Andreas Wanninger Editor Evolutionary Developmental Biology of Invertebrates Vol. 2 Lophotrochozoa (Spiralia) Evolutionary Developmental Biology of Invertebrates 2

Andreas Wanninger Editor

Evolutionary Developmental Biology of Invertebrates 2

Lophotrochozoa (Spiralia) Editor Andreas Wanninger Department of Integrative Zoology University of Vienna Faculty of Life Sciences Wien Austria

ISBN 978-3-7091-1870-2 ISBN 978-3-7091-1871-9 (eBook) DOI 10.1007/978-3-7091-1871-9

Library of Congress Control Number: 2015947925

Springer Wien Heidelberg New York Dordrecht London © Springer-Verlag Wien 2015 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifi cally the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfi lms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specifi c statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made.

Cover illustration: Scanning electron micrograph of an early trochophore larva of the tusk shell, Antalis entalis, a scaphopod mollusk. See Chapter 7 for details

Printed on acid-free paper

Springer-Verlag GmbH Wien is part of Springer Science+Business Media (www.springer.com) Pref ace

The evolution of life on Earth has fascinated mankind for many centuries. Accordingly, research into reconstructing the mechanisms that have led to the vast morphological diversity of extant and fossil organisms and their evolu- tion from a common ancestor has a long and vivid history. Thereby, the era spanning the nineteenth and early twentieth century marked a particularly groundbreaking period for evolutionary biology, when leading naturalists and embryologists of the time such as Karl Ernst von Baer (1792–1876), Charles Darwin (1809–1882), Ernst Haeckel (1834–1919), and Berthold Hatschek (1854–1941) realized that comparing ontogenetic processes between species offers a unique window into their evolutionary history. This revelation lay the foundation for a research fi eld today commonly known as Evolutionary Developmental Biology, or, briefl y, EvoDevo. While for many of today’s EvoDevo scientists the principle motivation for studying animal development is still in reconstructing evolutionary scenarios, the analytical means of data generation have radically changed over the cen- turies. The past two decades in particular have seen dramatic innovations with the routine establishment of powerful research techniques using micro- morphological and molecular tools, thus enabling investigation of animal development on a broad, comparative level. At the same time, methods were developed to specifi cally assess gene function using reverse genetics, and at least some of these techniques are likely to be established for a growing num- ber of so-called emerging model systems in the not too distant future. With this pool of diverse methods at hand, the amount of comparative data on invertebrate development has skyrocketed in the past years, making it increas- ingly diffi cult for the individual scientist to keep track of what is known and what remains unknown for the various animal groups, thereby also impeding teaching of state-of-the-art Evolutionary Developmental Biology. Thus, it appears that the time is right to summarize our knowledge on invertebrate development, both from the classical literature and from ongoing scientifi c work, in a treatise devoted to EvoDevo. Evolutionary Developmental Biology of Invertebrates aims at providing an overview as broad as possible. The authors, all renowned experts in the fi eld, have put particular effort into presenting the current state of knowl- edge as comprehensively as possible, carefully weighing conciseness against level of detail. For issues not covered in depth here, the reader may consult additional textbooks, review articles, or web-based resources,

v vi Preface

particularly on well- established model systems such as Caenorhabditis elegans ( www.wormbase.org ) or Drosophila melanogaster ( www.fl ybase.org ) . Evolutionary Developmental Biology of Invertebrates is designed such that each chapter can stand alone, and most chapters are dedicated to one phylum or phylum-like taxonomic unit. The main exceptions are the hexa- pods and the crustaceans. Due to the vast amount of data available, these groups are treated in their own volume each (Volume 4 and Volume 5, respec- tively), which differ in their conceptual setups from the other four volumes. In addition to the taxon-based parts, chapters on embryos in the fossil record, homology in the age of genomics, and the relevance of EvoDevo for recon- structing evolutionary and phylogenetic scenarios are included in Volume 1 in order to provide the reader with broader perspectives of modern- day EvoDevo. A chapter showcasing developmental mechanisms during regen- eration is part of Volume 2 . Evolutionary Developmental Biology of Invertebrates aims at scientists that are interested in a broad comparative view of what is known in the fi eld but is also directed toward the advanced student with a particular interest in EvoDevo research. While it may not come in classical textbook style, it is my hope that this work, or parts of it, fi nds its way into the classrooms where Evolutionary Developmental Biology is taught today. Bullet points at the end of each chapter highlight open scientifi c questions and may help to inspire future research into various areas of Comparative Evolutionary Developmental Biology . I am deeply grateful to all the contributing authors that made Evolutionary Developmental Biology of Invertebrates possible by sharing their knowledge on animal ontogeny and its underlying mechanisms. I warmly thank Marion Hüffel for invaluable editorial assistance from the earliest stages of this proj- ect until its publication and Brigitte Baldrian for the chapter vignette artwork. The publisher, Springer, is thanked for allowing a maximum of freedom dur- ing planning and implementation of this project and the University of Vienna for providing me with a scientifi c home to pursue my work on small, little- known creatures.

This volume covers the animals that have a ciliated larva in their life cycle (often united as Lophotrochozoa), as well as the Gnathifera and the Gastrotricha. The interrelationships of these taxa are poorly resolved and a broadly accepted, clade-defi ning autapomorphy is lacking. Spiral cleavage is sometimes assumed as the ancestral mode of cleavage of this grouping, and therefore the clade is named Spiralia by some authors, although others prefer to extend the term Lophotrochozoa to this entire assemblage. Aside from the taxon-based chapters, this volume contains a chapter that highlights similari- ties and differences in the processes that underlie regeneration and ontogeny, using the Platyhelminthes as a case study.

Tulbingerkogel, Austria Andreas Wanninger January 2015 Contents

1 Gnathifera...... 1 Andreas Hejnol 2 Gastrotricha ...... 13 Andreas Hejnol 3 Platyhelminthes ...... 21 Teresa Adell, José M. Martín-Durán, Emili Saló, and Francesc Cebrià 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study ...... 41 Francesc Cebrià, Emili Saló, and Teresa Adell 5 Cycliophora ...... 79 Andreas Wanninger and Ricardo Neves 6 Entoprocta ...... 89 Andreas Wanninger 7 Mollusca ...... 103 Andreas Wanninger and Tim Wollesen 8 Nemertea...... 155 Jörn von Döhren 9 Annelida ...... 193 Christoph Bleidorn, Conrad Helm, Anne Weigert, and Maria Teresa Aguado 10 Phoronida ...... 231 Scott Santagata 11 Ectoprocta...... 247 Scott Santagata 12 Brachiopoda ...... 263 Scott Santagata

Index ...... 279

vii Gnathifera 1 Andreas Hejnol

Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger. A. Hejnol Sars International Centre for Marine Molecular Biology , University of Bergen , Thormøhlensgate 55 , Bergen 5008 , Norway e-mail: [email protected]

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 1 DOI 10.1007/978-3-7091-1871-9_1, © Springer-Verlag Wien 2015 2 A. Hejnol

INTRODUCTION internal relationships using molecular data (Witek et al. 2008; Wey-Fabrizius et al. 2014 ). Figure 1.1 illustrates the likely phylogenetic The taxon Gnathifera was erected based on relationships as a consensus phylogeny that is morphological data by Ahlrichs ( 1995 , 1997 ). based on recent molecular as well as morpho- The taxon comprises the Gnathostomulida logical data. In all gnathiferans, fertilization is and Syndermata (which unites Rotifera, internal and the development direct. The para- Acanthocephala, Seisonida) (Fig. 1.1 ). With the sitic acanthocephalans have evolved additional discovery of Limnognathia maerski (Kristensen dispersal stages that allow infection and transi- and Funch 2000 ), the taxon Micrognathozoa has tions between hosts. been included into the Gnathifera. The name Gnathifera is based on the presence of a com- plex jaw apparatus in the pharynx of all groups, GNATHOSTOMULIDA except Acanthocephala (Sørensen 2003 ; Funch et al. 2005 ). Gnathifera are tiny, bilaterally Gnathostomulids are wormlike, microscopic, symmetric animals that live in aquatic habitats. marine, interstitial animals that are covered with Only the parasitic acanthocephalans reach body a monociliary epidermis. There are about 100 lengths of up to 80 cm. The acanthocephalans described species that are ordered into two have lost many morphological characters as taxa, the elongated Filospermoidea and the adaptations to their parasitic lifestyle, includ- more compact Bursovaginoida (Sterrer 1972 ; ing the jaw apparatus and the digestive tract. Sørensen et al. 2006 ). The animals have a mouth Gnathifera has been placed in the Spiralia, opening that contains the pharyngeal bulb with often affi liated with the Platyzoa (Funch et al. the cuticular jaw structure. Gnathostomulids 2005; Dunn et al. 2008; Witek et al. 2008 , 2009 ; have a blind gut – some species possess a “tem- Hejnol et al. 2009; Wey-Fabrizius et al. 2014 ). poral anus” (Knauss 1979) – an anterior brain, a However, the sister group of Gnathifera remains ventral ganglion that is affi liated with the mouth, unclear. Since gene sequences of most of the and one to three pairs of basiepithelial nerves gnathiferan species seem to evolve fast, it (Kristensen and Nørrevang 1977; Müller and remains diffi cult to resolve with confi dence the Sterrer 2004 ).

Fig. 1.1 Gnathiferan phylogenetic relationships. Phylogenetic relationships of gnathiferan taxa according to recent molecular and morphological studies (Images modifi ed from Funch et al. 2005 ) 1 Gnathifera 3

Early Development Gnathostomulid development needs urgent reinves- tigation with modern methods. The described pres- For only one gnathostomulid species – ence of a spiral cleavage program is unique to the Gnathostomula jenneri – the cleavage pattern has taxon Gnathifera and makes gnathostomulids a key been described (Riedl 1969 ). Eggs (55 μm in diam- taxon for the understanding of the development and eter) are laid by body rupture and start cleaving 2 h evolution of other gnathiferan taxa (Hejnol 2010 ). later (22 °C). The fi rst and second cleavages are “nearly equal” and meridional (Fig. 1.2A, B ). Riedl ( 1969) describes a spiral, dexiotropic arrangement MICROGNATHOZOA of the micromeres that results from the third round of cell divisions (Fig. 1.2C ). The spindle is alter- The freshwater species Limnognathia maerski is nating in angle and thus follows the pattern of a the only micrognathozoan described so far typical spiralian embryo (Fig. 1.2D ). Riedl (1969 ) (Kristensen and Funch 2000 ). With a size of only also described the appearance of an “annelid 100 μm in length, it is one of the smallest metazo- cross” on day 4 of development, which is a specifi c ans. No male individuals have been observed so far. arrangement of blastomeres in spiral embryos (see The females have paired gonads, of which one Chapters 6 , 7 , and 9) but is nowadays not consid- develops a single oocyte at a time. Nothing is ered of phylogenetic signifi cance (see Maslakova known about the reproduction mode, development, et al. 2004 ). Day 4 is also the time when gastrula- or interactions with its habitat, the submerged moss tion is completed, where two larger cells – which of freshwater springs on Disko Island, Greenland. Riedl (1969 ) interprets as the left and right mesen- doblast – are visible (Fig. 1.2E ). The hatchling of G. jenneri is 100 μm in length and lacks the jaws SYNDERMATA but has a primordium of the pharynx (Fig. 1.2F ). The juvenile has a gut, but no lumen is visible. The Syndermata are characterized by their so- called intrasyncytial lamina, a skeletal structure that is located inside the syncytial epidermis (Clément Postembryonic Development and Wurdak 1991 ). The taxon is comprised of three monophyletic groups: the Eurotatoria (comprising During postembryonic development, organ systems the Monogononta and Bdelloidea), the Seisonida, are successively developed until the juvenile is and the parasitic Acanthocephala (Fig. 1.1 ). The about 200 μm long. This is in striking contrast to phylogenetic interrelationship of these taxa is cur- other gnathiferan taxa that have eutely (Riedl 1969 ). rently under debate (see above).

Fig. 1.2 Cleavage program of the gnathostomulid are a view on the animal pole: (A ) two-cell stage, (B ) four- Gnathostomula jenneri after Riedl (1969 ). The naming of cell stage, ( C ) eight-cell stage, (D ) 16-cell stage, and (E ) the blastomeres is according to Riedl ( 1969) who applied approx. 64-cell stage. The shaded blastomeres are the the spiralian nomenclature to the blastomeres. All stages mesendoblasts. (F ) Hatchling with ciliated epidermis 4 A. Hejnol

Rotifera (Clément and Wurdak 1991). Several monogo- nont species have been studied regarding cell Rotifers (“wheel bearers”) are aquatic, tiny numbers (see references in Gilbert 1989). There (0.05–1 mm long) metazoans that are free-living have been several studies regarding the devel- and abundant in the marine and freshwater envi- opment of rotifers, which have mainly focused ronment. There are around 2,000 described spe- on monogonont species (see Table 1.1). Based cies, of which about 200 species live in the marine on the previous descriptions, the early develop- environment. Rotifers use the ciliated rows of ment of monogonont and bdelloid rotifers does cells at the anterior end (corona) for locomotion. not differ much and will be described together in The animals have an anterior brain, a lateral pair the following section. Several authors (Siewing of nerves, and a through gut (except Asplanchna ) 1969 ; Nielsen 2005 ) mention that rotifers have a with an anterior mastax that carries the jaw appa- modifi ed spiral cleavage. ratus. The body has a so-called hemocoel, a fl uid- fi lled cavity that is not surrounded by epithelia Cleavage and is thus not a true coelom. Rotifers deposit an oval, fertilized egg that is pro- Rotifers are divided into two groups, the tected by a resistant shell. In some representa- Monogononta and the Bdelloidea, which mainly tives (Brachionus , Keratella), the female carries differ in their arrangements of the gonads. the fertilized egg until hatching. The embryo Bdelloid rotifers possess two bilaterally arranged gives off one polar body before the onset of gonads, while the monogononts possess only one cleavage, and the position of the polar body gonad that is located medially. Both taxa further- marks the animal pole of the embryo. The fi rst more differ in their mode of reproduction. While cell division is unequal, dividing the egg into a monogononts are either gonochoric (dwarf large blastomere (called CD by some authors), males) or parthenogenetic, bdelloid rotifers have whose descendants will give rise to the germovi- no males and are exclusively parthenogenetic tellarium and other internal tissues. The smaller (Flot et al. 2013 ). Similar to all members of the blastomere – named AB – will mainly contribute Gnathifera, rotifers have direct development and to the ectoderm (Fig. 1.3A ). In all rotifer species no larval stage. As for other groups that have investigated so far, the AB blastomere divides been placed together as “Aschelminthes” (or equally into blastomeres A and B. The blasto- “Nemathelminthes”), rotifers are said to have mere CD divides unequally into the smaller blas- a fi xed number of cells when hatching (eutely) tomere C and the larger blastomere D (Fig. 1.3B ).

