving for retraction of the polypide Reversed polarity of avicularia – Avicularia with the rostrum oriented proximally Rhizoid – Heterozooidal polymorph (kenozooid) modified as a rootlet for stabilizing a colony to the substratum Rostrum – Distal part of an avicularium, bearing the mandible Sagittal (section) – Parallel to the lateral walls of the zooids Schizoporelloid ancestrula – Ancestrula with lepralioid frontal shield and a median sinus at the proximal margin of the skeletal orifice Scutum – Shield-like flattened, or branched spine partially covering opesia Secondary orifice – Opening at the outer extremity of the peristome Septulum, pl. septula – Uni- or multiporous pore-plate in mural and basal pore-chambers, i.e. a little septum Setiform – Elongated, thin, and narrow mandible of vibraculum Sexual zooid – Zooid producing eggs and/or sperm Sinus – Slit (or notch) at proximal edge of skeletal orifice Spine – Hollow pointed projection of gymnocystal wall Spine base – Circular pit or pore left after the spine detachment; characteristic of ephemeral articulated spines Spinocyst – Frontal shield (also termed pericyst) of contiguous costae Spiramen – Opening in the frontal shield proximal to the secondary orifice serving as an inlet to peristome (in some lepraliomorphs) or to ascus (in some umbonulomorphs) Subhypostegal (subfrontal) – A thin space or tiny cavity beneath the skeletal frontal wall in Euthyrisellidae coelom Subimmersed ovicell – Ovicell with less than half the brood cavity above the colony surface

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 Appendix

Subterminal orifice

– Orifice located proximally of the distal end of the zooid

Tatiform ancestrula

– Ancestrula with membranous frontal wall, often surrounded by periopesial spines – Kenozooid in a terminal position, i.e. last zooid budded in a row/colony – Orifice located in the distalmost position – Ovicell with ooecium in a terminal position in zooidal row – Parallel to the transverse walls of the zooids

Terminal kenozooid Terminal orifice Terminal ovicell Transverse (section) Transverse polarity of avicularia Umbo Unilamellar Unilaminar Uniserial Vibraculum, pl. vibracula Vicarious avicularium Vinculariiform Zooecium

– Avicularia with rostrum oriented transversely in respect to autozooid – – – – –

Prominence on the frontal wall proximal of the orifice Colony having one layer (lamina) of zooids See unilamellar Colony/colony branch consisting of one row of zooids Highly modified avicularium with setiform mandible slung between condyles or pivoting on a skeletal bar – Avicularium replacing an autozooid in zooid row, usually having the same size or larger than autozooid – Colony with erect, stalk-like branches – Skeletal wall of bryozoan zooid (used in older literature, especially by paleontologists)



Systematic classification of Cheilostomata used in this chapter 

 431

Systematic classification of Cheilostomata used in this chapter Phylum Bryozoa Ehrenberg, 1831 Order Cheilostomata Busk, 1852a Suborder Membraniporina Ortmann, 1890   Superfamily Membraniporoidea Busk, 1852b     Family Membraniporidae Busk, 1852b     Family Electridae Stach, 1937 (1851)     Family Sinoflustridae Gordon, 2009a Suborder Inovicellina Jullien, 1888   Superfamily Aeteoidea Smitt, 1868a     Family Aeteidae Smitt, 1868a Suborder Scrupariina Silén, 1941   Superfamily Scruparioidea Gray, 1848     Family Scrupariidae Gray, 1848     Family Eucrateidae Johnston, 1847     Family Leiosalpingidae d’Hondt & Gordon, 1996 Suborder Tendrina Ostrovsky, 2013   Superfamily Tendroidea Vigneaux, 1949     Family Tendridae Vigneaux, 1949 Suborder Thalamoporellina Ostrovsky, 2013   Superfamily Thalamoporelloidea Levinsen, 1902     Family Thalamoporellidae Levinsen, 1902     Family Steginoporellidae Hincks, 1884a Suborder Belluloporina Ostrovsky, 2013   Superfamily Belluloporoidea Ostrovsky, 2013     Family Belluloporidae Ostrovsky, 2013 Suborder Flustrina Smitt, 1867   Superfamily Flustroidea Lamoroux, 1816     Family Flustridae Lamouroux, 1816   Superfamily Buguloidea Gray, 1848     Family Bugulidae Gray, 1848     Family Beaniidae Canu & Bassler, 1927     Family Candidae d’Orbigny, 1851     Family Epistomiidae Gregory, 1893     Family Euoplozoidae Harmer, 1926     Family Jubellidae Reverter-Gil & FernándezPulpeiro, 2001     Family Rhabdozoidae MacGillivray, 1887a   Superfamily Calloporoidea Norman, 1903a     Family Calloporidae Norman, 1903a     Family Antroporidae Vigneaux, 1949