Table 1.1 Studies of rotifer development Reference Species Stages and methods Boschetti et al. (2005 ) Macrotrachela quadricornifera Early cleavage (confocal microscopy) (Bdelloidea) Tessin (1886 ) Eosphora digitata (Monogononta) Whole development, light microscopy Zelinka (1892 ) Mniobia ( Callidina ) russeola Whole development, light microscopy, histology (Bdelloidea) Jennings (1896 ) Asplanchna herrickii (Monogononta) Whole development, light microscopy, histology Tannreuther (1919 ) Asplanchna sieboldii (Monogononta) Cleavage and gastrulation of the male Tannreuther ( 1920 ) Asplanchna sieboldii (Monogononta) Whole development, light microscopy, histology Nachtwey (1925 ) Asplanchna priodonta (Monogononta) Whole development, light microscopy, histology Lechner (1966 ) Asplanchna girodi (Monogononta) Whole development, histology, UV ablation experiments Pray (1965 ) Lecane cornuta (Monogononta) Whole development, light microscopy Mrázek (1897 ) Asplanchna (Monogononta) Whole development, light microscopy Car (1899 ) Asplanchna brightwellii (Monogononta) Cleavage, gastrulation, light microscopy 1 Gnathifera 5

The size of blastomere C is similar to that of A mal cells contribute to the internalization by epi- and B. This four-cell stage looks similar to the bolic overgrowth of blastomere D, which gives four-cell stages of unequal cleaving spiralians, off two very small blastomeres that later undergo which also display a larger blastomere (D) and apoptosis (Fig. 1.4B ). During internalization, three smaller blastomeres (Chapter 7 ). The polar blastomere D divides, with one daughter cell body is located in the center of the four blasto- giving rise to the vitellarium and the other to the meres where all blastomeres are in contact with germarium of the adult (Fig. 1.4B ). After immi- each other (Fig. 1.3B ). The polar body marks the gration of the D blastomere, endodermal cells animal pole of the embryo, and interestingly, dur- follow the D blastomere that will form the diges- ing the subsequent cell divisions, bdelloids and tive tract of the adult (Fig. 1.4C). In monogonont monogononts differ in the further spatial interre- embryos, the polar body is transferred by these lationships of blastomeres and polar body. While cellular movements to the future anterior pole of in monogononts the polar body is located fi rst on the embryo. While the origin of the germovitel- the tip of the small blastomeres that are formed larium and the digestive tract has been described until the 16-cell stage (Jennings 1896 ; Nachtwey in rotifers, the origin of mesodermal tissues such 1925 ; Lechner 1966 ), the polar body in the bdel- as musculature and nephridia remains unknown loid species is located closely to the large blasto- (Fig. 1.4D ). Only a gross description of the fi nal meres that will gastrulate (Zelinka 1892 ). This fates of the early blastomeres is currently avail- difference seems to be fundamental at fi rst able for rotiferans (Fig. 1.5 ). glance, since many nomenclatures, e.g., the spi- ralian nomenclature developed by Conklin Organogenesis (1897 ), use the position of blastomeres in relation Endodermal cells form the tube of the digestive to the polar body for the naming of the cells. tract and differentiate into the stomach, also Following this rule, in monogononts the large called mastax (Fig. 1.4D ). The former blastopore cell that forms the germovitellarium would be a forms the opening that differentiates into the vegetal macromere, while in bdelloids this mouth on the anterior ventral side (Fig. 1.4D ). homologous blastomere would constitute an ani- The ectoderm on the dorsal side undergoes more mal micromere. Consequently, the names of the cell proliferation and will give rise to the cerebral individual homologous cells would change dra- ganglion of the adult. These cells will get inter- matically. However, since the fate of the individ- nalized from the dorsal ectoderm to form the ual blastomeres is comparable between both brain of the adult (Fig. 1.4D ). Future epidermis rotifer taxa and in both groups the polar body will cells close the immigration site and differentiate eventually lie at the future anterior pole of the into a syncytial integument. At this stage, the dif- embryo, the polar body cannot be used for nam- ferentiation of the major parts of the digestive ing the cells in the rotifer embryo, and thus other tract (mouth opening, pharynx, and mastax) is criteria, such as the fate of the blastomeres, have visible. In some embryos, the formation of an to be applied. oviduct has been described (Lechner 1966 ). The In both embryos, however, the four blasto- last phase of organogenesis begins with the com- meres form tiers, which may be called quadrants, plete closure of the site of immigration of the since they are descendants of the blastomeres of cerebral ganglion precursors. The cilia of the cili- the four-cell stage and lie in one row with the ary bands start beating and the pseudocoel mother cells (Fig. 1.3C, D ). becomes visible. Before hatching, the juvenile extends along the anterior-posterior axis and Gastrulation hatches. The hatchling possesses a functional Gastrulation in rotifers begins with the internal- excretory system with protonephridia. Due to the ization of the large blastomere D at the 16-cell eutelic condition, growth is achieved by exten- stage (Fig. 1.4A ). This is consistent between all sion of cell volume and not by further cell rotifer species investigated so far. The ectoder- proliferation. 6 A. Hejnol

Fig. 1.3 Rotifer cleavage up to the 16-cell stage. (A ) (1919 ) and Lechner ( 1966). The future anterior pole of the Two-cell stage. Polar body shaded in red . (B ) Four-cell embryo is marked with an asterisk . ( E ) 10-cell stage of the stage. ( C ) Eight-cell stage with blastomere D that will bdelloid Callidina (Zelinka 1892 ) which shows reverse form the germovitellarium shaded in brown . (D ) 16-cell polarity. Note the polar body close to the site of gastrula- stage of the monogonont Asplanchna after Tannreuther tion that also corresponds to the future anterior pole

Experimental Embryology cells, and cells of the digestive tract. Depending The only study using experimental approaches to on the degree and timepoint of the irradiation – investigate rotifer embryos is the work of Lechner the work does not clearly state the duration and (1966 ) on Asplanchna . He used UV irradiation to target – the embryo either does not gastrulate, destroy early blastomeres during cleavage. The arrests after gastrulation, or even manages to results are either a complete destruction of the hatch out of the egg shell with internal organs embryo or – results after less or partial irradia- nearly intact. Lechner’s (1966 ) series of experi- tion – partial differentiation of the embryo. After ments indicates that cell fates are determined partial ablation of individual blastomeres, the early and that blastomeres can differentiate into remaining cells were still able to differentiate to their fi nal fates without induction of functional late cellular fates, such as ciliated cells, epidermal neighbor cells. The rotifer embryo can thus be 1 Gnathifera 7

Fig. 1.4 Rotifer gastrulation. Gastrulation summarized from blastomere D before internalization are shaded in based on studies of different rotifer species ( Asplanchna dark brown . ( C ) Late gastrulation stage after further endo- girodi , Asplanchna priodonta , Callidina russeola ). (A ) dermal cells have immigrated through the blastopore (bp ). Blastomere D ( shaded in dark brown ) before internaliza- The polar body ( pb , shaded in red ) is closer to the future tion at the future anterior pole of the embryo. ( B ) The anterior pole ( asterisk ). ( D) Later stage with organ sys- large blastomere has divided into the germarium precur- tems already present. The mouth opening ( mo ) is anterior. sor (ger , shaded in light brown ) and the blastomere that The cells that will form the cerebral ganglion (cg ) have will form the vitellarium ( vit , shaded in brown gray ). been internalized. The gut lumen (gl ) is visible Small, degenerating blastomeres that have been given off 8 A. Hejnol

a 4,2

A3 a 5,2 a 4,1 Epibolic growth a 6,2 during gastrulation: a 5,1 a 7,2 ectoderm and endoderm a 6,1 a 7,1

b 4,2 anterior and dorsal ectoderm B3 b 5,2 stomach and stomach glands b 4,1 b 6,2 b 5,1 b 7,2 b 6,1 stomach b 7,1

c 4,2

c3 c 5,2

c 4,1 c 6,2 ectoderm, endoderm c 5,1 c 7,2 c 6,1 c 7,1

d 4,2 anterior and D3 d 5,2 ventral ectoderm

d 4,1 d 6,2 d 5,1 d 7,2 d 6,1 Germovittelarium d 7,1

Fig. 1.5 Rotifer fate map. The fate map of the monogonont Asplanchna girodi reconstructed after Lechner (1966 ) seen as a classic “mosaic” embryo in which cells rotifer Brachionus plicatilis. The fi rst ubiq- can differentiate autonomously. However, more uitously expressed vasa ortholog becomes detailed and sophisticated experiments are neces- restricted to the germovitellarium line and is later sary to characterize the underlying molecular expressed weakly in the vitellarium and stronger mechanisms and early inductional processes that in the oocytes and germ cells (Smith et al. 2010 ). are likely to be active also in this embryo. The same has been found for the nanos ortholog (Smith et al. 2010 ). The recent sequencing of the Molecular Approaches fi rst rotifer genome of the bdelloid Adineta vaga Molecular studies on rotifer development are (Flot et al. 2013 ) and reports of RNA interference rare. Smith and coworkers (Smith et al. 2010 ) in a monogonont rotifer (Snell et al. 2011 ) can studied the gene expression of the germ line provide tools to study the molecular mechanisms markers vasa and nanos in the monogonont of rotifer development in more detail. Here, bdel- 1 Gnathifera 9 loid rotifers provide a unique system to study The oval-shaped, fertilized egg has two polar the reduced tetraploidy (Flot et al. 2013 ) and bodies that mark the future anterior pole of the the multiplication of many key regulatory genes animal (Fig. 1.6A ). The fi rst cell division is equa- and signaling pathways and their impact on the torial to the longitudinal egg axis and gives rise to development of an embryo. The bdelloid species the animal blastomere AB and the slightly larger Adineta vaga has many more homeobox gene vegetal blastomere CD (Fig. 1.6B ). The two blas- families with four copies than tetraploid verte- tomeres divide equally in the next cell division brates; this could be explained by the gene con- round and produce the animal blastomere B3, the version that these animals undergo to eliminate median blastomeres A3 and C3, and the vegetal mutations under the absence of sexuality (Flot blastomere D3 (Fig. 1.6C ). The following cell et al. 2013 ). The Hox gene cluster is atomized divisions are unequal. Blastomere D3 is the fi rst and a posterior Hox gene is missing. to divide unequally into the micromere D4.1 and the macromere D4.2. The other macromeres fol- low the same division pattern in that they give off Seisonida smaller micromeres in the direction of the polar bodies. In different species, the embryo becomes The development of this peculiar rotifer lineage syncytial at different timepoints, ranging from has not been studied. The separate sex adults the four-cell to the 36-cell stage. A clear fate of deposit fertilized eggs close to their attachment the blastomeres has not been determined. site directly on the host the leptostracan crusta- Interestingly, up to the 25-cell stage of cean Nebalia . Only two species have been Gigantorhynchus gigas (Meyer 1928 ), only mac- described, Seison nebaliae and Seison annulatus . romeres divide – the micromeres are arrested. The quartet of macromeres forms thus “mother cells” of all micromeres before the micromeres Acanthocephala start to divide again in the 25-cell stage. Previous studies agree that during further devel- Acanthocephalans are a gnathiferan taxon that opment the micromeres condense and become is exclusively parasitic, and all 1,100 species internalized to form the “central nuclear mass” described show many adaptations to the parasitic (Fig. 1.6G ). Several authors described this as a form lifestyle (Meyer 1933 ). These wormlike animals of gastrulation, since these condensed nuclei will be live as adults in the digestive system of vertebrates part of all the internal tissues except the epidermis. (fi nal host) and take up nutrients through the integ- The embryo begins to form additional – up to four – ument. Acanthocephalans have a reduced diges- egg layers of different density that are composed tive tract and evolved reproductive strategies that out of polysaccharides and other material. allow them to infest the intermediate host (crusta- ceans) and fi nal host (Fig. 1.6 ). Acanthocephalans Acanthocephalan Life Cycle have separate sexes, males and females copulate, The acanthor is the infectious stage that pos- and eggs are fertilized internally. sesses a boring organ (aclid organ) that is used to penetrate the gut of the intermediate host (crusta- Development cean). The acanthor is fi rst surrounded by several The development of acanthocephalans has been membranes, and it only hatches when it has been studied in about ten species by several authors taken up by the intermediate host (Fig. 1.6I ). The (Schmidt 1985 ). The most detailed description of acanthor – after penetrating the gut and enter- the cell lineage of the embryo that gives rise to an ing the mesenteron – can be silent for hours or early stage called the acanthor (often called days. The acanthor begins to grow in size and “larva”), which then infests the intermediate host, the organs of the worm are formed by the cells has been conducted by Hamann (1891 ), Kaiser in the membranes, thus forming the acanthella (1893 ), and Meyer (1928 , 1933 ). (Fig. 1.6J ). In the acanthella, the brain begins 10 A. Hejnol 1 Gnathifera 11 to form, and the inner cell mass starts to form Ahlrichs WH (1997) Epidermal ultrastructure of Seison musculature (retractor muscle) and ovaries or tes- nebaliae and Seison annulatus, and a compari- son of epidermal structures within the Gnathifera. tes. Finally, hooks are formed on the proboscis. Zoomorphology 117:41–48 The worm – now called cystacanth – possesses Boschetti C, Ricci C, Sotgia C, Fascio U (2005) The all structures of the adult and is ready to infect development of a bdelloid egg: a contribution after the main host when the arthropod infested with 100 years. Hydrobiologia 546:323–331. doi: 10.1007/ S10750-005-4241-Z a cystacanth is consumed by the vertebrate host. Car L (1899) Die embryonale Entwicklung von Taken together, the data currently available on Asplanchna brightwellii . Biol Zentbl 19:59–74 gnathiferan ontogeny show that all species inves- Clément P, Wurdak E (1991) Rotifera. In: Harrison FW, tigated to date are direct developers. Some of the Ruppert EE (eds) Microscopic anatomy of inverte- brates. Wiler Liss, New York, pp 219–297 taxa are eutelic (Eurotatoria), while others show Conklin EG (1897) The embryology of Crepidula . J postembryonic development. Acanthocephalans Morphol 13:1–226 have evolved a complicated life cycle that allows Dunn CW, Hejnol A, Matus DQ et al (2008) Broad phy- the species to transfer between hosts and repro- logenomic sampling improves resolution of the ani- mal tree of life. Nature 452:745–749. doi: 10.1038/ duce in masses. The cleavage pattern differs nature06614 , nature06614 [pii] between groups. While spiral cleavage has been Flot JF, Hespeels B, Li X et al (2013) Genomic evidence described in gnathostomulids, this has so far not for ameiotic evolution in the bdelloid rotifer Adineta been confi rmed for rotifers or acanthocephalans. vaga . Nature 500:453–457. doi: 10.1038/nature12326 Funch P, Sørensen MV, Obst M (2005) On the phyloge- netic position of Rotifera – have we come any further? Hydrobiologia 546:11–28 OPEN QUESTIONS Gilbert JJ (1989) Rotifera. In: Adiyodi KG, Adiyodi RG (eds) Reproductive biology of invertebrates. Wiley, Chichester, pp 179–199 • Detailed cell lineage studies in all gnathiferan Hamann O (1891) Monographie der Acanthocephalen taxa (Echinorhynchen). Jena Z Naturw 25:113–231 • Organogenesis in all major taxa Hejnol A (2010) A twist in time – the evolution of spi- • The basic molecular mechanisms that trigger ral cleavage in the light of animal phylogeny. Integr Comp Biol 50:695–706. doi: 10.1093/icb/icq103 the development of gnathiferan groups Hejnol A, Obst M, Stamatakis A et al (2009) Assessing • Developmental gene expression of all major the root of bilaterian animals with scalable phyloge- gene families in all gnathiferan subtaxa nomic methods. Proc R Soc Ser B 276:4261–4270. doi: 10.1098/rspb.2009.0896 Jennings HS (1896) The early development of Asplanchna herrickii De Guerne. Bull Mus Comp Zool 30:1–117 Kaiser J (1893) Die Acanthocephalen und ihre References Entwickelung. Bibl Zool 2:1–374 Knauss E (1979) Indication of an anal pore in Ahlrichs WH (1995) Ultrastruktur und Phylogenie von Gnathostomulida. Zool Scr 8:181–186 Seison nebaliae (Grube 1859) und Seison annulatus Kristensen RM, Funch P (2000) Micrognathozoa: a new (Claus 1876). PhD thesis, Georg August University class with complicated jaws like those of Rotifera and Göttingen Gnathostomulida. J Morphol 246:1–49

Fig. 1.6 Acanthocephalan development and life cycle. mass is established and an additional layer has been All embryonic stages are oriented with the polar bodies formed around the embryo. ( I ) Hatched acanthor from the ( pb) up; cleavage pattern of Giganthorhynchus gigas after arthropod host. Hooks are visible at the anterior pole. Meyer ( 1928 ). (A ) Fertilized zygote. (B ) Two-cell stage. Central cell mass is still undifferentiated. ( J ) Developed ( C ) Four-cell stage. (D ) Eight-cell stage. (E ) 13-cell stage. acanthella (male). The central cell mass has formed ( F ) 17-cell stage. (G ) Syncytial embryo with multiple retractor muscles (rm ) and the gonads (te , testes). (K ) Life cells. The central nuclear mass ( cnm ) begins to form by cycle of the acanthocephalan Echinorhynchus gadi (for immigration of nuclei at the future anterior pole of the description, see text) embryo. ( H ) Later stage embryo. The central nuclear 12 A. Hejnol

Kristensen RM, Nørrevang A (1977) On the fi ne structure Smith JM, Cridge AG, Dearden PK (2010) Germ cell spec- of Rastrognathia macrostoma gen. et sp.n. placed in ifi cation and ovary structure in the rotifer Brachionus Rastrognathiidae fam.n. (Gnathostomulida). Zool Scr plicatilis . EvoDevo 1:5. doi: 10.1186/2041- 6:27–41 9139-1-5 Lechner M (1966) Untersuchungen zur Embryonalentwicklung Snell TW, Shearer TL, Smith HA (2011) Exposure to des Rädertieres Asplanchna girodi De Guerne. Roux’ dsRNA elicits RNA interference in Brachionus man- Arch f Entwicklungsmech 157:117–173 javacas (Rotifera). Mar Biotechnol (NY) 13:264–274. Maslakova SA, Martindale MQ, Norenburg JL (2004) doi: 10.1007/s10126-010-9295-x Fundamental properties of the spiralian developmental Sørensen MV (2003) Further structures in the jaw appara- program are displayed by the basal nemertean Carinoma tus of Limnognathia maerski (Micrognathozoa), with tremaphoros (Palaeonemertea, Nemertea). Dev Biol notes on the phylogeny of the Gnathifera. J Morphol 267:342–360. doi: 10.1016/j.ydbio.2003.10.022 255:131–145 Meyer A (1928) Die Furchung nebst Eibildung, Reifung Sørensen MV, Sterrer W, Giribet G (2006) Gnathostomulid und Befruchtung des Gigantorhynchus gigas. Zool Jb phylogeny inferred from a combined approach of four Anatomie 50:117–218 molecular loci and morphology. Cladistics 22:32–58 Meyer A (1933) Acanthocephala. Akademische Sterrer W (1972) Systematics and evolution within the Verlagsgemeinschaft, Leipzig Gnathostomulida. Syst Zool 21:151–173 Mrázek A (1897) Zur Embryonalentwicklung der Tannreuther GW (1919) Studies on the rotifer Asplanchnia Gattung Asplanchna . Sitz-Ber Kgl Böhm Gesell Wiss ebbesbornii, with special reference to the male. Biol 58:1–11 Bull 37:194–207 Müller MCM, Sterrer W (2004) Musculature and Tannreuther GW (1920) The development of Asplanchna nervous system of Gnathostomula peregrina ebbesbornii (Rotifer). J Morphol 33:389–419 (Gnathostomulida) shown by phalloidin labeling, Tessin G (1886) Über Eibildung und Entwicklung der immunohistochemistry, and cLSM, and their phyloge- Rotatorien. Z Wiss Zool 44:273–302 netic signifi cance. Zoomorphologie 123:169–177 Wey-Fabrizius AR, Herlyn H, Rieger B, Rosenkranz Nachtwey R (1925) Untersuchungen über die Keimbahn, D, Witek A, Welch DBM, Ebersberger I, Hankeln T Organogenese und Anatomie von Asplanchna pri- (2014) Transcriptome data reveal Syndermatan rela- odonta Gosse. Z Wiss Zool 126:239–492 tionships and suggest the evolution of endoparasitism Nielsen C (2005) Trochophora larvae: cell-lineages, in Acanthocephala via an epizoic stage. PLoS One ciliary bands and body regions. 2. Other groups and 9:e88618. doi: 10.1371/journal.pone.0088618 general discussion. J Exp Zool B Mol Dev Evol Witek A, Herlyn H, Meyer A, Boell L, Bucher G, Hankeln 304:401–447. doi: 10.1002/jez.b.21050 T (2008) EST-based phylogenomics of Syndermata Pray F (1965) Studies on the early development of the questions monophyly of Eurotatoria. BMC Evol Biol rotifer Monostyla cornuta MÜLLER. Trans Am 8:345. doi: 10.1186/1471-2148-8-345 Microsc Soc 84:1965 Witek A, Herlyn H, Ebersberger I, Mark Welch DB, Riedl RJ (1969) Gnathostomulida from America. Science Hankeln T (2009) Support for the monophyletic 163:445–452 origin of Gnathifera from phylogenomics. Mol Schmidt G (1985) Development and life cycles. In: Crompton Phylogenet Evol 53:1037–1041. doi: 10.1016/j.ympev. D, Nickol B (eds) Biology of the Acanthocephala. 2009.07.031 Cambridge University Press, Cambridge, pp 273–305 Zelinka C (1892) Studien über Räderthiere. III. Zur Siewing R (1969) Lehrbuch der vergleichenden Entwicklungsgeschichte der Räderthiere nebst Entwicklungsgeschichte der Tiere. Verlag Paul Parey, Bemerkungen über ihre Anatomie und Biologie. Z Hamburg/Berlin Wiss Zool 53:1–159 Gastrotricha 2 Andreas Hejnol

Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger.

A. Hejnol Sars International Centre for Marine Molecular Biology , University of Bergen , Thormøhlensgate 55 , 5008 Bergen , Norway e-mail: [email protected]

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 13 DOI 10.1007/978-3-7091-1871-9_2, © Springer-Verlag Wien 2015 14 A. Hejnol

INTRODUCTION The brain is connected via lateral axon tracts or neurite bundles to the posterior end of the Gastrotricha is a clade of aquatic microscopic body (Schmidt-Rhaesa 2015 ). The alimentary animals that are among the smallest metazoans. canal has an anterior mouth with a muscular Gastrotrichs mainly live in the marine and fresh- pharynx, a midgut that is composed of less than water interstitial environment and on the surface 100 cells and ends with a posterior anal opening of aquatic plants. There are around 700 described (Ruppert 1991 ). Most species lack an ectodermal species of gastrotrichs, which are divided into the hindgut and some even lack an anal opening marine taxon Macrodasyida and the marine and (Urodasys ). No specialized hemal or respiratory freshwater taxon Chaetonotida. So far, no fossils system is present in gastrotrichs, but the excretory have been assigned to the taxon Gastrotricha. All system is composed of one to six pairs of gastrotrichs are direct developers, and the adults protonephridia (Ruppert 1991 ). Gastrotrichs pos- measure between 60 and 600 μm, with some sess characteristic adhesive tubes that are com- exceptions growing to a size of several millime- posed of two to three gland cells that secrete ters. The acoelomate body is wormlike, bilaterally adhesive substances (“dual-gland system”). symmetric, and sometimes covered with scales Gastrotrichs can be dioecious or protandric her- and a cuticle (Fig. 2.1; Ruppert 1991). The ventral maphrodites, and some obligate parthenogenetic side is covered with locomotory cilia (“gaster,” species have been described (Hummon 1974 ). belly; “thrix,” hair). The animals have a brain that Fertilization is internal and eggs are deposited is mainly composed of an elaborated dorsal through a female opening or body rupture. The commissure, whereas a ventral commissure is phylogenetic position of Gastrotricha is still under present in some species (Schmidt-Rhaesa 2015 ). debate, and affi liations with the Platyzoa,

Fig. 2.1 Live adult gastrotrichs. (A ) The macrodasyoid All Rights Reserved). mo mouth opening, ph pharynx, at Macrodasys sp. ( B) The chaetonotoid Halichaetonotus sp . adhesive tube, gu gut, go gonad Scale bar: 50 μm (© Andreas Schmidt-Rhaesa 2015 . 2 Gastrotricha 15 a suggested monophylum within the Spiralia, along the animal-vegetal axis and gives rise to the have been hypothesized based on the results of animal blastomere AB and the vegetal blasto- large- scale phylogenomic analyses (Dunn et al. mere CD (Fig. 2.2A ). In the next division round, 2008 ; Hejnol et al. 2009 ). AB is the fi rst that divides slightly unequally into the smaller blastomere A and the larger blasto- mere B, followed by the division of CD into the EARLY AND LATE DEVELOPMENT vegetal blastomeres C and D (Fig. 2.2B ). After these divisions, the animal blastomeres are posi- There are only a few studies of gastrotrich devel- tioned about 90° twisted to the vegetal ones opment. Besides the most comprehensive studies (Fig. 2.2B ). In the next round of cell division to of the chaetonotoid species Lepidodermella squa- the eight-cell stage, all blastomeres divide per- mata by Sacks ( 1955 ) and the macrodasyoid spe- pendicular to the animal-vegetal axis. The fi rst cies Turbanella cornuta by Teuchert ( 1968 ), other cell to divide is AB, followed by CD (Fig. 2.2C ). data are very scarce (de Beauchamp 1929 ; Brunson The four animal blastomeres are located in the 1949; Swedmark 1955 ). Gastrotrich development furrows between the four vegetal blastomeres. In has been video recorded (Teuchert 1975 ; Schnabel the subsequent division rounds, the animal blas- et al. 2006 ), and the results confi rm that the early tomeres cleave before the vegetal blastomeres development is highly stereotypic (Schnabel et al. (Fig. 2.2D ). The 32-cell stage has a blastocele, 2006 ). So far, no developmental gene expression and gastrulation starts with the immigration of studies have been published. two sister cells, A5.3 and A5.4 (Fig. 2.2E ; Sacks Eggs are deposited with an egg shell that is 1955 ). The animal pole of the embryo will form smooth in marine species and can contain spines the anterior region of the juvenile. The site where or other appendages in the freshwater species. the two cells entered the blastocele will later The fertilized eggs are attached to sand grains or form the stomodeum of the adult (Fig. 2.2F ). to algae with the use of secretory glands of the During later development, a posterior opening is adult. Both gastrotrich groups differ regarding formed that gives rise to the proctodeum their development in terms of duration and their (Fig. 2.2F, G ). The organ systems begin to form cleavage patterns. 6 h after egg deposition, and the embryo under- goes an extension, which bends the future juve- nile inside the egg shell (Fig. 2.2H ). The Development of Chaetonotoida endodermal gut connects the anterior stomodeum and the posterior proctodeum to each other. The most detailed description of chaetonotoid A pharynx is clearly visible after 9 h of oviposi- development is that of the parthenogenetic tion. Cell differentiation of epidermal scales, freshwater species Lepidodermella squamata cilia, gut cells, and caudal forks occurs between 9 (Sacks 1955) that confi rmed the partial descrip- and 23 h of development. The juvenile hatches tions of the early and late cleavage made by after 23–31 h. Ludwig (1875 ), de Beauchamp ( 1929 ), and Brunson (1949 ). The cleavage pattern of Lepidodermella squa- Development of Macrodasyoida mata is similar to that of Neogossa antennigera (de Beauchamp 1929 ). Eggs are deposited as fer- The literature about macrodasyoid gastrotrich tilized zygotes and with an egg shell. At 22 °C development is very scarce. The main work stems the second meiotic division occurs after egg from Teuchert (1968 , 1975 ) on several species, deposition, and a polar body is extruded and with the main description of Turbanella cornuta . localizes to the longitudinal end of the oval egg, A short note on Macrodasys had been published marking the animal pole of the embryo earlier (Swedmark 1955). The development in (Fig. 2.2A ). The fi rst cell division is equal and macrodasyoid gastrotrichs takes longer than in 16 A. Hejnol

Fig. 2.2 Development of the chaetonotoid Lepidodermella squamata after Sacks (1955 ) and the macrodasyoid Turbanella cornuta after Teuchert (1968 ). Comparative stages of the embryos of L. squamata ( left ) and T. cornuta ( right ). Animal pole to the top , ventral to the left . ( A , A ’) Two-cell stage. ( B , B ’) Four-cell stage. ( C , C ’) Eight-cell stage. ( D , D ’) 16-cell stage. ( E , E ’) Gastrulation. ( F , F ’) Stomodeum formation. (G , G ’) Digestive tract visible. ( H , H ’) Late stage of a juvenile in the egg shell. bp blastopore, st stomo- deum, pr proctodeum. Identically colored blastomeres indicate their homology and demonstrate the differences between the cleavage patterns of the two species 2 Gastrotricha 17 the investigated chaetonotoid species. Turbanella The mesodermal bands will likely form the cornuta develops in 3–4 days until hatching, and musculature of the adult. The anterior ectoderm Macrodasys caudatus up to 10 days (Teuchert proliferates and ectodermal cells will contribute 1968 ). The eggs are fertilized internally, and two to the brain. Cell differentiation and organ system polar bodies are extruded sequentially after egg formation begins 50–55 h after egg deposition. deposition at the animal pole of the embryo The juvenile hatches after 4 days at 22 °C. (Fig. 2.2A ’). The fi rst cleavage is equatorial and equal and divides the embryo into the animal blastomere AB and the vegetal blastomere CD Comparison Between Chaetonotoid (Fig. 2.2A ’). In the next cleavage round, blasto- and Macrodasyoid Early mere AB divides before CD. Both cells divide Development meridionally, but the cleavage planes are perpen- dicular to each other. In the early four-cell stage, The described cleavage patterns for chaetonotoid the animal blastomeres A and B lie between the gastrotrichs are consistent, and only minor dif- vegetal blastomeres C and D (Fig. 2.2B ’). After ferences have been detected (Sacks 1955 ). The the cell divisions, one of the vegetal blastomeres, differences between chaetonotoid and macro- C, moves toward the animal pole, leaving blasto- dasyoid gastrotrichs are more fundamental mere D at the vegetal pole (Fig. 2.2B ’). According (Fig. 2.2 ). Teuchert points out that one of the to Teuchert (1968 ), C exclusively forms ecto- major differences between the embryos of both derm, while D – besides some mesodermal and taxa is the origin of the cells that form the stomo- ectodermal portions – will give rise to all endo- deum (Teuchert 1968). In chaetonotoids, the ani- derm. After the shift of the C blastomere, the mal (“oral”) cells form the blastopore and the embryo begins the next round of divisions. The endoderm, while in macrodasyoids, the endo- animal blastomeres divide fi rst along the animal- derm and blastopore are formed from the vegetal vegetal axis, and the dorsally located cell C cells (“caudal,” i.e., posterior). This fundamental divides along the same axis (Fig. 2.2C ’). The last difference led Teuchert to turn Sacks’ descrip- cell to divide in this round is the ventral blasto- tions of the Lepidodermella squamata embryo mere D. In the next round of divisions, all cells 180°, so that in both embryos the endoderm divide perpendicular to the previous cleavage seemingly originates from the vegetal pole plane (Fig. 2.2D ’). At the 30-cell stage, 11–12 h (Teuchert 1968 ). However, video recordings of after egg deposition, two ventral descendants of L. squamata development (Schnabel et al. 1997 ) blastomere D begin to immigrate into the blasto- show that indeed Sacks’ ( 1955) observations are cele and will later form the endoderm (Fig. 2.2E ’). correct. Other differences in the development of According to Teuchert ( 1968), the mesodermal both groups concern the angle during early cleav- cells follow after the cells have undergone age (see different arrangements in Fig. 2.2 ) and another division. The other blastomere identities how the animal blastomeres get shifted against and the later cell divisions in the T. cornuta the vegetal ones in the early embryo. In macro- embryo remain unclear. The blastopore elon- dasyoid embryos, the spindles place the blasto- gates, closes from the posterior end, and forms meres between the vegetal ones, while in the opening of the stomodeum (Fig. 2.2F ’). The chaetonotoid embryos the cells shift after the embryo is now 40–50 h old. The proctodeum will placement on top of the sister cell. be formed later at a new site at the posterior tip of In both groups, the entire endoderm seems to the extended embryo (Fig. 2.2G ’). be formed by two gastrulating cells. The meso- According to Teuchert (1968 ), mesodermal derm is formed by precursors that gastrulate bands are formed by mesodermal precursors that later – possibly at different positions in the gastrulate after the two endodermal precursors. embryo. 18 A. Hejnol

ORGANOGENESIS OPEN QUESTIONS

Organogenesis in gastrotrichs has not been described in detail. Sacks’ work about • Reinvestigation of macrodasyoid develop- Lepidodermella squamata describes the forma- ment regarding fate map and cell lineage tion of the digestive tract (Fig. 2.2; Sacks 1955). • Virtually all aspects of organogenesis, No data about mesoderm or nervous system including neuromuscular development development is available for a chaetonotoid spe- • Molecular characterization including gene cies. Teuchert (1968 ) provides data about the expression patterns of basic developmental development of some organ systems in the mac- processes, cell type differentiation by investi- rodasyoid Turbanella cornuta . The brain is gation of transcription factors, signaling formed by ectodermal thickenings at the anterior cascades, and structural genes in both gastro- end of the animal (Fig. 2.2 ). Cells proliferate trich groups from the epidermis to the inside of the embryo and become arranged in a crescent-shaped dorsal commissure above the pharynx. The T. cornuta References embryo has one pair of protonephridia anlagen, which get multiplied in a serial arrangement after Brunson R (1949) The life history and ecology of two hatching (Teuchert 1968 ). The pharynx is formed North American gastrotrichs. Trans Am Microsc Soc 68:1–20 by two cell rows that will form a tube composed de Beauchamp P (1929) Le développement des of 70–80 cells. The lumen-less gut is composed Gastrotriches. Bull Soc Zool Fr 54:549–558 of approximately 20 large cells that are arranged Dunn CW, Hejnol A, Matus DQ, Pang K, Browne WE, in a row. During later development of the embryo, Smith SA, Seaver E, Rouse GW, Obst M, Edgecombe GD, Sorensen MV, Haddock SH, Schmidt-Rhaesa A, a small lumen becomes visible. The mesoderm is Okusu A, Kristensen RM, Wheeler WC, Martindale composed of two lateral bands. Since gastro- MQ, Giribet G (2008) Broad phylogenomic sampling trichs are acoelomate, these cells will mainly improves resolution of the animal tree of life. Nature give rise to the musculature of the animal. 452:745–749 Hejnol A, Obst M, Stamatakis A, Ott M, Rouse GW, Teuchert ( 1968) also mentions that it is possible Edgecombe GD, Martinez P, Baguñá J, Bailly X, that posterior cells of the mesodermal bands Jondelius U, Wiens M, Müller WEG, Seaver E, might form the gonads. The cells of the lateral Wheeler WC, Martindale MQ, Giribet G, Dunn CW mesodermal bands proliferate and fi ll the space (2009) Assessing the root of bilaterian animals with scalable phylogenomic methods. Proc Roy Soc Ser B on the dorsal and ventral sides of the gut. 276:4261–4270 Teuchert ( 1968) assumes that these cells will Hummon W (1974) Gastrotricha. In: Giese A, Pearse J form the longitudinal dorsal and ventral muscu- (eds) Reproduction of marine invertebrates. Academic, lature. Another mesodermal derivative that is New York Ludwig H (1875) Ueber die Ordnung Gastrotricha specifi c to macrodasyoid gastrotrichs is the so- Metschn. Z Wiss Zool 16:193–225 called “Y-organ” of unknown function (Ruppert Ruppert EE (1991) Gastrotricha. In: Harrison FW, 1991 ; Teuchert 1968 ). Ruppert EE (eds) Microscopic anatomy of inverte- The differences between both groups and the brates: aschelminthes. Wiley-Liss, New York Sacks M (1955) Observations on the embryology of an overall paucity of data on gastrotrich develop- aquatic gastrotrich, Lepidodermella squamata ment make it diffi cult to reconstruct the ancestral (Dujardin, 1841). J Morphol 96:473–495 mode of development for gastrotrichs and at Schmidt-Rhaesa A (2015) Gastrotricha. In: Schmitdt- present render detailed comparisons with other Rhasa A, Harzsch S, Purschke G (eds) Structure and evolution of invertebrate nervous systems. Oxford larger taxonomic units diffi cult. University Press, Oxford (in press) 2 Gastrotricha 19