    Family Bryopastoridae d’Hondt & Gordon, 1999     Family Chaperiidae Jullien, 1888     Family Cupuladriidae Lagaaij, 1952     Family Cymuloporidae Winston & Vieira, 2013     Family Doryporellidae Grischenko, Taylor, & Mawatari, 2004     Family Ellisinidae Vigneaux, 1949     Family Farciminariidae Busk, 1852b     Family Foveolariidae Gordon & Winston, 2005 in Winston (2005)     Family Heliodomidae Vigneaux, 1949     Family Hiantoporidae Gregory, 1893     Family Mourellinidae Reverter-Gil, Souto, & Fernández-Pulpeiro, 2011     Family Pyrisinellidae Di Martino & Taylor, 2012     Family Quadricellariidae Gordon, 1984   Superfamily Cellarioidea     Family Cellariidae Lamouroux, 1816     Family Membranicellariidae Levinsen, 1909   Superfamily Lunulitoidea Lagaaij, 1952     Family Lunulariidae Levinsen, 1909     Family Otionellidae Bock & Cook, 1998     Family Selenariidae Busk, 1854   Superfamily Microporoidea Gray, 1848     Family Microporidae Gray, 1848     Family Alysidiidae Levinsen, 1909     Family Aspidostomatidae Jullien, 1888     Family Calescharidae Cook & Bock, 2001     Family Calpensiidae Canu & Bassler, 1923     Family Chlidoniidae Busk, 1884     Family Onychocellidae Jullien, 1882b     Family Poricellariidae Harmer, 1926     Family Setosellidae Levinsen, 1909   Superfamily Monoporelloidea Hincks, 1882b     Family Monoporellidae Hincks, 1882b     Family Macroporidae Uttley, 1949   Superfamily Adeonoidea Busk, 1884     Family Adeonidae Busk, 1884     Family Inversiulidae Vigneaux, 1949   Superfamily Arachnopusioidea Jullien, 1888     Family Arachnopusiidae Jullien, 1888     Family Exechonellidae Harmer, 1957   Superfamily Bifaxarioidea Busk, 1884     Family Bifaxariidae Busk, 1884     Family Mixtopeltidae Gordon, 1994a

432 

 Appendix

  Superfamily Catenicelloidea Busk, 1852b     Family Catenicellidae Busk, 1852b     Family Eurystomellidae Levinsen, 1909     Family Petalostegidae Gordon, 1984     Family Savignyellidae Levinsen, 1909   Superfamily Celleporoidea Johnston, 1838     Family Celleporidae Johnston, 1838     Family Colatooeciidae Winston, 2005     Family Hippoporidridae Vigneaux, 1949     Family Phidoloporidae Gabb & Horn, 1862   Superfamily Chlidoniopsoidea Harmer, 1957     Family Chlidoniopsidae Harmer, 1957   Superfamily Conescharellinoidea Levinsen, 1909     Family Conescharellinidae Levinsen, 1909     Family Batoporidae Neviani, 1901     Family Lekythoporidae Levinsen, 1909     Family Orbituliporidae Canu & Bassler, 1923   Superfamily Cribrilinoidea Hincks, 1879     Family Cribrilinidae Hincks, 1879     Family Euthyroididae Levinsen, 1909     Family Polliciporidae Moyano G., 2000   Superfamily Didymoselloidea Brown, 1952     Family Didymosellidae Brown, 1952   Superfamily Euthyriselloidea Bassler, 1953     Family Euthyrisellidae Bassler, 1953     Family Clathrolunulidae Gordon & Sanner, 2020     Family Neoeuthyrididae Gordon & Sanner, 2020   Superfamily Hippothooidea Busk, 1859b     Family Hippothoidae Busk, 1859b     Family Chorizoporidae Vigneaux, 1949     Family Haplopomidae Gordon in De Blauwe, 2009     Family Pasytheidae Davis, 1934 (1881)     Family Trypostegidae Gordon, Tilbrook & Winston in Winston, 2005     Family Vitrimurellidae Winston, Vieira, & Woollacott 2014   Superfamily Lepralielloidea Vigneaux, 1949     Family Lepraliellidae Vigneaux, 1949     Family Atlantisinidae Berning, Harmelin & Bader, 2017     Family Bryocryptellidae Vigneaux, 1949     Family Dhondtiscidae Gordon, 1989a     Family Hincksiporidae Powell, 1968     Family Jaculinidae Zabala, 1986     Family Metrarabdotosidae Vigneaux, 1949     Family Romancheinidae Jullien, 1888     Family Sclerodomidae Levinsen, 1909

    Family Tessaradomidae Jullien in Jullien & Calvet, 1903     Family Umbonulidae Canu, 1904b   Superfamily Mamilloporoidea Canu & Bassler, 1927     Family Mamilloporidae Canu & Bassler, 1927     Family Ascosiidae Jullien, 1882a     Family Cleidochasmatidae Cheetham & Sandberg, 1964     Family Crepidacanthidae Levinsen, 1909   Superfamily Pseudolepralioidea Silén, 1941     Family Pseudolepraliidae Silén, 1941   Superfamily Schizoporelloidea Jullien, 1882a     Family Schizoporellidae Jullien, 1882a     Family Acoraniidae López‐Fé, 2006     Family Actisecidae Harmer, 1957     Family Buffonellidae Jullien, 1888     Family Calwelliidae MacGillivray, 1887     Family Cheiloporinidae Bassler, 1936     Family Cryptosulidae Vigneaux, 1949     Family Cyclicoporidae Hincks, 1884b     Family Echinovadomidae Tilbrook, Hayward, & Gordon, 2001     Family Eminooeciidae Hayward & Thorpe, 1988c     Family Escharinidae Tilbrook, 2006     Family Fatkullinidae Grischenko, Gordon, & Morozov, 2018     Family Fenestrulinidae Jullien, 1888     Family Gigantoporidae Bassler, 1935     Family Hippaliosinidae Winston, 2005     Family Hippopodinidae Levinsen, 1909     Family Lacernidae Jullien, 1888     Family Marcusadoreidae Winston, Vieira, & Woollacott 2014     Family Margarettidae Harmer, 1957     Family Mawatariidae Gordon, 1990     Family Microporellidae Hincks, 1879     Family Myriaporidae Gray, 1841     Family Pacificincolidae Liu & Liu, 1999     Family Petraliidae Levinsen, 1909     Family Phoceanidae Vigneaux, 1949     Family Phorioppniidae Gordon & d’Hondt, 1997     Family Porinidae d’Orbigny, 1852     Family Robertsonidridae Rosso in Rosso et al., 2010     Family Stomachetosellidae Canu & Bassler, 1917     Family Tetraplariidae Harmer, 1957     Family Teuchoporidae Neviani, 1896b



    Family Vicidae Gordon, 1988   Superfamily Siphonicytaroidea Harmer, 1957     Family Siphonicytaridae Harmer, 1957   Superfamily Smittinoidea Levinsen, 1909     Family Smittinidae Levinsen, 1909     Family Bitectiporidae MacGillivray, 1895     Family Lanceoporidae Harmer, 1957 (1927)     Family Powellithecidae Di Martino, Taylor, Gordon, & Liow, 2016

Systematic classification of Cheilostomata used in this chapter 

    Family Watersiporidae Vigneaux, 1949   Superfamily Urceoliporoidea Bassler, 1936     Family Urceoliporidae Bassler, 1936     Family Prostomariidae MacGillivray, 1895 Suborder Skylonina Viskova in Viskova & Morozova, 1988   Superfamily Skylonoidea Sandberg, 1963     Family Bicorniferidae Keij, 1977

 433

Index acanthostegal 110, 116 acanthostyles 15, 19, 21 acidification 5 acidophil 72 Acoraniidae 384 Actinoporidae 245 Actisecidae 384 Adeonidae 357 Adeonoidea 357 adventitious avicularia 168 AEO larvae 134 Aeteidae 328 Aeteoidea 328 Aethozoidae 285 Aethozooides uraniae 95 Aeverrillia 153 Aeverrilliidae 299 Afrindella 213 aktinotostyles 15 Alcyonidiidae 275 Alcyonidioidea 275 alkaloids 178 alphaproteobacterial symbionts 136 Alveoli 234 Alyonushkidae 255 Alysidiidae 352 amathamides 178 Amathia 292 Amathia gracilis 79 amoebocytes 95 anal budding direction 271 anal density receptors 85 anal side 59 Anasca 326 anastomosing 280 anastomosing funicular cords 318 ancestral ectocyst 218 ancestrula 1, 58, 101, 231 ancestrular cystid 138 Anguinella 308 Anguinella palmata 285 Annectocymidae 244 annular muscles 60, 86, 153, 226 annulus 195 anoxia 29, 33 anterior median muscles 131 Antroporidae 340 anus 71 Anyutidae 255 apertural 58 apertural area 84, 85, 309 Apertural muscles 86 apertural papilla 270, 289 apertural rims 307 aperture 270

https://doi.org/10.1515/9783110586312-013

apical disc 128 apical organ 128 Arachnidiidae 280 Arachnidioidea 280 arachnidioideans 77 Arachnidium 280 Arachnopusiidae 357 Arachnopusioidea 357 aragonitic 35, 38, 39 areola 321, 322 areolar pores 322 Articulata 235, 240, 249 Articulated colonies 238 artificial shrinkage 77 Asajirella 207 Asajirella gelatinosa 93 Ascophora 326 ascopore 323 Ascosiidae 381 ascus 320, 322 ascus formation 322 asexual buds 309 Aspidostomatidae 352 astogeny 1, 58 Atlantisinidae 373 atrial sphincter 86 atrium 86, 133 attachment apparatus 196 attachment filaments 226 attachment ligaments 90 attachment organ 59, 85, 90, 226 augmented feeding 177 Autozooid movements 175 Avenella 296 avicularia 19, 24, 42, 165 avicularian behavior 169 avicularium 323 aviculomorphs 19, 23, 42 avoidance response 157 A zooids 168 basal diaphragms 14, 15, 232, 229 basal pore-chambers 318 basal side 58 basal transversal muscle 91 basophil 72 Bathyalozoidae 300 Bathymetric distribution 1 Batoporidae 366 Beaniidae 336 behavioral movements 180 behavior of feeding 165 Bellulopora 116 Belluloporidae 333 Belluloporina 333

436 

 Index

Belluloporoidea 333 Benedenipora 275 Benedeniporidae 274 Benedeniporoidea 274 benedeniporoidean 309 Bicorniferidae 404 Bifaxariidae 359 Bifaxarioidea 359 Biflustridae 326 bilateral 301 bimineralic 35 biofoulers 5 bioimmuration 11, 24 bioimmured 308 biradial cleavage 123 Bird’s head avicularia 167, 168 biserial branches 162 Bitectiporidae 402 blastemata 128 blastomere fates 127 Bockiella 277 body cavities 59 body wall muscles 88 body wall musculature 86, 192 boring 292 boring bryozoan 307 boring forms 280 boring ones 270 bristle or spine type 167 brood 273 brood chambers 110, 123, 234 brooding 14, 15, 21, 23, 35, 37, 41, 42, 101, 110 brooding zooids 289 brood pouches 285 brown body 1, 226 Bryocryptellidae 373 Bryopastoridae 340 bryostatins 5, 136, 178 bryozoan food 1 bryozoology 5 buccal dilatators 62, 93, 91 budding sites 58 Buffonellidae 384 Bugulidae 335 Buguloidea 168, 335 Buskiidae 292 B zooids 167 caecal cells 72 caecum 69 calcification 307 calcified cuticles 308 Calescharidae 352 Calloporidae 339 Calloporoidea 339 Calpensea nobilis 95 Calpensiidae 353 Calwelliidae 384 camouflage 292 campylonemidan 150 Canaliporidae 255