Schnabel R, Hutter H, Moerman D, Schnabel H (1997) Swedmark B (1955) Développment d’un Gastrotriche Assessing normal embryogenesis in Caenorhabditis Macrodasyoide, Macrodasys affi nis Remane. CR elegans using a 4D microscope: variability of Acad Sci Paris 240:1812–1814 development and regional specifi cation. Dev Biol Teuchert G (1968) Zur Fortpfl anzung und Entwicklung 184:234–265 der Macrodasyoidea (Gastrotricha). Z Morphol Tiere Schnabel R, Bischoff M, Hintze A, Schulz AK, Hejnol A, 63:343–418 Meinhardt H, Hutter H (2006) Global cell sorting in Teuchert G (1975) Organisation und Fortpfl anzung von the C. elegans embryo defi nes a new mechanism for Turbanella cornuta (Gastrotricha). Film C1176. pattern formation. Dev Biol 294:418–431 Institut für den wissenschaftlichen Film, Göttingen Platyhelminthes 3 Teresa Adell , José M. Martín-Durán , Emili Saló , and Francesc Cebrià

Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger. T. Adell • E. Saló (*) • F. Cebrià Department of Genetics, Faculty of Biology , Institute of Biomedicine, University of Barcelona , Av. Diagonal 643, edifi ci Prevosti, planta 1 , Catalunya, Barcelona 08028 , Spain e-mail: [email protected] J. M. Martín-Durán Sars International Centre for Marine Molecular Biology, University of Bergen , Thormøhlensgate, 55 , Bergen 5008 , Norway

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 21 DOI 10.1007/978-3-7091-1871-9_3, © Springer-Verlag Wien 2015 22 T. Adell et al.

INTRODUCTION (marine fl atworms) and terrestrial species usually display bright colours and patterns. Molecular The phylum Platyhelminthes comprises dor- phylogenetic studies place the Platyhelminthes soventrally fl attened worms commonly known within the Spiralia clade. The most recent inter- as fl atworms (from the Greek platys , meaning nal phylogenies support the subdivision of the fl at, and helminthos , meaning worm) (for a gen- Platyhelminthes into two main groups: the earliest eral overview of this phylum, see Hyman 1951 ; branching lineages grouped into the paraphyletic Rieger et al. 1991 ). Platyhelminthes are one of ‘Archoophora’ and the more divergent mono- the largest animal phyla after arthropods, mol- phyletic Neoophora (Laumer and Giribet 2014 ; lusks, and chordates and includes more than Riutort et al. 2012). The ‘Archoophora’ includes 20,000 species, more than half of which are para- those groups with endolecithal eggs. They are sitic fl atworms. Free-living fl atworms (classically exclusively free-living organisms and are classi- referred to as ‘Turbellaria’) live in a large vari- fi ed into three orders: Catenulida, Polycladida, ety of habitats, from freshwater springs, rivers, and Macrostomida (Fig. 3.1 ). The Neoophora lakes, and ponds to the ocean and moist terres- includes all groups with ectolecithal eggs. It com- trial habitats. Their size ranges from microscopic prises several free-living orders, together with the worms to the 30 m long tapeworms found in the parasitic groups (the classes Trematoda, Cestoda, sperm whale. Free-living fl atworms are most and Monogenea) united under the monophyletic often white, brown, grey, or black; polyclads Neodermata.

Catenulida

Polycladida Catenulida (Stenosum sthenum)

Archoophora Macrostomorpha

Polycladida (Prosthiostomum siphunculus) Prorhynchida Lecithoepitheliata Gnosonesimida

Macrostomorpha (Marcostomum lignano) Proseriata

Bothrioplanida

Prothynchida (Geocentrophora sphyrocephala) Neodermata Trematoda, Cestoda, Monogenea

Neoophora Proseriata (Monocelis fusca) Rhabdocoela

Fecampiida Rhabdocoela (Rhynchomesostoma rostratum) Tricladida

Prolecithophora Tricladida (Procerodes littoralis)

Fig. 3.1 Phylogenetic consensus tree of Platyhelminthes. new orders (Prorhynchida and Gnosonesimida). For sim- The phylogenetic classifi cation is based on Laumer and plicity, the old Lecithoepitheliata name is used here. Giribet ( 2014 ), in which the former Lecithoepitheliata Images of representative species are shown (Adapted appears as a paraphyletic group, now divided into two from Martín-Durán and Egger (2012 ) ) 3 Platyhelminthes 23

Platyhelminths lack a coelom; circulatory, COMPARATIVE PLATYHELMINTH skeletal, and respiratory systems; and a defi ni- DEVELOPMENT tive anus. They have a central nervous system (CNS), a blind gut, and fl ame-bulb protone- The embryonic development of fl atworms has long phridia. The space between the organs is fi lled attracted the attention of embryologists and phylo- with mesenchyme (or parenchyma). Their epi- geneticists. The structure of the oocyte was used as dermis is a monolayer of multiciliated cells one of the main features to classify Platyhelminthes: that provide locomotion. A characteristic trait entolecithal species (with eggs that contain all the of this phylum is the presence of a large num- yolk needed for development) were classifi ed as ber and variety of adhesive secretions from Archoophora, since this condition is considered specialized cells and organs. Platyhelminths basal, whereas ectolecithal species (with eggs that have an extreme morphological plasticity, and must take up yolk from outside) were classifi ed as some are capable of regenerating a whole Neoophora, as they required the invention of spe- organism from a small piece or can change cialized yolk cell- producing organs called vitellaria their dimensions continuously according to the (a detailed and recent revision can be found in availability of food. This property is related to Martín-Durán and Egger 2012 ). the existence of a unique population of adult Since the advent of modern molecular phylo- totipotent stem cells, termed ‘neoblasts’, which genetic analyses, the inclusion of Platyhelminthes provide the cellular basis for this morphologi- in the Spiralia has been well supported and cal plasticity. widely accepted (Dunn et al. 2008 ). The spiral Platyhelminths are with few exceptions her- cleavage pattern of Platyhelminthes is character- maphroditic (with cross-fertilization), with ized by an oblique orientation of the spindles rather complex reproductive systems. However, with respect to the embryonic animal-vegetal some free-living species have the ability to axis. Due to the stereotypic nature of this cleav- reproduce asexually by fi ssion and subsequent age pattern, it is possible to identify and name regeneration; moreover, for some parasitic fl at- each cell according to an established nomencla- worms, asexual multiplication is an obligatory ture. Depending on the species, spiral cleavage part of their life cycle. In contrast to other ani- may be either equal or unequal. In the latter case, mals, most platyhelminths have eggs that are one of the blastomeres at the two-cell stage (blas- devoid of yolk (ectolecithal eggs), which is sup- tomere CD) and four-cell stage (blastomere D) is plied by specialized yolk cells. This trait is one typically larger than the others. A feature of the of the main causes for the greatly modifi ed spiral development is the early specifi cation embryonic development observed in many of blastomeres (determinative development). fl atworms. Comparative embryology shows that polyclads This modifi ed embryonic development and follow a relatively stereotypical spiral cleavage their high regenerative capacity (see Chapter 4 ) pattern, while in some neoophorans, spiral cleav- makes them of widespread biological interest. age is replaced by an irregular and disperse cleav- This chapter reviews the embryonic develop- age, referred to as Blastomerenanarchie (Thomas ment of the different subtaxa of Platyhelminthes 1986 ; Baguñà and Boyer 1990 ). Currently, it is from the perspective of evolutionary develop- assumed that the acquisition of ectolecithal eggs mental biology. Chapter 4 uses Platyhelminthes involved the modifi cation of the cleavage pattern. as a model phylum to study the regenerative The observation that quartet spiral cleavage pat- potential across the animal kingdom, with a terns can still be recognized in some neoopho- particular focus on the CNS and the photore- rans (lecithoepitheliates and proseriates), despite ceptors, highlighting the pivotal role played by the presence of extra-embryonic yolk cells within the neoblasts. The role of evolutionarily con- the egg, agrees with the view that spiral cleavage served signalling pathways (Wnt, BMP, Hippo) most likely constitutes the plesiomorphic cleav- in controlling adult plasticity, maintenance of age pattern in Platyhelminthes (Thomas 1986 ; axial polarity, and growth is also discussed in Baguñà and Boyer 1990 ). Interestingly, the strik- Chapter 4 . ing diversity of platyhelminth embryology 24 T. Adell et al.

Table 3.1 Comparison of the main embryonic traits of different subtaxa of Platyhelminthes Egg type Cleavage pattern Larva/direct development Catenulida Entolecithal Spiral (early) Luther’s larva and direct development Polycladida Entolecithal Spiral Müller’s, Goette’s, and Kato’s larva and direct development Macrostomorpha Entolecithal Spiral (early) Direct development Lecithoepitheliata Ectolecithal Spiral Direct development Proseriata Ectolecithal Spiral (early) Direct development Bothrioplanida Ectolecithal Disperse Direct development Rhabdocoela Ectolecithal Irregular Direct development Fecampiida Ectolecithal Irregular Direct development Prolecithophora Ectolecithal Disperse Direct development Tricladida Ectolecithal Disperse Direct development (the embryo has been considered a ‘cryptic larva’) Neodermata Ectolecithal Disperse Larva and direct development

contrasts with the similarity of adult body plans larval stage within Platyhelminthes will be dis- observed, at least among the free-living groups cussed later. (Fig. 3.1 ). Although most of the available data corresponds Polycladida to classical descriptive embryological approaches, The Polycladida consists of large, almost molecular techniques are beginning to be imple- exclusively marine animals. It is divided into two mented in representative species from several taxa suborders: the Cotylea, with a prominent sucker of Platyhelminthes, such as the polyclads and the posterior to the female genital opening, and the triclads (Rawlinson 2010 ; Lapraz et al. 2013 ). The Acotylea, which lack a sucker. Polyclad fl atworms following sections discuss the main fi ndings on provide an interesting system to explore the evolu- embryonic development in each taxon of fl at- tion of development within Platyhelminthes and worms, focusing on early development. Table 3.1 Spiralia, because, unlike most other fl atworms, summarizes the most important known embryonic they undergo spiral cleavage similar to that seen in traits of each group. Emphasis is placed on embry- some other spiralian taxa and some lineages also onic features with evolutionary signifi cance and display indirect development through the forma- topics that deserve further attention. tion of Müller’s and Goette’s larvae. To date, the most comprehensive description of their early embryonic development comes from the species Archoophorans Hoploplana inquilina (Surface 1907 ; Boyer et al. 1998 ). Recently, a new species, Maritigrella croz- The ‘Archoophora’ (Catenulida, Macrosto- ieri , has been established as a suitable species that morpha, and Polycladida) is a paraphyletic group can be maintained in the laboratory (Rawlinson including all fl atworms with endolecithal eggs. 2010 ; Lapraz et al. 2013 ). Archoophorans exhibit quartet spiral cleavage, at Polyclad cleavage is spiral, usually with no least during the early zygotic divisions. size difference between the cells of the four-cell stage, although the D cell can be slightly larger in Catenulida some species. However, the cleavage pattern dif- Knowledge of embryonic development in catenu- fers from that of canonical spiralians in that the lids is scarce. The only points worthy of note here micromeres of the fourth quartet (4a–4d) are are that their early development looks similar to atypically large and the macromeres (4A–4D) are polyclads and, although most of them hatch as a smaller. During gastrulation, the animal micro- juvenile, a so-called Luther’s larva has been meres 4a–4c and 4A–4D become internalized described for Rhynchoscolex simplex (Reisinger completely through epiboly over the vegetal 1924 ). The signifi cance of the existence of this macromeres (Fig. 3.2 ). Micromere 4d produces 3 Platyhelminthes 25

Fig. 3.2 Embryonic development of Polycladida, which will be located on the future ventral side of the Macrostomorpha, Rhabdocoela, and Tricladida. Polyclads embryo. The epidermis differentiates from this embryonic exhibit a conserved quartet spiral mode of development, blastema, which will ingest external yolk cells. In triclads, although the macromeres ( 4A–D) are smaller than the no signals of spiral cleavage are observed, but blastomeres micromeres ( 4a–d ). Gastrulation occurs through epiboly cleave in a disperse manner and remain isolated within the of the animal micromeres over the vegetal macromeres. yolk mass. Afterwards, some blastomeres differentiate Another peculiarity of polyclad development is that mac- into the primary epidermis and embryonic pharynx, which romeres 4A – D and micromeres 4a – c degenerate and the will be used to ingest the yolk cells. After yolk ingestion, whole endoderm and a large part of the mesoderm origi- the remaining blastomeres proliferate and differentiate nate from the 4d micromere. Macrostomids show the typi- into the defi nitive organs. In all schemes, an idealized cal quartet spiral cleavage pattern up to the eight-cell cross section of the animal-vegetal axis is presented (ani- stage. Afterwards, the vegetal macromeres 2A – D fl atten mal to the top , vegetal to the bottom ). Yolk granules are in and cover the embryo. This transient membrane is later light blue , hull cells in orange , and embryonic cells in replaced by the defi nitive epidermis. The rest of the blas- grey. bl blastomere, eb embryonic blastema, ec ectoderm, tomeres remain in the inner region and form the embry- ep epidermis, eph embryonic pharynx, epp embryonic onic blastema, in which organogenesis takes pace. In pharynx primordium, es egg shell, eym embryonic yolk rhabdocoeles, the fi rst cell division gives rise to an animal mantle, ma macromere, mec mesoectoderm, men mesoen- micromere and a vegetal macromere. Proliferation of doderm, mi micromere, yc yolk cell, ys yolk syncytium these two initial cells forms an embryonic blastema, (Adapted from Martín-Durán and Egger (2012 ) ) 26 T. Adell et al.