Cancellata 235, 240, 254 cancelli 234 Candidae 337 Candidatus Endobugula 136 cap cell 230 capitulum 277 cap-like apparati 15 capsule 195 capsule formation 202 Carbasea 178 cardelle 307 cardiac sphincter 69, 285, 296, 299, 309 cardiac valve 69 Catenicellidae 360 Catenicelloidea 360 Cauloramphus 153 cavernozooecia 19, 23 Cavidae 252 Cellariidae 348 Cellarioidea 348 Celleporella hyalina 165 Celleporidae 362 Celleporoidea 362 cell lineage 123 Celluliporidae 243 cementing body 196 cementing layer 202 centripetal flow 160 cerebral ganglion 77 Cerioporidae 249 Cerioporina 235, 240, 249 Chaetopterus 301 Chaperiidae 341 Cheiloporinidae 386 Cheilostome 11, 37 cheilostome budding 319 cheilostome evolution 172 cheilostome growth forms 317 cheilostome phylogeny 324 cheilostome radiation 317 cheilostomes 12, 14, 15, 24, 35, 37, 38, 39, 40, 41, 42 cheilostome suborders 326 cheilostome systematics 326 chemical defense 213, 177 chemical ecology 177 chemical triggers 170 chimney formation 162 chimneys 158, 211 chitinous rods 308 Chlidoniidae 353 Chlidoniopsidae 365 Chlidoniopsoidea 365 Chorizoporidae 370 ciliary reversal 148, 156 ciliated groove 128 ciliated ridges 133 ciliation 69 Cinctipora 108 Cinctiporidae 102, 230, 246 cincture cells 65 circular lophophore 207, 216

Index 

circulation 63 circum-oral nerve ring 79 Clathrolunulidae 369 Clausidae 252 Clavoporidae 277 Cleavage pattern 127 Cleidochasmatidae 381 cleithral 113 coelomocytes 95, 195 coelomopore 101 Colatooeciidae 363 collar 59, 265, 270, 300 colonial animals 65 colonial feeding patterns 157 colonial fission 190 colonial integration 163 coloniality 180 colonial system 77 colony chimney 160 colony feeding currents 163 colony morphology 191 colony surface topography 160 Commensalism 180 common bud 232 communication areas 65 communication pores 13, 27, 35, 229, 318 compensation space 322 competition 175, 177, 324 competitive dominance 176 compound colonies 211, 238 compound tentacle sheath neurite bundle 85 compound walls 227 condyles 317 Conescharellinidae 365 Conescharellinoidea 365 confluent body cavity 205 cormidium 113 corona 128 coronal cells 128 coronal retractors 131 coronate larva 299 Coronate larvae 134 Corymboporidae 252 corynotrypid 225 Corynotrypidae 241 costal shield 320 costate frontal wall 165 Crassodiscoporidae 254 creeping 211 creeping mobility 218 creeping movement 205, 211 creeping sole 205 creeping stolons 309 Crepidacanthidae 381 Cribrilinidae 320, 367 Cribrilinoidea 367 Crisiidae 249 Crisinidae 254 Cristatella mucedo 95 Cristatellidae 205 Crisuliporidae 249

Crownoporidae 241 cruciform budding pattern 216 cryptocyst 321 cryptocystal opercula 317 Cryptopolyzoon 292 cryptostomes 14, 15, 19, 25, 27, 29, 31, 42 Cryptosulidae 387 ctenostome 11, 12, 13, 24, 27, 29, 35, 40, 54, 55, 225 ctenostome-like ancestor 317 Ctenostome phylogeny 308 Cupuladriidae 342 cupuladriids 177 curling of branches 173 cuticle 65 cuticular septae 65 cuticular wrinkles 301 Cyclicoporidae 387 Cyclostomata 11, 13, 15, 21, 35 Cyclostome development 127 cyclostome larvae 129 cyclostomes 11, 12, 13, 14, 15, 24, 33, 35, 37, 38, 39, 41, 42 cyclostome taxonomy 240 cyclozooecia 19, 23 Cymuloporidae 343 cyphonautes 123, 133, 134, 265 cyst formation 198 cystid 57 cystid anastomoses 280 cystid appendages 280, 285, 299, 305, 309 cystigenic 200 cystiphragms 14, 15, 19, 21 cystoporates 13, 15, 25, 29, 31, 33, 38, 39 Cytididae 254 cytophore 104 Dactylethrae 233 deep sea 280 defecation 155 degeneration-regeneration cycles 19 Densiporidae 252 depressor bars 321 deutoplasmatic 200 Dhondtiscidae 375 diagonal layer of muscles 192 Diaperoeciidae 244 diaphragm 86 diaphragmatic sphincter 59, 85, 86, 90 Didymosellidae 368 Didymoselloidea 368 diffuse plexus 83, 85 digestion 69 Digestive system 69 Digestive tract muscles 93 Diploclemidae 241 Discoporella 173 dispersal 190 disporellids 226 dissolved organic matter 143 Dogger Bank Itch 277 dormant buds 273, 274 dormant stages 190

 437

438 

 Index

Doryporellidae 343 double-walled 228 duplicature band 84, 85, 88, 90, 93 Echinella 289 Echinovadomidae 387 ecology 324 economic significance 5 ectocyst 63, 191 ectocyst tubes 216 ectooecium 110 Electra pilosa 163 Electridae 327 Eleidae 246 eleozooids 42, 234 Ellisinidae 343 Elzerina 277 embryonic development 123 embryonic incubation 110 emerged larvae 163 Eminooeciidae 387 enantiomorph 285, 292 encrusters 270, 289 encrusting 309 encrusting cyclostomes 176 encrusting sheets 1 endemism 24, 27, 29 endosaccal cavity 59, 226 endosymbiotic microorganisms 178 endotoichal 115 endozooidal 114 Entalophoridae 245 entooecium 110 epidermal blastemata 137 epidermal cells 67 epistome 91, 192 epistomial canal 62 Epistomiidae 102, 108, 338 epithelial disc 200 epithelial vesicle 200 epizooic 1 erect bilaminate 162 erect colony branches 213 Escharinidae 388 esophagus 71 Esthonioporata 11, 13, 14, 17, 21, 24 esthonioporine 24, 25 Eucrateidae 330 Euctenostomata 274 Euoplozoidae 338 Eurystomata 7 Eurystomellidae 360 Euthyrisellidae 369 Euthyriselloidea 369 Euthyroididae 368 Evolutionary trends 238 excretory bladder 205 excretory organs 95 excretory system 95 excurrent chimney 156, 205, 159 Exechonellidae 358

exilazooecia 14, 19, 21, 40 exosaccal cavity 59, 226 expansion 153 exploratory behavior 136 Exterior walls 231 extinction 25, 27, 29, 31, 33, 37, 38, 44 extracellular matrix 67 extraembryonic nutrition 102 extrazooidal skeleton 229 extrazooidal structures 233 eyespots 129 Facilitation 180 Farciminariidae 343 Farrellidae 301 fascicles 238 Fatkullinidae 389 fecal pellets 71, 155 feeding 143 feeding behavior 150, 155 feeding current 69, 15, 39, 41 Feeding interference 177 feeding modes 71 feeding patterns 145 feeding process 192 fenestra 195 fenestrates 14, 15, 23, 24, 25, 29, 31, 35, 38, 41, 42 Fenestrulinidae 390 fertilization 101, 104, 109, 123 fighting buds 177 filter pump 148 filtration mechanism 133 fingerlike protrusions 177 Firmatopores 233 fixed-walled 228 Flabellotrypidae 241 flask-shaped cells 129 flattening zooids 173 flexor muscles 301 flick particles 161 float cells 201 float chambers 196 Floatoblasts 195 Flustrellidra hispida 277 Flustrellidridae 277 Flustridae 334 Flustrina 334 Flustroidea 334 foliated fabric 231 follicular cells 104 food selection 148 food source 143 forceps type 167 foregut epithelium 71 foregut plexus 85 forked canal 61, 95, 192, 206 formation of statoblasts 196 forms of statoblasts 195 fouling 180, 191 Foveolariidae 344 fragmentation 173, 190

Index 

Fredericella tenax 196 Fredericella toriumii 207 Fredericellidae 207 fredericellid-plumatellid clade 218 free living 324 free living cheilostomes 173 free-walled 11, 228 frenaculum 305 freshwater 296 freshwater cyphonautes 289 Frondiporidae 245 frontal budding 160, 317, 319 frontal shields 317 frontal side 58 frontal wall 11, 42, 319 fungiform 238 funicular strands 104 funicular system 72, 292, 309 funiculi 289 funnel-shaped cystiphragms 15 gamete release 109 ganglionic horns 79, 91 ganglionic lumen 79 gas exchange 322 gastrulation 124 Gelatinella 213 germinated zooids 205 germination of statoblasts 204 gigantism 31 Gigantoporidae 390 gizzard 69, 153, 230, 292, 296, 299, 308 glandular cells 63 glandular field 128 gnawing apparatus 301 Gondwana 24, 31 gonochorism 102 gonozooids 102, 110, 116, 127, 225, 273 granulocytes 95 gravity responses 135 group actions 158 gut contents 143 gymnocyst 320 gymnocystal frontal shield 321, 322, 323 Gymnolaemata 12, 21, 24 gymnolaemate ancestor 268 Gymnolaemate origins 266 Haplopomidae 370 Harmeriellidae 301 Hastingsiidae 246 Haywardozoon 308 Heliodomidae 345 hemiphragms 14, 15, 21, 42, 232 hemisepta 15, 21, 23, 31, 33, 232 hermaphrodites 101 heterochrony 42 heterozooids 23, 40, 41, 233, 323 hexagonal semi-nacre fabric 231 Hiantoporidae 346 hibernacula 273