Fig. 3.3 Larval types and juveniles of Polycladida. juvenile of Pseudostylochus obscurus (Adapted from Müller’s larva of Prosthiostomum siphunculus , Kato’s Martín-Durán and Egger (2012 ) ) larva of Planocera reticulata, and directly developing both endoderm and mesoderm. Micromeres 4a–c et al. 2011 , 2012 ). Some species directly develop are the main yolk-containing cells that are incor- into small juvenile polyclads (Fig. 3.3 ). porated into the gut. The macromeres degenerate (Surface 1907 ; Thomas 1986 ; Boyer et al. 1996 ). Macrostomorpha This differs from a suggested spiral archetype The Macrostomorpha comprises small fl at- (e.g., mollusks and annelids; see Chapters 7 and worms mainly from the taxon Macrostomida. 9 ), where 4d makes only mesoderm and 4a–c and To date, studies on their embryonic develop- the macromeres form the endoderm. The apical ment are restricted to the genus Macrostomum. micromeres give rise to the brain and the epider- Macrostomum lignano has recently been intro- mis. An inner cell layer is formed by the progeny duced as a new model organism for studying their of 4d (called mesentoblast) and the animal micro- development and evolution, since it can be easily mere 2b (mesectoderm) (van den Biggelaar et al. reared in the laboratory, produces a multitude of 1997; Boyer et al. 1998 ). The musculature, gland eggs year round, and has a very short generation cell, protonephridial system, and pharynx muscu- time (2–3 weeks at 20 °C). Its embryogenesis lature are derived from this inner layer. The has been described in detail (Morris et al. 2004 ). dorsoventral (DV) and anterior- posterior (AP) Early cleavage, up to the eight-cell stage, occurs body axes are established by the mesentoblasts in a typical spiral cleavage pattern. In later stages, and their progeny, which form a ventral plate of development starts to deviate from this pattern, cells. A series of blastomere ablation studies mainly because of the formation of an external showed that polyclad development is determina- yolk mantle from the four vegetal yolky mac- tive (Boyer et al. 1998 ). romeres, the so-called hull cells, which cover Later development shows much variation the embryo (Willems et al. 2009 ). This process from almost direct to indirect with free- swimming was called ‘inverse epiboly’ by Thomas ( 1986 ). larvae, which are classifi ed according to the num- Within the yolk mantle, the remaining blasto- ber of lobes and eyes that are present. Many poly- meres form a proliferating mass, the embryonic clads have a spherical Müller’s larva with eight primordium, from which the defi nitive structures, lobes and three eyes. Some species have Goette’s such as the epidermis that will replace the hull larva or Kato’s larva which only differ from each cells, and the gut primordium will arise (Fig. 3.2 ). other in the number of lobes and eyes (Gammoudi After 4–7 days (at 20 °C), the juvenile hatches. 3 Platyhelminthes 27

Hull cells are not only found in macrostomids other spiralians, and in contrast to the situation but in all fl atworm taxa, with the exception of observed in polyclads, the macromeres are large polyclads. The presence of hull cells is the most and give rise to the endoderm (Reisinger 1972 ). obvious deviation from the typical spiral develop- In Lecithoepitheliata, gastrulation involves epib- mental pattern in Platyhelminthes. By defi nition, oly of the micromeres over the vegetal macro- hull cells are large, yolk-rich embryonic cells that meres at the 25–30-cell stage. The micromeres surround the blastomeres after the early divisions. then fl atten and differentiate into the hull mem- Although hull cells look similar and occupy the brane, which engulfs a portion of the yolk. In same position within fl atworm embryos, they do Proseriata, no pattern is discernable in late cleav- not share any ontogenetic origin, indicating that age stages. In both groups, the inner mass of blas- they are not homologous structures. Hull cells tomeres differentiates into an embryonic blastema may originate from the vegetal macromeres (as in that occupies the future ventral side of the Macrostomum), from animal micromeres (as in embryo, where the defi nitive organs and the epi- proseriates or lecithoepitheliates), or from vitel- dermis that will replace the hull membrane will laria yolk cells (as in triclads, prolecithophorans, differentiate (Reisinger 1972 ). and rhabdocoeles). Bothrioplanida, Rhabdocoela, Fecampiida, and Prolecithophora Neoophorans The embryonic development of Bothrioplanida, Rhabdocoela, Fecampiida, and Prolecithophora The common feature of Neoophora is the laying species has not been studied in detail. However, of ectolecithal eggs. Neoophora have oogonia since they share essential features, they are con- that are divided into the germarium, which pro- sidered here together. These four groups show duces the oocytes, and the vitellarium, which no signs of a spiral quartet. In Rhabdocoela and produces the yolk cells. Thus, the egg contains Prolecithophora, the fi rst cell division gives rise both oocytes and extra-embryonic yolk cells, and to an animal micromere and a vegetal macro- all Neoophora embryos have developed mecha- mere, which have been proposed to be homolo- nisms to engulf the external yolk cells, usually by gous to blastomeres AB and CD of canonical forming a temporary epidermis known as hull spiralians (Hartenstein and Ehlers 2000 ; Younossi- membrane. Ectolecithality determines the highly Hartenstein et al. 2000 ). Late cleavage does not divergent mode of development within this group follow a regular pattern and results in an embryonic and with respect to the rest of the spiralians. blastema from which a transitory epidermis differ- Whereas some of them retain a kind of quartet entiates to engulf the external yolk cells (Fig. 3.2 ). spiral cleavage (Lecithoepitheliata and Afterwards, the embryonic blastema differentiates Proseriata), others (Bothrioplanida, Rhabdocoela, to form the defi nitive epidermis and organ primor- Fecampiida, Prolecithophora, and Tricladida) dia, as observed in all other neoophoran fl atworms. have a disperse mode of cleavage that, in the case All directly hatch as juvenile worms. of the Tricladida, has been classically called Blastomerenanarchie (Seilern-Aspang 1958 ). Tricladida Juveniles directly develop from an embryonic Tricladida includes marine, freshwater, and ter- blastema. Obviously, such divergent cleavage restrial species. Due to the amazing regenerative patterns result in divergent gastrulation and lin- abilities of the adults in some species, Tricladida eage fate processes. is by far the best-studied platyhelminth taxon, not only at the morphological but also at the molecu- Lecithoepitheliata and Proseriata lar level. The majority of species analysed with Lecithoepitheliata and Proseriata retain a kind of respect to their potential to regenerate are fresh- stereotypical quartet spiral cleavage pattern, just water animals (Brøndsted 1969), but embryologi- up to the eight-cell stage in Proseriata. As in cal studies have been reported from representatives 28 T. Adell et al.

Stage 1 Stage 2 Stage 3 Stage 4 yc bl gb pe gb zy yc bl pe yc ys ys yc bl

ti

epp pe ti ep es pe ep yc ep pnc ep stage 5-6 stage 7 stage 8

yc mc ed yc ed pa e yc pa ep e pgb pa pgb

pe dpp cns cns mo agb ms dp np pnc agb ms dp gc

Stage 1 Stage 3 Stage 5 Stage 6 Stage 7 Stage 8

dp e dp e

Fig. 3.4 Overview of embryogenesis in the planarian phology observed in stage 8 embryos and hatchlings. agb Schmidtea polychroa . Stage 1, blastomeres divide within anterior gut branch, bl blastomere, bp brain primordium, the mass of yolk cells. Stage 2, each zygote gives rise to a cns central nervous system, dp defi nitive pharynx, dpp yolk-feeding embryo with its transient embryonic struc- defi nitive pharynx primordium, e eye, ed epidermis, ep tures (i.e., primary epidermis, embryonic pharynx). Stage embryonic pharynx, epp embryonic pharynx primordium, 3, due to the ingestion of yolk cells, the embryo grows in es egg shell, gb germband, gc gastrodermal cell, mc mus- size, and the initial yolk-derived syncytium becomes cle cell, mo mouth, ms muscle, np neural precursor, pa restricted to the periphery of the embryo, forming the so- parenchyma, pe primary epidermis, pgb posterior gut called germband, composed mainly of proliferating blas- branch, pnc pioneer nerve cord, ti temporary intestine, vnc tomeres. Stage 4, blastomeres in the germband start ventral nerve cord, yc yolk cells, ycI type I yolk cells, ycII differentiating, and the defi nitive nerve cords become vis- type II yolk cells, ys yolk syncytium, zy zygote. Images of ible. Stages 5–6, specifi cation of defi nitive axial identities planarian embryos corresponding to different stages are and differentiation of the adult cell types and organs shown. Anterior is to the left . Scale bars: 500 μm except (defi nitive pharynx and brain primordium) start. Stage 7, for stage 8 where it is 1 mm (Adapted from Martín-Durán the defi nitive organs arise and develop to acquire the mor- et al. (2012a ) ) of all three habitat types. Whereas the freshwater the syncytium, where cleavage takes place. As planarian Schmidtea mediterranea has emerged in the Prolecithophora and Bothrioplanida, as the model for the study of regeneration mecha- Blastomerenanarchie -type cleavage is observed nisms and stem-cell biology, its sister species with no canonical pattern and none of the blasto- Schmidtea polychroa has become the model of meres remaining next to the other. After several choice for embryological studies (reviewed in divisions, transient structures organize, such as Martín-Durán et al. 2012a ). the embryonic epidermis (the hull membrane) Triclads lay cocoons from which one to ten and the embryonic pharynx, through which the juveniles may hatch (Tekaya et al. 1999). The yolk syncytium is engulfed (Figs. 3.2 and 3.4 ). cocoons contain several zygotes surrounded by The rest of the blastomeres remain in the periph- a large number of extra-embryonic yolk cells ery of the embryo in an undifferentiated state. It (Cardona et al. 2005), which fuse and form a is known that they express stem-cell-associated syncytium after deposition (Figs. 3.2 and 3.4 ). gene markers such as vasa and tudor (Solana and The early blastomeres become embedded inside Romero 2009 ). Once the embryo has internalized 3 Platyhelminthes 29

A

B CD EF G

Fig. 3.5 Gene expression during planarian embryonic expression of the respective genes in adult animals. (B – G ) development. (A ) Overview of major developmental Expression pattern of specifi c genes analysed by whole events ( coloured bars ) and associated gene expression mount in situ hybridization (stage 2 in B , stage 5 in C , ( coloured curves) with respect to stages and days of devel- stage 6 in D – F , and stage 7 in G ). See main text for opment at 20 °C (Modifi ed from Martín-Durán et al. references ( 2012a )). Broken lines to the right indicate continued enough yolk, the blastomeres migrate and dif- any specifi c AP position (Cardona et al. 2005 ; ferentiate into the defi nitive structures that will Martín-Durán and Romero 2011 ). Molecular replace the embryonic ones, that is, the defi ni- studies show that the stages at which the pri- tive epidermis, pharynx, nervous, digestive, and mordia of the defi nitive organs appear, the so- muscle systems (Fig. 3.4 ). The process of organ- called fi ve to six stages, are the ones in which ogenesis was formerly described as involving the the molecular markers of axial polarity (Wnt formation of three main ventral primordia (an and BMP pathway elements) appear (Marín- anterior brain primordium, a central pharynx pri- Durán et al. 2010 ), supporting consideration of mordium, and a posterior primordium), as in other this stage as the point at which the body plan neoophoran fl atworms. However, recent studies of the adult is established (for a review of gene on S. polychroa demonstrate that the appearance expression during Schmidtea polychroa embryo- of the defi nitive structures does not occur in genesis, see Fig. 3.5 ). Interestingly, it is at these 30 T. Adell et al. stages when triclads start to display an ability to this process seems to be controlled by the genes regenerate, suggesting that neoblasts appear at sine oculis and eye absent but appears to be inde- the same time as other defi nitive cell types. The pendent of Pax6 , a proposed master regulator of most obvious event of adult organogenesis is animal eye development (see Chapter 4 ). the formation of the defi nitive pharynx primor- Although the formation of the hull mem- dium ventrally to the degenerating embryonic brane is a common trait in Platyhelminthes pharynx (Martín-Durán and Romero 2011). The embryos, the presence of a transient embryonic pharynx primordium, which strongly expresses pharynx, with the associated neural structures, the gut marker foxA (Martín-Durán et al. 2010 ), is only observed in triclads. An interesting fi rst occupies a posterior position, and it moves question is thus the extent to which this com- to more central regions as the embryo elon- plex transient yolk-feeding embryo can be con- gates anterior-posteriorly. Simultaneously to sidered an encapsulated larva and whether its the formation of the pharynx, gastrodermal cells formation can be considered a process of gas- ( GATA456a - positive cells) appear scattered trulation. Although it had traditionally been through the margins of the embryo and eventu- accepted that triclads, and other Platyhelminthes ally enclose the ingested yolk cells and defi ne with disperse cleavage, lack true gastrulation the defi nitive gastrodermis, which is already movements, the anatomy of their embryos divided into the three main gut branches. The indicates that well-defi ned tissues belonging appearance of the musculature seems to follow to the ectoderm (primary epidermis), meso- a similar dynamic, with muscular precursors derm (pharyngeal muscle), and endoderm ( myosin heavy chain- positive , mhc, cells and cells (temporary intestine) are present at this time. immunolabeled by the specifi c antibody TMUS- Moreover, recent molecular studies have dem- 13) being fi rst specifi ed diffusely throughout onstrated the expression of evolutionarily con- the embryo and then progressively forming an served gastrulation-related genes during the orthogonal grid as the embryo elongates and formation of the embryonic pharynx, including fl attens (Cardona et al. 2005 ; Martín-Durán snail, twist, foxA, and β- catenin (Martín-Durán et al. 2010 ). Neurogenesis starts with the for- et al. 2010 ). These fi ndings suggest that ancient mation of a defi nitive neural primordium, made mechanisms of early cell fate specifi cation are of two bilaterally symmetrical condensations of still present in triclad embryos and, therefore, neural progenitors. As embryogenesis contin- that planarian development can be separated ues, this primordium develops into the anterior into two morphogenetic stages: a fi rst gastrula- bilobed brain and the two main ventral nerve tion process and a subsequent ‘metamorpho- cords (Cardona et al. 2005 ). In contrast to stud- sis’. During gastrulation, the three germ layers ies on the embryonic development of the diges- segregate and establish the primary organiza- tive and muscular systems, and despite the great tion of the feeding embryo or ‘cryptic’ larva amount of data regarding adult nervous system (Cardona et al. 2006). Metamorphosis then regeneration (see Chapter 4 ), gene expression establishes the defi nitive adult body plan studies on planarian embryonic neurogenesis through the involvement of totipotent blasto- are still lacking. As a fi rst step, the embryonic meres. During this second stage, the morpho- development of the photoreceptors was charac- genetic and patterning mechanisms are similar terized at the genetic level (Martín-Durán et al. to those used during regeneration and 2012b ). Simultaneously to the early formation of homeostasis in the adult, e.g., in the use of the the brain, an anterior pair of eye precursor cells Wnt and BMP signalling pathways to establish (opsin- and tryptophan hydroxylase (tph)-positive the AP and the DV axis (Martín-Durán et al. cells, detected by the antibody against arrestin) 2010 ) (see Chapter 4 ). The cryptic larva may appears beneath the dorsal epidermis, which be considered a vestigial larva, or, since sev- develops into the eyes of the juvenile (Martín- eral embryos share the same maternal resources Durán et al. 2012b ). As in adult eye regeneration, in each cocoon, it could be the evolutionary 3 Platyhelminthes 31

Fig. 3.6 Life cycle of Schistosoma . Eggs are expelled residence in the veins (8 , 9 ). Adult worms in humans with faeces ( 1 ). Under optimal conditions, miracidia reside in the mesenteric venules in various locations, ( 2 ) hatch from the eggs and swim and penetrate the snail which seem to be specifi c for each species ( 10 ). The intermediate host ( 3). The stages in the snail include two females deposit eggs in the small venules of the portal and generations of sporocysts ( 4) and the production of perivesical systems. The eggs are moved progressively cercariae (5 ). Upon release from the snail, the infectious towards the lumen of the intestine (S. mansoni and S. cercariae swim, penetrate the skin of the human host (6), japonicum) and are eliminated with faeces (1 ) (Adapted and become schistosomulae ( 7 ). The schistosomulae from Wikimedia Commons) migrate through several tissues and stages to their result of their competition for the limited primary ciliated epidermis, which is replaced maternal yolk (Cardona et al. 2006 ). Further during development. Other specializations of discussion about the gastrulation and the direct the Neodermata, a result of its adaptation to the versus indirect mode of development in endoparasitic lifestyle, are the reduction or Platyhelminthes is presented below. absence of a gut, a simplifi ed nervous system, and the existence of a complex life cycle that Neodermata involves several larval stages inhabiting differ- Neodermata comprises the parasitic fl atworms, ent animal hosts (Fig. 3.6 ). Their life cycle has classifi ed in three taxa: Monogenea, Cestoda, and evolved to give rise to a large number of Trematoda. The body wall of Neodermata, an descendants. Embryological studies of these unciliated syncytium called the neodermis, repre- taxa are scarce, since most of the effort is focused sents the defi ning morphological criterion of the on the understanding of the life cycle of the spe- clade. Ontogenetically, the neodermis is formed cies which represent a public health problem, secondarily underneath an earlier developing either directly producing human diseases, such 32 T. Adell et al. as schistosomiasis (Fig. 3.6), or affecting human the embryophore, may retain its ciliation. Upon resources, e.g., as pests in fi sh farms. hatching in the gut of the primary host, the larva sheds its protective envelopes and penetrates Cestoda through the intestinal wall into the body cavity of Cestoda are commonly known as tapeworms, and the host. Once inside the body cavity, this pri- about 4,000 species have been described. Almost mary larva metamorphoses into a secondary all vertebrate species may function as potential larva, called the cysticercoid. hosts. Tapeworms usually require at least two A well-known example is the beef tapeworm hosts, and humans represent the primary host of Taenia saginata , the adult form of which lives in several species, with infestation occurring the human small intestine and can attain a length through uptake of uncooked meat such as pork of 20 m, folded back and forth in the host intes- ( Taenia solium), beef (Taenia saginata), or fi sh tine. Fertilized eggs shed from the human host ( Diphyllobothrium spp.). Adult tapeworms are are ingested by cows, the intermediate hosts, parasitic in the digestive tract of vertebrates, and where six-hooked larvae, the oncospheres, hatch often one of the intermediate hosts is an inverte- and migrate to skeletal muscles where they encyst brate. Tapeworms usually have long fl at bodies to become ‘bladder worms’ or cysticercoids. composed of many reproductive units or proglot- Each of these juveniles remains quiescent until tids. This represents a specifi c trait of Cestoda, the uncooked muscle is eaten by humans, where since each proglottid contains the male and it attaches to the intestine and matures. More female reproductive structures and can reproduce dangerous to humans is the species Taenia independently; therefore, it could be considered solium, or pork tapeworm, because humans can that a tapeworm is actually a colony of proglot- act also as secondary hosts. Cysticercoids can tids. Tapeworms completely lack a digestive sys- move from the muscle to the brain and cause cys- tem, and, as in Trematoda, there are no external ticercosis, which can be lethal. motile cilia. In contrast to monogeneans and Embryogenesis of Taenia and Hymenolepis trematodes, however, their entire surface is cov- species has been described at the morphologi- ered with small projections that resemble the cal level (Hartenstein and Jones 2003 ). The gen- microvilli of the vertebrate small intestine. As in eral mode of fl atworm development can still be the intestine, these microvilli serve to expand the discerned in both of them. The oocytes are sur- absorptive surface. This represents an adaptation rounded by a hard outer capsule formed within the for a tapeworm, since it must absorb all of its ovary. They divide totally and unequally. Similar nutrients across the tegument. Tapeworms have to what has been described for the embryos of well-developed muscles, and their excretory and many neoophoran fl atworms, no spiral cleavage nervous systems are similar to those of other fl at- pattern is apparent. During the early cleavage worms. They have no special sense organs but do divisions, large macromeres separate from the have modifi ed cilia that function as sensory end- micromeres and give rise to two syncytial sheaths ings in the tegument. One of their most special- surrounding the embryo. The inner envelope is ized structures is the scolex, the organ of considered as the primary epidermis or embryo- attachment, which usually has suckers and spiny phore, which is only ciliated in coracidium lar- tentacles. vae of aquatic tapeworms. Like in free-living Tapeworms have a complex life cycle that neoophoran platyhelminths, non-morphogenetic begins with an infectious larva, called a coracid- movements that resemble a classical gastrulation ium in aquatic tapeworms and oncosphere in ter- are observed. During organogenesis the hooklets, restrial tapeworms. This simplifi ed passive larva specifi c structures that cestode larvae use to pen- enters the primary host by being ingested. The etrate the epidermis of the host, are differentiated, larva is extremely small, containing only a few together with the muscle and the nervous system. embryonic cells (50–100) surrounded by several Oncosphere larvae have a complex system of protecting envelopes. The inner envelope, called muscle fi bres connected to the hooklets, and this 3 Platyhelminthes 33 enables them to generate coordinated movement the major infectious diseases in the world and is to enter the body cavity of the host. A pair of neu- widely prevalent in developing countries. As in rons located in the anterior tip of the larva sends all trematodes, the life cycle of Schistosoma is axons posteriorly and branch profusely along the very complex (Fig. 3.6 ). It has separate sexes that deep muscle fi bres to innervate the musculature live for several years in blood vessels. Inside the (Hartenstein and Jones 2003 ). host vasculature, each female can release about Despite its extreme reduction in overall cell 300 eggs per day. When eggs are excreted into number and different cell types, the oncosphere fresh water by the primary host, a ciliated larva larvae of cestodes exhibit similarities to the lar- called miracidium hatches in about 5 days. In vae or juveniles of other fl atworms, notably an snails of the genus Biomphalaria , miracidia anterior ‘brain’ (consisting of only two neurons undergo cycles of asexual reproduction, leading in cestode larvae), as well as other resemblances to the emergence of a second larval type, the cer- during ontogenesis. A common feature of many caria. Cercariae swim actively and penetrate the cestode and trematode species is the simultane- defi nitive host, again through the skin to enter ous apoptotic death of several micromeres to blood vessels, where they develop into adults form the embryonic envelopes and the multipli- (Jurberg et al. 2009 ). Through this complex cation and differentiation of other blastomeres. cycle, a single zygote of parasitic fl atworms can This genetically programmed cell death leads to give rise to an enormous number of progeny. The a simplifi cation of the infective larval stage and disease is caused not by the transmitted eggs but likely represents an ontogenetic adaptation to the rather by chronic infl ammation that occurs when parasitic life strategy (Młocicki et al. 2010 ). eggs released by female worms get trapped in Mesocestoides corti represents a different small blood vessels of the liver or other organs model for studying cestode biology and develop- (Fig. 3.6 ). ment. It is particularly interesting because of its Schistosomes are neoophorans. Therefore, intermediate larval stage (tetrathyridium), com- the yolk cells, produced by the vitellarium, are posed of a scolex and an unsegmented body that found outside the oocyte. In general, their is able to proliferate asexually by longitudinal fi s- embryology resembles that of free-living fl at- sion in the intermediate hosts. This enables to worms. Schistosome eggs are not retained in the study asexual reproduction, proglottid formation, female uterus and remain uncleaved when laid. and strobilar development, leading to the forma- Cleavage starts in the blood of the primary tion of serially arranged genital organs, one in host and produces two blastomeres that differ each segment (Koziol et al. 2010 ). slightly in size. During the fi rst cleavages, the vitelline cells fuse into a single yolk syncytium. Trematoda Subsequent cleavages are asynchronous and Trematodes are all parasitic fl ukes, and as adults form a discoidal embryonic primordium with an they are almost all found as endoparasites in ver- animal-vegetal axis. This grows to form the ste- tebrates. However, they share several characteris- reoblastula, which comprises a central prolifer- tics with free-living Platyhelminthes, such as a ating embryonic primordium surrounded by an well-developed alimentary canal and similar outer envelope, produced by the migration of reproductive, excretory, muscle, and nervous three to four macromeres. At this point, cellular systems. differentiation begins, giving rise to an inner Trematodes have a complex life cycle. The envelope and the appearance of early organ pri- intermediate host is a mollusk, and the defi nitive mordia. Afterwards, organogenesis takes place, host is a vertebrate. In some species, a second and with the differentiation of the neural mass, the sometimes even a third intermediate host is epidermis, the musculature, and the miracidial involved. The genus Schistosoma has been the glands. Hatching releases a fully formed larva subject of several studies, since it is the causative that displays muscle contraction, beating cilia, agent of schistosomiasis, which ranks as one of and fl ame cells. 34 T. Adell et al.

Although almost nothing is known about the such as in some cestodes is not required in molecular control of trematode embryogenesis, a Aspidogastrea since the different organization of recent report demonstrated the importance of the life cycle makes a locomotor function TGFb signalling during egg development. It was unnecessary. Almost simultaneously with blasto- shown that the protein encoded by SmInAct, the mere multiplication, the progressive degenera- Schistosoma homolog of activin, is produced only tion of some micromeres begins, which is a when females are paired with males in an immu- common trait in neodermatans. nologically competent setting (Freitas et al. 2007). In this study, RNA interference showed that SmInAct plays a crucial role in egg development. EVOLUTIONARY IMPLICATIONS Recently, the early development of an Aspidogastrea species (A. limacoides ) has been In the following sections, the most reliable studied at the ultrastructural level (Świderski embryonic characters are summarized and dis- et al. 2011). The subclass Aspidogastrea is a cussed, putting forward evolutionary hypotheses small group of parasitic fl atworms generally con- and future directions that will be useful for devel- sidered to belong within the Trematoda, although opmental and evolutionary studies. several differential traits call this into question. Aspidogastreans occur worldwide in marine and freshwater environments, where they use a mol- Spiral Cleavage in Ectolecithal lusk as an obligate host. Although in some cases Embryos a vertebrate is a facultative or obligate fi nal host, their entire life cycle may occur within the mol- Current phylogeny supports the hypothesis that lusk. In contrast to what is found in trematodes quartet spiral cleavage is the ancestral develop- and cestodes, there are no asexual generations in mental mode in Platyhelminthes. The basal posi- known life cycles of aspidogastreans. Also in tion of Polycladida in the most recent phylogenetic contrast to other parasites, it shows a rather direct analysis further supports this view (Laumer and development, since it produces a ‘larva’ called a Giribet 2014 ). The developmental deviations cotylocidium that exhibits the fundamental orga- found in polyclads, such as the degeneration of nization of the adult. The eggs of A. limacoides fourth quartet macromeres and of micromeres are ectolecithal and contain a fully developed 4a–c, are probably apomorphies of this group, cotylocidium when laid, revealing some degree since they are not present in those taxa of neooph- of ovoviviparity. The fi rst cleavage is equal, but oran Platyhelminthes that retain a quartet spiral subsequent cell divisions are unequal and asyn- cleavage. Further study of the embryonic devel- chronous. These cleavages end in the formation opment in the Catenulida will be essential to of the early embryo, which is composed of sev- understand the evolutionary history of cleavage eral blastomeres of different sizes. The largest patterns in the Platyhelminthes. blastomeres, the three macromeres, undergo a The presence of external yolk cells in neooph- very early fusion of their cytoplasm, forming a orans does not necessarily imply the loss of spiral syncytial layer of the embryonic envelope directly cleavage, as exemplifi ed in lecithoepitheliates and beneath the egg shell. The presence of this early proseriates. The complete absence of spiral cleav- and unique envelope is the most important age in the rest of the Neoophora (Neodermata, difference between Aspidogastrea and other Bothrioplanida, Rhabdocoela, Fecampida, Neodermata. The lack of an inner embryonic Prolecithophora, and Tricladida) might thus be envelope in aspidogastreans may be explained by explained by a single evolutionary event at the base the close contact between the vitelline syncytium of this group. However, the signifi cant differences and the differentiating blastomeres, which pro- between the cleavage in rhabdocoeles and the vides a direct passage for nutrients. Moreover, remaining taxa suggest that they could have evolved the existence of an inner ciliated epithelium independent adaptations to their ectolecithal 3 Platyhelminthes 35

condition. Whether the disperse cleavage in the formation of the planarian feeding embryo meets remaining taxa shares a common origin remains these criteria. Despite the ‘anarchic’ situation to be clarifi ed. A recent study demonstrated the that might be apparent at fi rst sight, defi ned absence of centrosomes in the triclad Schmidtea groups of blastomeres are formed during planar- mediterranea and in the neodermatan Schistosoma ian development. Blastomeres that form the pri- mansoni (Azimzadeh et al. 2012 ), suggesting that mary epidermis, the embryonic pharynx, or this loss occurred concomitantly with the loss of remain undifferentiated in the syncytium each spiral cleavage and the emergence of disperse have a specifi c molecular profi le. Thus, the cleavage in the ancestor of triclads and schisto- expression of β catenin - 1, a master regulatory somes. This hypothesis will need to be further gene for the formation of the endomesoderm tested by studying the presence of centrosomes in across metazoans, is restricted to the cells that other groups of fl atworms. form the embryonic pharynx, and these cells It remains unclear how the changes in the express genes related to specifi cation of the ancestral quartet spiral cleavage affect cell fates mesoderm (twist and myosin heavy chain ) and during early embryogenesis. Polyclads follow a endoderm (foxA ) (Martín-Durán et al. 2010 , determinative mode of cleavage, whereby the 2012a ). This segregation of different groups of loss of blastomeres during early development blastomeres is also directly related to the estab- cannot be compensated by the remaining ones lishment of the primary embryonic organization, (Boyer et al. 1998). Due to experimental diffi cul- which represents the scaffold along which the ties, ablation experiments are still lacking in the defi nitive embryo will develop. neoophorans, and thus it remains unclear whether Related to gastrulation, one of the long- disperse cleavage is determinative or regulative. standing questions concerning the evolution of spiralian development relates to the origin of the mesoderm. In many spiralians, the 4d cell (the Gastrulation in Neoophoran mesentoblast) produces the endomesoderm and Platyhelminthes some of the endoderm. The second and third quartet of micromeres, which generate largely The ancestral mode of gastrulation in ectodermal derivatives, also provide a second Platyhelminthes seems to be the epibolic move- source of mesoderm referred to as ectomesoderm ment of animal micromeres over the vegetal-most (Boyer et al. 1998 ). In polyclads, three peculiari- blastomeres, as observed in polyclads, lecitho- ties that deviate from other spiralians are found: epitheliates, and other phyla with quartet spiral the complete endoderm is formed by 4d, the 4d2 cleavage. However, this mode of gastrulation is cell and not 4d forms the mesentoblast, and the not conserved in neoophorans, since it is infl u- macromeres are smaller than the micromeres and enced by the amount of yolk and the disperse do not contribute to any embryonic tissue forma- cleavage. This raises the question of how they tion. On the other hand, although only prelimi- gastrulate or indeed whether they gastrulate at nary data are available, current observations all. In the groups with disorganized cleavage, indicate that the mesoderm is formed differently such as the triclads, in which the blastomeres are in Macrostomum, since the progenitor of the not attached to each other (Blastomerenanarchie ), mesoderm precursor (4d cell) forms a hull cell whether or not a true gastrulation occurs is still and is then lost. Since the only data about the ori- an open question. An attempt to answer this ques- gin of the endomesoderm in the groups that have tion requires that we fi rst defi ne gastrulation. If lost spiral cleavage is the formation of the tran- we consider that gastrulation involves a series of sient embryonic pharynx of triclads from the coordinated cell and tissue movements that lead early segregation of a subset of blastomeres, fur- to the formation of the distinct cell layers of an ther studies are necessary to determine whether embryo (ectoderm, mesoderm, and endoderm) the molecular signals that specify endomesoder- and to a basic body plan, we must accept that the mal tissues are conserved in Platyhelminthes 36 T. Adell et al. despite the evolution of such divergent develop- As expected, the neoophoran groups that exhibit mental modes. In particular, attention should be disperse cleavage do not show any apparent axial paid to the role of pathways such as canonical correlation. However, conclusions are diffi cult to Wnt signalling in archophoran and neoophoran draw, since the groups with disperse cleavage, species. such as triclads, have yet to be studied in detail. The existence of stereotypic cell-cell interactions between blastomeres and any inductive signal Correlation Between Zygotic, between them is unlikely to occur in triclads. Embryonic, and Adult Polarities However, some studies have shown a primary polarity in the oocyte (Benazzi 1950 ; Anderson Spiralian development relies on maternal deter- and Johann 1958 ), and the presence of maternal minants and cell-cell interactions to specify the determinants, like β catenin - 1, has also been dem- different cell lineages and axial organizers during onstrated (Martín-Durán et al. 2010 ). cleavage. In some taxa, the axial organizer, the D To conclude, no specifi c correlation between quadrant and its descendants, is already defi ned the animal-vegetal axis and the anterior-posterior as early as in the four-cell embryo, due to the axis is present in Platyhelminthes, except for transmission of maternal determinants to one of macrostomids and polyclads. Clearly, the appear- the four cells through asymmetric cell divisions. ance of ectolecithal development led to dramatic These embryos are unequal-cleaving spiralians. changes in the specifi cation of the embryonic However, other spiralian lineages are equal- polarity. However, a combination of morphologi- cleaving, and therefore, early cleavage does not cal and molecular studies in representatives of lead to specifi cation of the D quadrant. each taxon will be required to resolve the mecha- Nevertheless, it is the process of cell-cell com- nism of axial establishment and its evolutionary munication that sets up the embryonic organizer, signifi cance in Platyhelminthes. during which one of the vegetal blastomeres con- tacts the animal micromeres and becomes speci- fi ed as the D quadrant (van den Biggelaar and Direct Versus Indirect Development: Guerrier 1979 ). Regardless of the type of cleav- Was a Larva Part of the Ancestral age, in spiralians the descendants of the D quad- Platyhelminth Life Cycle? rant will occupy a dorsoposterior position in the embryo and will guide the cell lineages of the A, Indirect development has been described in three B, and C quadrants to follow their specifi c posi- groups of free-living platyhelminths: catenulids tions. In this way, the animal-vegetal axis of the with Luther’s larva; polyclads with Müller’s, oocyte represents the future anterior-posterior Goette’s, and Kato’s larva; and fecampiids. axis of the embryo. The extent to which these However, Luther’s larva of catenulids has only events are conserved in Platyhelminthes, how- been observed in a single species, and it is ever, remains unclear. In Macrostomum , this rela- virtually identical to the adult, suggesting that tionship seems to be conserved (Seilern-Aspang catenulids would be better considered as direct 1957 ). In polyclads, although the D quadrant is developers (Martin-Duran and Egger 2012 ). specifi ed as in other equal-cleaving spiralians, a Similarly, the existence of a true larval stage in 90° shift of the anterior-posterior axis in relation fecampiids is under debate. Polyclad larvae are to the animal-vegetal axis has been described the only nonparasitic platyhelminths that show (Younossi-Hartenstein and Hartenstein 2000 ). In unique features not found in the adult worm, the neoophoran groups that retain spiral cleavage such as the lobes and an apical organ, which (lecithoepitheliates and proseriates), the D quad- are resorbed during metamorphosis. In Müller’s rant is also specifi ed, but the animal-vegetal axis and Goette’s larvae, a dorsoventral fl attening corresponds to the defi nitive dorsoventral axis, and an increase in the number of eyes also take where the animal pole becomes the ventral side. place during metamorphosis (Ruppert 1978 ). 3 Platyhelminthes 37