hierarchical modules 181 Hiller’s plexus 85 Hincksiporidae 375 Hippaliosinidae 390 Hippopodinidae 391 Hippoporidridae 363 Hippoporina 162 Hippothoidae 369 Hippothooidea 369 Hislopiidae 289 Hislopioidea 289 homeomorphy 19, 42 homogeneous fabric 231 Horneridae 254 horseshoe shaped 79, 91, 192 Hyalinella 213 hydrostatic apparatus 319 hydrostatic pressure 154 hydrostatic protrusion 152 hyperstomial 114 hypertubercles 214 Hypophorellidae 301 hypostegal coelom 319 Immergentia 271 incurrent cells 159 individual behavior 150, 161 individual fabrics 231 individualized behavior patterns 157 ingestion rate 148 inner epithelial layer 200 interconnecting cords 77 interconnecting funicular cords 77 intercoronal cells 128, 129, 135 interior walls 13, 33, 231 internal sac 128, 129, 133 Internectella 190 Internectella bulgarica 218 intertentacular bases 67, 265 intertentacular fork 83 intertentacular membrane 67, 192, 207, 214 intertentacular organ 101, 109, 163, 266, 277, 280, 285 intertentacular perikarya 83 intertentacular pits 67, 83, 265 intertentacular radial nerve 83 Interzooidal avicularia 167 interzooidal budding 319 interzooidal nutrient exchange 228 interzooidal pore 65 Interzooidal pore complexes 265 interzooidal septa 206 interzooids 299 intraovarian 101 invasive 5 invasive species 190, 211 Inversiulidae 357 Jaculinidae 375 Jebramella angusta 164 Jebramellidae 301 jelly-like ectocyst 216

 439

440 

 Index

Joint retractions 161 Jubellidae 338 Jurassic 11, 19, 24, 33, 35, 36, 37, 39, 40, 42, 54 Kamptozoa 6 Kellwasser event 29 kenozooids 19, 40, 271, 323 Kinetoskias 173 kneading 93 Kukersella 241 Lacernidae 392 Lake Baikal 289 Lanceoporidae 402 larvae 15, 27, 37, 41 larval behavior 135 larval morphology 128 larval musculature 131 larval types 133 lateral nerves 129 lateroabfrontal neurite bundles 83 laterofrontal neurite bundles 83 lecithotrophic 123 Leiosalpingidae 331 Leiosoeciidae 251 Lekythoporidae 366 Lepraliellidae 372 Lepralielloidea 372 lepralioid shield 321, 322 Leptoblasts 196 leucocytes 95 Lichenoporidae 255 life histories 118 limiting cells 65 living chamber 226 Lobiancopora 277 long peristomes 285 lophophoral arms 67, 79, 192, 207 lophophoral arms muscles 91 lophophoral base 59 lophophoral base muscles 91 lophophoral concavity 61 lophophoral muscles 90 Lophophorata 6 lophophore 13, 41 lophophore shape 150 lophophore size 150 Lophopodella 207 Lophopodella carteri 95 Lophopodidae 207 lunaria 14, 17, 19, 21, 35, 42 Lunulariidae 350 lunulitiform colonies 163 Lunulitoidea 350 lymphocytes 95 Macroporidae 356 macrozooecia 14, 15, 19 maculae 14, 41, 42, 165, 233 Malacostegina 317, 326

male zooids 165 Mamilloporidae 380 Mamilloporoidea 380 mandible 165 mantle cavity 128 mantle larva 190 Marcusadoreidae 393 Margarettidae 394 matrotrophy 101, 110, 116 Mawatariidae 395 medioabfrontal neurite bundles 83 mediofrontal neurite bundle 83 melicerititid 230, 235 Membranicellariidae 349 Membranipora membranacea 148 Membraniporidae 326 Membraniporina 317, 324, 326 Membraniporoidea 326 membranous sac 59, 225 mesodermal cells 124 mesopsammal 285 mesotheca 19, 23, 25 mesozooecia 14, 19, 21, 23 metachronal laeotropic pattern 135 metachronal waves 146 metamorphic competence 127, 135 metamorphic patterns 136 metamorphosis 7, 136 metaxizooecia 19, 23 Metrarabdotosidae 376 microbial films 136 Microporellidae 395 Microporidae 351 Microporoidea 351 microzooecia 19, 23 midventral groove 153 Mimosellidae 301 mineralogy 5 mixed skeletal morpholo 230 Mixtopeltidae 360 molecular phylogenetic 7, 309 molting 177 Monastesia 305 Monobryozoon 285 Monobryozoidae 285 Monoporellidae 355 Monoporelloidea 355 monticulae 14 monticules 158 morphological characters 269 mortality 178 motility 172 Mourellinidae 347 mouth shape 150 mucus 69 mucus entrapment 69 multiporous 65, 296 multiporous pore plates 279 multiporous septula 318 Multisparsidae 243

Index 

multizooidal budding 319 mural hoods 233 mural pore-chambers 318 mural spines 15, 232 Muscular system 86 mutualism 180 myoepithelial pharynx 69, 93 myoepithelium 71, 93 Myolaemata 8, 308 Myriaporidae 395 myxozoans 190 nanozooecia 15 nanozooids 172, 234 nautizooids 289 Neoeuthyrididae 369 neozooecia 19, 21 nephridia 95 nerve nodule 128, 129 nervous system 57, 77 neuromuscular strand 124, 128, 129, 131 Nolella 271 Nolella alta 285 non-buoyant floatoblasts 207, 214 nonciliated tentacle 172 Nudibranch 325 nurse cells 101 obligate dormancy 205 obliquely truncate 150, 159 occlusor muscles 317 Oncousoeciidae 243 ontogenetic characters 265 Onychocellidae 353 oocyte doublet 104 oocytic doublets 104, 107 ooecial spines 116 ooeciopores 235 ooeciostomes 235 ooecium 110, 111 oogenesis 104 oogonia 102 opercula 42 operculum 24, 307 operculum occlusor 90 opesia 321 opesiules 321 oral groove 67 oral side 59 oral tentacles 91 Orbituliporidae 366 organic shells 128 orifical sphincter 88 orifice 59 orificial sphincter 88, 275 osmoconformers 189 Otionellidae 350 outer epithelial layer 201 outer nerve ring 79 ovarian wall 104