Nevertheless, most parts of the larval body are commonalities in the formation of the adult body retained during postembryonic development, plan in all neoophorans and macrostomorphs similar to, e.g., polychaete annelids or most mol- (Martín- Durán and Egger 2012 ). In these groups, lusks (see Chapters 7 and 9 ). the defi nitive cell types and tissues develop from All other free-living Platyhelminthes undergo embryonic anlagen rather late in embryogenesis, direct development. However, the presence of an after the extra-embryonic yolk cells have been intracapsular larva has been proposed in triclads. incorporated into the embryo (in the Neoophora) This is based on the presence of specifi c organs in or the hull membrane has covered the embryo (in the early yolk-feeding embryo (embryonic phar- the Macrostomorpha). As an example of neooph- ynx and primary epidermis) that are later replaced oran organogenesis, organ development seems to by the defi nitive organs in the hatchling planari- occur quite diffusely in the triclad Schmidtea ans in a process that could be considered a meta- polychroa, with the different cell types forming morphosis. The difference in the morphology of simultaneously throughout the entire germband. the triclad embryo and juvenile is supported by Only the brain and the pharynx primordia appear the existence of a different molecular profi le as a specifi c cluster of cells in the anterior and pos- (Martín-Durán et al. 2010 ). Nevertheless, the terior region, respectively. In the Macrostomorpha, absence of indirect development in the lineages defi nitive organs also develop from an embryonic that branched off between polyclads and triclads primordium (Morris et al. 2004 ). In this group, indicates that the polyclad larvae and the putative however, a close association between myoblasts intracapsular larva of triclads are likely indepen- and neuroblasts during differentiation has been dent adaptations of both lineages. described, perhaps refl ecting the intricate func- All neodermatan fl atworms have one or more tional relationship of muscle and nerve cells in the larval stages, but these are probably adaptations body wall musculature (Reiter et al. 1996 ). to their parasitic life style and not homologous to Polyclads are an exception to the other fl at- the polyclad larvae. Therefore, only polyclads worms since they exhibit a typical determinative and neodermatans show a prototypical larval spiralian development, in which cell types are stage, and the existence of cryptic larvae has been specifi ed early in ontogeny and proper organo- proposed for triclads. The distribution of indirect genesis starts right after gastrulation. This situa- development in the current internal phylogeny of tion is thus more similar to other studied Platyhelminthes (Laumer and Giribet 2014 ) spiralians and can be considered to be the ances- together with the presence of direct development tral state to all Platyhelminthes. Studies are, in the Gastrotricha, a proposed sister taxon to however, scarce and have mostly focused on the fl atworms (Dunn et al. 2008 ; Struck et al. 2014 ), formation of the larval neuromuscular system suggests that direct development may be the (Younossi- Hartenstein and Hartenstein 2000 ; ancestral life style in the Platyhelminthes. Bolaños and Litvaitis 2009 ; Rawlinson 2010 , 2014 ; Semmler and Wanninger 2010 ). In indi- rect developing polyclads, an initial apical heli- Organogenesis in Platyhelminthes coid muscle develops into an orthogonal grid of longitudinal and circular muscles, which corre- Little attention is paid in this chapter to late sponds to the adult body wall musculature. embryogenesis of Platyhelminthes, when organo- Longitudinal muscles seem to derive from the genesis takes place. The main reason is that, vegetal 4d blastomere (endomesoderm), while besides for Tricladida (see above), not much is circular fi bres develop from the 2b micromere known about this process either at the descriptive (ectomesoderm) (Boyer et al. 1996 ). The devel- or the molecular level in the majority of the fl at- opment of the dorsoventral parenchymatic mus- worm lineages. Nevertheless, although early cles occurs after hatching in the planktonic phase development among different Platyhelminthes is and seems to be associated with the metamor- highly variable, there seem to be signifi cant phosis into the juvenile, as this process likely 38 T. Adell et al. controls the dorsoventral fl attening of the larva. • What is the developmental origin of neoblasts Direct developing polyclads seem to lack the ini- (adult stem cells)? tial apical helicoid muscle, but the embryo • How do the different developmental strategies hatches with a muscular pattern similar to the observed in Platyhelminthes relate to the dif- one observed in the adult. Neuronal develop- ferent regenerative capacities of each lineage? ment has been studied in indirect developing species, in particular regarding the development Acknowledgements We thank Bernhard Egger for of the apical organ. Neurogenesis starts with the providing platyhelminth images showed in Fig. 3.1 , the specifi cation of a few apical serotonin- and schemes of platyhelminth embryogenesis in Fig. 3.2 , and the images of larvae in Fig. 3.3 . We thank Iain Patten for FMRFamide-like-positive cells that establish the advice on the English. This work was supported by grant brain and apical plate primordia and the forma- BFU2012-31701 (Ministerio de Economía y tion of posterior neuronal cell bodies. From the Competitividad, Spain) to F.C; grant BFU2008-01544 brain, dorsolateral projections extend posteriorly (Ministerio de Economía y Competitividad, Spain) to ES and TA; grant 2009SGR1018 (Agència de Gestió d’Ajuts to meet the posterior neurons, and ventro-lateral Universitaris i de Recerca) to ES, FC, and TA; and grant projections innervate the mouth and larval lobes. AIB2010DE-00402 (Ministerio de Economia y How this larval neural pattern metamorphoses Competitividad Accion Integrada). 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Francesc Cebrià , Emili Saló , and Teresa Adell

Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger.

F. Cebrià • E. Saló (*) • T. Adell Department of Genetics, Faculty of Biology , Institute of Biomedicine, University of Barcelona , Av. Diagonal 643, edifi ci Prevosti, planta 1 , Catalunya, Barcelona 08028 , Spain e-mail: [email protected]

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 41 DOI 10.1007/978-3-7091-1871-9_4, © Springer-Verlag Wien 2015 42 F. Cebrià et al.

ADULT DEVELOPMENTAL PROCESSES: REGENERATION IN PLATYHELMINTHES

Some species of Platyhelminthes have become model systems in which to study whole-body regeneration in adults. Before describing how this capacity is distributed and varies within the phylum, however, it is important to introduce the adult pluripotent stem cells that confer this remarkable ability in fl atworms, the so-called neoblasts.

Fig. 4.1 Electron micrograph of a planarian neoblast. A neoblast is a small cell (close to 10 μm) with a large Neoblasts: The Pluripotent Stem Cells nucleus ( N ) and a small cytoplasm (C ). Arrow indicates of Adult Platyhelminthes the chromatoid bodies usually observed in this cell type. Scale bar: 0.5 μm A common feature that characterises the phylum Platyhelminthes is the presence of neoblasts, the modifi cation are also present in neoblasts (Rossi only cell type that proliferates in adult somatic tis- et al. 2007 ; Fernández-Taboada et al. 2010 ; sues. The term was coined by Harriet Randolph in Rouhana et al. 2010; Labbe et al. 2012; Onal et al. 1892 to refer to undifferentiated embryonic- like 2012 ; Wagner et al. 2012 ). Although the ontoge- cells observed during the formation of new meso- netic origin of neoblasts is not clear, it has recently derm after fi ssion in earthworms (Annelida). been suggested that they correspond to primordial Later, in 1897 , she extended the term neoblasts to stem cells (PriSCs) (Solana 2013 ). These PriSCs similar staining observed in smaller cells present are thought to constitute stem cells intercalated in planarians (Tricladida) (for a historical review, between the zygote and the germ line. Depending see Baguñà 2012 ; Rink 2013 ; Adell et al. 2014). on the reproductive mode and regenerative capa- The term ‘neoblast’ refers to adult stem cells that, bilities of the different animals, the PriSCs may in triclads, are scattered throughout the fl atworm give rise to mainly primordial germ cells or other body and are only absent in the animal head tip types of somatic cells. The pluripotency of the and the pharynx. Neoblasts are the only cells that neoblasts was fi rst hypothesised many decades exhibit mitotic activity and the capacity to differ- ago (reviewed in Brϕndsted 1969; Baguñà 2012 ; entiate into all somatic and germinal cell types. see also, Baguñà et al. 1989a; Ladurner et al. Neoblasts can be easily identifi ed by their small 2000 ; Newmark and Sánchez-Alvarado 2000 ). size (7–12 μm in diameter), spindle shape, and the However, it has only recently been shown that an presence of a single fi lopodium and a high nuclear individual neoblast is capable of differentiating to cytoplasmic ratio (Fig. 4.1 ). At the ultrastruc- into all cell types in the planarian Schmidtea med- tural level, the neoblasts contain chromatoid bod- iterranea (Wagner et al. 2011 ). ies close to the nucleus (Auladell et al. 1993 ); Although all Platyhelminthes possess neo- these round structures lack membranes and are blasts, not all of them are capable of regenerat- composed of ribonucleoproteins, and they resem- ing. In Platyhelminthes that do not regenerate, ble the germ granules found in the germ cells of neoblasts are probably required for the homeo- other animals. Germ granules are thought to func- static cell turnover observed in adult animals. As tion as centres for post-transcriptional regulation in the case of their free-living relatives, pluripo- of mRNA (Extravour and Akam 2003 ), and sev- tent stem cells are fundamental in the develop- eral studies have shown that genes involved in ment of parasitic Platyhelminthes, where they are post-transcriptional regulation and chromatin called germinative, germinal, or regenerative 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 43 cells. Despite the different nomenclature, the several species of triclads and polyclads have basic processes of cell proliferation and differen- been linked to the history of regeneration research tiation are shared between free-living and para- since its very beginning, as Pallas already sitic Platyhelminthes. Neoblasts are considered described these striking abilities in 1774 (Pallas fundamental for the complex plastic life cycles of 1774 ; for a historical revision, see Brϕndsted parasitic species, which are subject to regulation 1969 ). The fact that some species are capable of by host signals. In cestodes, systems have been regenerating a complete animal from a tiny piece developed for the short-term culture of neoblast of their bodies (something quite unique among populations from asexually multiplying larval bilaterians) led Dalyell to claim that planarians stages (Brehm 2010 ). Despite some similarities, were ‘almost immortal under the edge of the the germinative cells do not contain chromatoid knife’ (Dalyell 1814 ). bodies and they do not express vasa and piwi Regeneration has quite a broad distribution in gene orthologs, suggesting fundamental differ- the animal kingdom (Birnbaum and Sánchez ences between these and other neoblast cell pop- Alvarado 2008 ; Bely and Nyberg 2010 ; Poss ulations (Collins et al. 2013; Koziol et al. 2014 ). 2010 ), since many phyla contain species with In Taenia solium , there is a germinative region some regenerative capability. However, it is also posterior to the apex of the scolex, the neck, in true that closely related species can display a which stem cells proliferate continuously, differ- very different regenerative potential (e.g., the entiate into all cell types, and migrate to the tegu- ability to regenerate entire limbs of some amphib- ment (Merchant et al. 1997 ). In trematodes, two ians versus the complete absence of regeneration populations of neoblast-like cells have been iden- in most other vertebrates). Also, regeneration can tifi ed in the species Schistosoma mansoni . occur at different biological levels of organisa- Somatic stem cells have been identifi ed in adult tion: cell, tissue, organ, structure, or the whole schistosomes (Collins et al. 2013 ) and in the spo- body (Bely and Nyberg 2010). Thus, on the one rocyst, the larval stage that lives in the intermedi- hand, regeneration is broadly distributed, but on ate mollusk host (Wang et al. 2013 ). These data the other, the distribution and potential of this suggest that schistosome adult stem cells persist biological property is uneven. This has raised the through its life cycle. Transcriptomic analysis question of whether regeneration is an ancestral supports the existence of a molecular signature trait that was lost multiple times through evolu- in free-living Platyhelminthes and Trematoda, tion (Bely 2010; Bely and Nyberg 2010 ) or, suggesting an ancient role for these genes in reg- based on some taxon-specifi c components, that it ulating the stem cell population of Platyhelminthes appeared independently in different regeneration- (Collins et al. 2013 ; Wang et al. 2013 ). competent lineages (Garza-García et al. 2010 ). Based on the observation that different models use the same conserved signalling pathways or Regenerative Capabilities genetic programmes to regenerate similar struc- tures, placing regeneration as an ancestral trait at Several species of Platyhelminthes are known to the base of metazoans seems plausible. In this have amazing regenerative capabilities, under- sense, the loss of the capacity to reactivate those stood as the capacity to regrow parts of their bod- conserved pathways after tissue loss could ies lost after a traumatic amputation. Among the explain the loss of regeneration in many species. most popular current animal models for regener- Recently, studies on annelids have provided some ation, triclads such as Schmidtea mediterranea examples of evolutionary loss of regeneration and Dugesia japonica (Newmark and Sánchez (Bely 2010 ). However, if regeneration is not Alvarado 2002; Agata 2003 ; Saló 2006 ; Gentile ancestral but evolved independently in different et al. 2011 ), as well as the macrostomid lineages, one might expect to identify taxon- Macrostomum lignano (Egger et al. 2006 ; Morris specifi c factors, genes, or programmes in those et al. 2006 ), occupy a prominent position. In fact, regeneration-competent species. Not many 44 F. Cebrià et al. examples of such specifi c regeneration- promoting 2007 ). Among model fl atworm species for regen- genes have been described (Poss 2010 ). One of eration studies, Schmidtea mediterranea and them is Prod1 , which appears to be restricted to Dugesia japonica can regenerate a whole animal salamanders and is essential for limb regenera- from almost any tiny piece of their bodies, whereas tion in these animals (Garza-García et al. 2010 ). Macrostomum lignano can regenerate as long as As more transcriptomic and genomic data the brain and pharynx are present. Even among tri- become available for a larger number of species clads, which include many species that can regen- with and without regenerative abilities, compara- erate a whole animal from a tiny piece of their tive studies should provide insights into the evo- bodies, some species show more restricted regen- lutionary origin of regeneration. erative abilities than others. In fact, up to fi ve to As in other phyla, not all species of eight types of triclads have been proposed accord- Platyhelminthes display the same regenerative ing to their ability to regenerate (Sivickis 1930 ; abilities (Fig. 4.2). Free-living fl atworms fall in Teshirogi et al. 1977). This classifi cation is mainly one of three different categories. Some groups, based on the capacity to regenerate a head (with such as many rhabdocoels and some lecithoepithe- the cerebral ganglia), which varies depending on liates, cannot regenerate at all. Other groups can the level of amputation along the anterior-posterior regenerate some organs or individual parts but not (AP) axis. Thus, some species will regenerate a others. For example, some species of triclads and head from any body piece, others will regrow the macrostomids can regenerate the posterior region anterior region only if amputated in front of the but not the anterior part with the brain and eyes. pharynx, and others will never regenerate the head Finally, some species of catenulids, polyclads, if amputated behind the cephalic ganglia. macrostomids, and triclads can regenerate any Dendrocoelum lacteum , Procotyla fl uviatilis, and organ and structure (reviewed in Egger et al. Phagocata kawakatsui are good examples of

Fig. 4.2 Phylogenetic Regenerative ability consensus tree of Platyhelminthes. The phylogenetic classifi cation Catenulida Excellent is based in Laumer and Giribet (2014 ), in which Polycladida Good the former

Lecithoepitheliata appears Archoophora Macrostomorpha Excellent as a paraphyletic group, now divided into two new orders (Prorhynchida and Prorhynchida Poor Gnosonesimida). For Lecithoepitheliata simplicity, the old Gnosonesimida ? Lecithoepitheliata name is used here. The regenerative Proseriata Good ability of each order is indicated next to it Bothrioplanida Good (Adapted from Egger et al. ( 2007 ) ) Neodermata Poor Trematoda, Cestoda, Monogenea Neoophora Rhabdocoela Poor

Fecampiida ?