overgrowth 175 ovicell 41, 110, 163 ovicell type 115 ovipositor 164 Pachyzoontidae 280 Pacificincolidae 395 paedomorphosis 42 palaeostomates 225 palatal structures 172 Paleotubuliporina 240 pallial epithelium 128 pallial sinus 128, 133 Paludicella 270 Paludicella articulata 77, 79, 269 Paludicella pentagonalis 291 Paludicellidae 289 Paludicelloidea 289 Panolicella 305 paraphyletic 269 parazooecia 19, 23 parietal muscles 60, 85, 86, 265 parietodiaphragmatic muscle 90 parietovaginal bands 88, 90 parieto-vestibular 90 particle handling 156 particle rejection 157 Pasytheidae 371 Pectinatella 190 Pectinatella magnifica 172 Pectinatellidae 211 peduncle 277 pedunculate autozooids 307 Penetrantia 266, 271 Penetrantiidae 307 pentagonal 270, 291 pentagonal aperture 280 pentagonal arrangement 299 perennials 118 periblast 195 peripharyngeal plexus 86 Peripheral innervation 83 peristaltic contractions 71 peristaltic movements 71, 93 peristome 270 peristome enlargement 309 peristomial budding 285, 296, 299, 308, 309 peristomial tube 270 peromorphosis 42 Petaloporidae 254 Petalostegidae 360 Petraliidae 395 phagocytes 95 pharmacological 5 pharyngeal structure 309 Pherusella 277 Pherusellidae 277 Phidoloporidae 363 Phoceanidae 396 Phorioppniidae 397

 441

442 

 Index

photokinesis 135 photopositive 163 photoreceptors 128, 129, 135 phototactic larvae 163 phototaxis 135 Phylactolaemata 107 phylactolaemate development 127 phylactolaemate larvae 129, 134 phylactolaemates 11, 40 phylogenomic 7 phylogeny 6 phytoplankton 24, 37 Pierella 305 piptoblasts 196, 216 placental analogue 116 placental cheilostomes 77 placentation 107 placentotrophy 117 Plagioeciidae 244 planktonic 275 Planktotrophic 123 Platypolyzoon 308 pleated collar 305 Plumatella fruticosa 218 Plumatella osburni 218 Plumatellidae 213 Polliciporidae 368 polyembryony 41, 116, 127, 225 polymorph development 240 polymorphism 77, 172, 180, 233, 271, 323 polymorphs 14, 15, 19, 42 polypide 57, 58 polypide behaviors 155 polypide cycle 226 polypide eversion 319 polypide formation 266 polypide recycling 1 polypide rudiment 168 Polyzoa 5 Pore-chambers 318 pore complex 285 pore plate 65, 299 pore plugs 271 Poricellariidae 354 Porinidae 397 positive phototaxis 173 positive phototropism 173 Postgastrulation 127 post-Palaeozoic stenolaemates 225 postparitive incubation 110 Pottsiella 289 Pottsiellidae 299 Powellithecidae 402 preancestrula 136 predation 29, 40, 41, 42, 178, 325 predator 2, 29, 42 preoral cavity 128 preparitive 110 presumptive cystid 129 primary polypide 136

proancestrula 231 progressive cycles 226 Prostomariidae 404 protandrous 102 Protobenedenipora 275 protoctenostomes 266, 274 protoecium 225, 231 protogynous 102 protostomy 124 protrusion 153 proventriculus 69, 289 Pseudocerioporidae 252 pseudocoel 228 pseudocyphonautes 134, 277, 279 Pseudolepraliidae 382 Pseudolepralioidea 382 pseudolunaria 233 pseudopore distribution 232 pseudopores 14, 24, 65, 230, 322 ptilodictyines 15, 23, 25 pustules 232 Pustuloporidae 245 pycnogonids 325 pylorus 69 pyriform organ 128 Pyrisinellidae 348 Quadricellariidae 348 quiescence 204 radial aperture 307 radial fascicles 238 radial nerves 79 radial symmetry 275 ramets 180 raphe 191, 213 rasping apparatus 301 reciprocal overgrowth 175 Rectangulata 235, 240, 255 regeneration cycles 226 Rejection mechanisms 154 rejection movements 154 Reproductive behavior 163 Reproductive patterns 101 retraction 153 retractor muscles 59, 93 rhabdomesine 14, 19, 23, 25, 29, 31, 33 Rhabdozoidae 338 rheotropic 173 rheotropism 136 rhizoids 233, 271, 273 rhombic semi-nacre fabric 231 rigidly calcified colonies 161 ring canal 61, 91 ring canal musculature 91 ring nerve 79 Robertsonidridae 398 Romancheinidae 376 rosette-like 211 rotating food 71

Index 

Rumarcanella 213 runner-like 305 runners 270 sac zooids 271 saddle-shaped 213 Sagenellidae 241 sandy tubes 285 Savignyellidae 361 scanning activity 157 scanning behavior 153 Schizoporellidae 383 Schizoporelloidea 383 Sclerodomidae 377 Scrupariidae 329 Scrupariina 329 Scruparioidea 329 secondary metabolites 178 second funicular strand 77 Selenariidae 350 self-overgrowth 175 Semiceidae 246 sensory cells 71, 85 sensory function 192 septulum 318 sessile avicularia 167, 168 sessoblast 196 sessoblast formation 218 setigerous collar 153, 299 Setosellidae 355 settlement behavior 135 sexual dimorphism 102 shell adductors 131 shield evolution 321 shift of the polypide 265 sieving function 69 Sineportella 296 single-walled 228 Sinoflustridae 328 Siphonicytaridae 400 Siphonicytaroidea 400 Siphoniotyphlidae 246 skeletal apertures 226 skeletal frontal wall 320 skeletal mineralogy 225 skeletal organization 227, 229 skeletal ultrastructure 231 Skylonina 404 Skylonoidea 404 Smittinidae 401 Smittinoidea 401 solitary 280, 309 solitary bryozoans 270, 285 sources of yolk 104, 107 space balloons 301 Spathipora 271, 273 Spathiporidae 292 spatulate avicularia 168 spawned ova 165 special cells 65

species number 270 Spermatogonia 102 sperm collection 164 sperm release 164 spines 289 spinoblasts 196 spinocyst 320 spinose kenozooids 277, 299 spinozooids 271, 273, 300 Spiroporidae 244 standoff 175 stationary polypide cycles 226 statoblast 72 statoblast anlagen 196 statoblast evolution 218 statoblast release 195 Steginoporella 167 Steginoporellidae 332 Stenolaemata 11, 12, 13, 41, 44 Stephanellidae 216 Stigmatoechidae 254 Stolella 213 stolonal forms 270 stolonate colonies 291 stolon-like processes 216 Stomachetosellidae 399 stomatopod 308 Stomatoporidae 243 striation patterns 95 subepithelial 6 sublophophoral current 155 subperitoneal cell 79 substratum type 324 suction feeding 71 Sulcocavidae 252 Sundanellidae 296 supraneural coelomopore 109 suture 196 Swarupella 196, 213 swimming zooids 190 symbiotic relationships 180 synecium 115 Synnotum 153, 173 systematic categorization 270 systematics 7 Taghanic event 29 Tapajosella elongata 216 Tapajosellidae 216 teeth-like structures 289 temperature effects 135 Tendridae 332 Tendrina 331 Tendroidea 332 tentacle ciliation 230 tentacle curving 153 tentacle flicking 69, 146, 148, 157 tentacle innervation 79 tentacle muscles 90 tentacle neurite bundles 67

 443

444 

 Index

tentacle sheath 59 tentacle sheath length 152 tentacle sheath muscles 93, 309 tentacular neurite bundles 79 Terebriporidae 292 Tergozooecia 233 terminal diaphragms 229 Terviidae 244 Tessaradomidae 377 testing position 157 Tetraplariidae 399 Teuchoporidae 400 Thalamoporella 115 Thalamoporellidae 332 Thalamoporellina 332 Thalamoporelloidea 332 Theonoidae 245 Timanodictyina 11 topographic complexity 180 transverse fibrous fabric 231 transverse muscle fibers 299 transverse stolon muscle 301 trepostomes 14, 15, 19, 24, 25, 27, 29, 31, 33, 38, 39, 40 triangular mandibles 168 triangular shells 134 Triassic 11, 12, 13, 14, 21, 23, 33, 36, 39 trifid nerve 79, 85 triradiate 93 Triticellidae 305 true stolons 271 Trypostegidae 372 Tubuliporidae 243 Tubuliporina 235, 240, 241 turgescent 71 twin ancestrula 138 Type AE larvae 134 Type E larvae 134 Type O larvae 134 ultrastructural fabrics 231 Umbonulidae 378 umbonuloid shield 321, 323 uncalcifed bryozoans 269 unidirectional flow 161 upstream-feeding 6 Urceoliporidae 404 Urceoliporoidea 404 vacuolated cells 95 Varunella 213

vegetal quartet 124 vesicular collarette 128 Vesiculariidae 292 Vesicularioidea 291 Vesicularioideans 77 vestibular glands 308, 317 vestibular muscles 90 vestibular pore 110, 192, 211 vestibular wall 59, 86, 88 vestibulum 86 vestibulum dilatators 90 vestigial polypide 167 vibracula 165, 168 vibraculan behavior 170 vibracularia 15, 19 vibracular movement 171 vibratile plume 128 vicarious avicularia 167 Vicidae 400 Victorella 271 victorellid 93 Victorellidae 296 Victorelloidea 296 Visceral innervation 85 visceral neurite bundles 86 vitellogenesis 104, 107 Vitrimurellidae 372 viviparity 101, 110 v-shaped muscles 91 Walkeriidae 305 Walkerioidea 77, 299 walkerioidean 93 waterfowl 190 Watersiana 296 Watersiporidae 403 white spot 63, 195, 211, 213 Wilbertopora 172 winter species 190 zooarium 1 zooecium 1 zoogeographic 189 zooidal budding 319 zooidal polymorphism 317 zooidal shape 270 zooids 57 zoophytes 5, 143