Tricladida Excellent

Prolecithophora Good 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 45

triclads in which postpharyngeal amputation gen- pronounced in taxa containing species with sex- erates tail pieces that are not capable of regenerat- ual reproduction than in taxa that reproduce asex- ing a new head region. In contrast, the anterior half ually, as regeneration is absolutely required for can fully regenerate a new tail. Remarkably, it has survival of the mother animal during asexual recently been shown that the silencing of a single reproduction (Egger et al. 2007 ). Some species, gene, a β -catenin homolog, is able to rescue ante- such as Microstomum lineare , combine sexual rior regeneration in these tail pieces, indicating and asexual modes of reproduction. In these fl at- that they retain regenerative potential but that, worms, a good regenerative capacity is seen dur- under natural conditions, this programme is not ing the asexual stage of their life cycle, but this activated (Liu et al. 2013 ; Sickes and Newmark capacity is reduced with the appearance of sexu- 2013 ; Umesono et al. 2013 ). ality (Reuter and Kreshchenko 2004 ). Although parasitic species of Platyhelminthes Asexual reproduction by fi ssion may occur by outnumber free-living fl atworms, much less is paratomy or architomy. During paratomy, the dif- known about their regenerative capabilities. Senft ferentiation of the organs, such as the new and Weller (1956 ) cultured immature specimens of cephalic ganglia or eyes, precedes the fi ssioning Schistosoma mansoni and observed that in some itself. Often, the ‘mother’ worm is transformed cases in which the animals had been damaged in the into a chain of zooids that will later split into sev- isolation procedure, they could regenerate the miss- eral independent ‘daughter’ fl atworms. In con- ing parts in vitro. Also, during the life cycle of trast, during architomy, fi ssioning precedes the schistosomes, infectious cercariae transform into differentiation of the new organs of the daughter schistosomula (the larval stage of Schistosoma in fl atworm. In fact, no external signs can be distin- their vertebrate hosts) after they lose their tails when guished before the fi ssion process starts. Despite they enter their defi nitive host. In vitro, tails of this lack of external signs and organ differentia- detached cercariae are able to grow some tissues tion before fi ssioning, some molecular changes resembling a head in shape although they die within have been described in animals committed to a few days (Coultas and Zhang 2012 ). It has been asexual reproduction (Bueno et al. 2002 ). also described that the cestode Diphyllobothrium Architomy is common in triclads, whereas para- erinacei can regenerate. Isolated larvae of this spe- tomy is often found in several species of catenu- cies were subject to different types of injuries and lids and macrostomids (Reuter and Kreshchenko amputations and were able to regenerate after they 2004). After architomic fi ssioning, the tiny body had been transferred to their intermediate and defi n- piece that is produced must regenerate a new itive hosts (Vorontsova and Liosner 1960). More organism. In this sense, architomy and regenera- recently, it has been shown that primary cells from tion after amputation are very similar, as they Echinococcus multilocularis are able to form proceed through similar stages. cell aggregates from which young metacestode (lar- val stage of tapeworms) vesicles develop in vitro (Spiliotis et al. 2008). Nevertheless, further experi- Mechanisms of Regeneration ments are needed to determine the regenerative capabilities of larval and adult parasitic As described above some triclads and macrosto- Platyhelminthes. mids are among current model species for regen- The regeneration potential within this phylum eration. In these species, regeneration requires is often associated with the reproductive mode. cell proliferation and blastema formation, and Platyhelminthes are, with few exceptions, her- they can be categorised as examples of epimor- maphroditic (Hyman 1951 ). However, asexual phic regeneration, together with, for example, reproduction by fi ssion occurs in some catenulids amphibian limb regeneration. The term epimor- and macrostomids, and it is quite common among phosis was coined by Morgan (1901 ) to refer to triclads (Hyman 1951 ; Reuter and Kreshchenko the regeneration mode ‘in which proliferation of 2004 ). As expected, regenerative abilities are less material precedes the development of the new 46 F. Cebrià et al. part’. This was in contrast to morphallaxis ‘in blastema formation, which fi ts with epimorphic which a part is transformed directly into a new regeneration. The term morphallaxis, however, is organism or part of an organism without prolif- more problematic because it has been mainly eration at the cut-surfaces’. However, as Morgan associated with tissue remodelling in the absence already realised, these two modes of regeneration of cell proliferation. However, it has been shown are not mutually exclusive. In fact, planarians are that tissue remodelling in planarians also depends an example in which a mixed epimorphic/mor- on the presence of stem cells (Gurley et al. 2010 ). phallactic model has been proposed (Saló and Therefore, it would probably be better to use the Baguñà 1984 ) because, in addition to cell prolif- term ‘remodelling’ instead of ‘morphallaxis’, eration at the cut surface, there is a remodelling without linking it necessarily to the presence or of the pre-existing tissues far from the wound in absence of cell proliferation. order to attain the correct body proportions Regeneration implies three main stages: (1) (Fig. 4.3 ). Over the years, some debate has arisen wound closure and healing, (2) cell proliferation about the convenience of using these terms as and blastema formation, and (3) cell differentia- originally defi ned by Morgan. Most fl atworms tion and morphogenesis. In triclads, wound clo- regenerate through cell proliferation-dependent sure is mediated by the contraction of the body

Fig. 4.3 Remodelling of pre-existing tissues during pre-existing structures remodel to reach a new propor- regeneration in Schmidtea mediterranea . During the pro- tioned animal. Scale bar: 1 mm cess of regeneration while new tissue is regenerating, the 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 47 wall muscle fi bres at the cut surface, followed by cell types (Kragl et al. 2009; Knopf et al. 2011 ). the spreading of epithelial cells surrounding the This suggests that the blastema is in fact heteroge- amputation site (Pedersen 1976 ; Hori 1989 ). A neous and formed by distinct populations of lin- similar process was observed in Microstomum eage-restricted progenitor cells (Tanaka and (Palmberg 1986 ), Catenula (Moraczewski 1977 ), Reddien 2011). Remarkably, this scenario may and Macrostomum lignano (Egger et al. 2009 ). also occur in triclads, based on the existence of However, in Microstomum , the new epidermis is distinct cell populations of progenitor- like neo- formed underneath the wound epidermis, whereas blasts that express specifi c transcription factors in other fl atworms the new epidermis forms by required for their differentiation into eye pigment, the insertion of new cells from the parenchyma photoreceptors, protonephridia, or different neuro- (Palmberg 1986 ; Rieger et al. 1991 ). nal types (Lapan and Reddien 2011 ; Scimone In most cases, cell proliferation at the cut- edges et al. 2011 ; Cowles et al. 2013 ; Currie and Pearson results in the formation of a blastema, an unpig- 2013 ; März et al. 2013 ; Reddien 2013 ). mented mass of undifferentiated cells where the Another difference between vertebrate and missing organs and structures will differentiate. some fl atworm blastemas concerns the prolifera- Remarkably, there are also a few examples (e.g., tive activity of their constituent cells that form Microstomum lineare and some catenulids) in them. Thus, in amphibians and zebrafi sh, cell which this cell proliferation does not result in blas- proliferation drives blastema growth. In contrast, tema formation and the new tissues and organs proliferation in triclads is mainly concentrated in regenerate within the pre-existing tissues after the the stump region adjacent to the blastema, and migration of proliferating cells from regions far very few mitoses are observed within the blas- away from the wound (Palmberg 1986 ; Rieger tema itself (Baguñà 1976; Saló and Baguñà et al. 1991 ). In general, most fl atworms regrow the 1984 ). This is more obvious in anterior blastemas missing parts from a regenerative blastema, simi- than posterior ones, where more mitotic cells are lar to the process of amphibian limb regeneration. observed (Wenemoser and Reddien 2010 ). Thus, However, there are some important differences in triclads such as Schmidtea mediterranea , ante- between the amphibian and fl atworm blastemas. rior blastemas mainly grow by the migration of One main difference concerns the origin of the post-mitotic neoblasts (probably already com- cells that form the blastema. In amphibians, fol- mitted) from the stump region. In contrast, the lowing amputation, pre- existing differentiated blastema of Macrostomum lignano and the caten- cells of the stump dedifferentiate, re-enter the cell ulid Paracatenula galateia contains proliferative cycle, proliferate, and re-differentiate into new cells (Nimeth et al. 2007 ; Egger et al. 2009 ; Dirks cells to restore the missing structures. In contrast, et al. 2012 ; Verdoodt et al. 2012 ). Despite this regeneration in fl atworms depends upon the pres- difference in the distribution of the proliferating ence of neoblasts. Upon amputation, neoblasts neoblasts, upon amputation, a similar biphasic proliferate at the wound region, giving rise to the mitotic and S-phase pattern is observed in tri- blastema where they will differentiate (Saló and clads and M. lignano , respectively. An initial Baguñà 1984 ; Wenemoser and Reddien 2010 ). peak in cell proliferation that is more or less uni- Thus, although some examples of transdetermina- form throughout the animal and represents a sys- tion between germ and somatic cells have been temic injury response is followed by a second observed (Gremigni et al. 1980 ), cell dedifferenti- peak restricted to the wound region (Saló and ation does not seem to play a role in fl atworm Baguñà 1984 ; Wenemoser and Reddien 2010 ; regeneration. Verdoodt et al. 2012 ). The classical view of a blastema as a mass of In summary, Platyhelminthes include some undifferentiated homogeneous cells has been chal- of the most popular model species in which to lenged after the fi nding in amphibians and zebraf- study regeneration. As in other phyla, there is a ish that the dedifferentiated cells keep a memory great variability in regenerative capabilities of their origin and re-differentiate into the same within Platyhelminthes, ranging from species 48 F. Cebrià et al. that are incapable of any regeneration to others heterogeneous population of undifferentiated that can regenerate a whole animal from almost but already committed progenitor cells. Also, in any small part of their bodies. A strong associa- vertebrates and some Platyhelminthes, cell pro- tion exists between asexual reproduction and liferation occurs within these blastemas. In tri- regeneration. In Platyhelminthes, regeneration clads, however, very few mitotic cells are depends upon the presence of adult totipotent observed within them. Whereas cell dedifferen- somatic stem cells and, in most cases, proceeds tiation is the main source of cells for the form- through the formation of a regenerative blas- ing blastemas in vertebrates, in Platyhelminthes tema. In both Platyhelminthes and vertebrates, blastemas appear by proliferation of adult plu- these blastemas appear to be formed by a ripotent stem cells.

Schmidtea mediterranea : A Regeneration Star regions (e.g., anterior or posterior blastemas), Regeneration is one of the most fascinating or individual cell types (e.g., neoblasts and gut phenomena in biology. That some animals are cells) are available. Also, RNAi to study gene capable of regrowing a limb or the whole body function in fl atworms was originally estab- from a tiny piece of them awakens our imagina- lished in S. mediterranea and then adapted to tion and may even make us feel jealous! other species. Neoblasts were fi rst labelled with Freshwater planarians (Tricladida) have enor- BrdU in this fl atworm and the totipotency of mous regenerative power. You can cut them single neoblasts in vivo was also probed for the into many pieces and get many complete worms fi rst time in this species. S. mediterranea has in just a few days. Because of the increasing served as a model to understand how the ante- interest in stem cell research, a growing num- rior-posterior and dorsoventral axes are re- ber of laboratories around the world have established during fl atworm regeneration. All turned their attention to these animals. The spe- these resources, tools, and studies have contrib- cies Schmidtea mediterranea has emerged as a uted to the establishment of S. mediterranea as regeneration star. This tiny fl atworm was origi- an excellent model system to study different nally described in the 1970s and has become a aspects of regeneration, such as neoblast main- model species used in many laboratories. tenance and self-renewal, blastema formation, Remarkably, most of the clonal lines used in axial polarity, neural regeneration, growth and research nowadays derive from a population degrowth, and the relationship between regen- originally obtained from a small fountain in eration and cancer. The results obtained so far Barcelona. S. mediterranea exists in two forms: have been relevant not only to understand sexual and asexual strains. Asexual animals regeneration in fl atworms but also from a reproduce by fi ssion and are the only reported broader evolutionary perspective. Moreover, case of asexuality in this genus. The difference the fact that S. mediterranea exists in sexual between the two strains is a heteromorphic and asexual forms renders it an ideal model for chromosomal translocation. Like other triclads, comparative studies on how patterning and S. mediterranea is easily cultured in the labora- morphogenesis occur in embryonic and post- tory; however, whereas other species with high embryonic contexts. regenerative capabilities are mixoploids or polyploids, S. mediterranea is a stable diploid (2 n = 8). This important feature was taken into account when searching for an appropriate spe- cies to enter the genomics era. The genome of S. mediterranea has been sequenced and is cur- rently under assembly. Moreover, several tran- scriptomes from whole animals, specifi c 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 49

The Central Nervous System (CNS) bodies are peripheral and surround a central neuro- of Platyhelminthes: Pattern pil formed by nerve fi bres (Rieger et al. 1991 ; and Regeneration Reuter and Gustafsson 1995 ; Kotikova et al. 2002 ). The brain is dorsal to the ventral nerve cords, Structure of the CNS which, in some species, project into regions ante- Platyhelminthes have a bilaterally symmetrical rior to the brain. In many species, especially among CNS consisting of anterior cephalic ganglia Tricladida, the brain displays a typical spongy tex- (‘brain’) and one or more pairs of longitudinal ture, as it is traversed by muscles and processes nerve cords that usually run along the length of from secretory cells (Baguñà and Ballester 1978 ). the animal (Fig. 4.4 ); in some catenulids, how- Early phylogenetic offshoots usually have smaller ever, the nerve cords do not reach the posterior brains with few cells lateral to a fi brillar neuropil; end (Rieger et al. 1991 ). The longitudinal cords in later-branching Platyhelminthes the brain is are usually interconnected by commissures that formed by clearly distinguishable bilateral cephalic are more or less regularly spaced in a ladder-like ganglia connected by one or more commissures. In pattern referred to as ‘orthogon’ (Bullock and most of the major systematic groups, there are spe- Horridge 1965 ; Reuter and Gustafsson 1995 ). cies with or without a variably structured brain cap- Submuscular, subepidermal, subepithelial, and sule (Rieger et al. 1991 ). An encapsulated brain has infraepithelial peripheral plexuses are intercon- been described in Polycladida, Lecithoepitheliata, nected with the CNS in most fl atworm taxa and Prolecithophora (Rieger et al. 1991 ). In (Baguñà 1974 ; Baguña and Ballester 1978 ; Rieger trematodes the brain usually contains a pair of et al. 1991; Reuter and Gustafsson 1995). The cephalic ganglia with a central neuropil connected pharynx contains one or two nerve rings that form by a commissure. Miracidium larvae already have a nerve net usually close to the distal part of this a bilobed brain (Bullock and Horridge 1965 ; organ (Bullock and Horridge 1965 ; Baguña and Leksomboon et al. 2012 ). Ballester 1978 ; Rieger et al. 1991 ; Reuter 1994 ). The basic orthogonal plan of the nervous sys- The brain of platyhelminths varies substantially tem of Platyhelminthes varies mainly in the num- in form and size, as well as in the number and ber of longitudinal cords. Whereas Bullock and arrangement of neurons. In most cases (with the Horridge (1965 ) described nine types of exception of adult Polycladida), the neuronal cell orthogons, Kotikova ( 1986 , 1991 ) distinguished

Fig. 4.4 Structure of the CNS in Platyhelminthes. more prominent than in P. ullala (Tricladida, Immunostaining with anti-synapsin in Phagocata ullala , Continenticola) The image of M. lignano was originally Camerata robusta , and Schistosoma mansoni . published by Egger et al. (2007 ). The image of S. mansoni Immunostaining with anti-GYRFamide in Macrostomum was originally published by Collins et al. (2011 ). cg lignano. In P. ullala , C. robusta, and S. mansoni, the ven- cephalic ganglia, vnc ventral nerve cords, lnc lateral nerve tral nerve cords are the main cords. In M. lignano the lat- cords, ph pharynx. Anterior is to the left in all aspects. eral nerve cords are the main cords. In C. robusta Scale bars: P. ullala and C. robusta 0.5 mm; M. lignano (Tricladida, Maricola) the lateral nerve cords are much 100 μm; S. mansoni 50 μm 50 F. Cebrià et al. eight different types. According to Kotikova, the longitudinal cords, which is accompanied by a body shape mainly determines the type of strengthening of the remaining cords. There is orthogon, which would be basically homologous also a trend towards concentration of plexal fi bres and derived from the pattern observed in to longitudinal cords. Whereas the main cords are Catenulida. The presence of three to fi ve pairs of localised ventrally in various lineages of cords (seen in some Rhabdocoela and Proseriata) Neoophora, the main cords occupy a more lateral is considered more primitive, with an evolution- position in the earlier-branching taxa (Catenulida ary tendency towards reducing the number of and Macrostomida) (Reisinger 1972 ; Joffe and cords to one prominent pair (in macrostomids, Reuter 1993 ; Reuter and Gustafsson 1995 ; Reuter Rhabdocoela, and Tricladida) (Rieger et al. and Halton 2001 ). 1991 ). The nerve cords can be placed at different Traditionally, the CNS of the Platyhelminthes positions: dorsal, lateral, marginal, ventrolateral, has been considered as primitive (Bullock and and ventral. Reuter and Gustafsson ( 1995 ) pro- Horridge 1965 ) and, therefore, most similar to posed that, independently of their position, the the urbilaterian nervous system. However, cur- term main cord should be applied to those cords rent phylogenetic analyses do not support this that (1) start as multifi bre outgrowths from the basal position of the Platyhelminthes within brain and (2) are thicker and possess more neu- Bilateria (Riutort et al. 2012 ); moreover, Acoela rons positive for serotonin and catecholamine and Nemertodermatida, previously included markers. In contrast, all other cords are thinner within the Platyhelminthes, are now placed in a and have less contact with the brain. separate phylum, the Acoelomorpha (Ruiz-Trillo In Catenulida, Macrostomida, and et al. 1999 ). The position of the Acoelomorpha is Monocelididae (Proseriata), the main cords are still under debate, as they have been considered usually lateral (Reuter et al. 1986 ; Reuter 1988 ; basal bilaterians or, more recently, basal deutero- Joffe and Kotikova 1991 ; Reiter and Wikgren stomes (Philippe et al. 2011 ). Remarkably, 1991 ; Joffe and Reuter 1993 ). In Tricladida and although acoelomorphs and platyhelminths have Lecithoepitheliata the ventral cords are the main been separated in evolution at least by half a bil- ones. In the Prolecithophora and Rhabdocoela, lion years (Peterson and Eernisse 2001 ; Peterson either the lateral or ventral cords dominate or the et al. 2008 ), their CNS displays many similarities lateral and ventral ones are equally developed. For in the architecture of their brains and orthogons, example, in the rhabdocoel Castrella truncata , the as well as in some developmental features (Bailly lateral and ventral cords are about the same thick- et al. 2013). This has led some authors to suggest ness, although the ventral ones are associated with that the morphological features of the platyhel- more serotonergic neurons and would then be con- minth CNS could refl ect some characteristics of sidered the main cords (Kotikova et al. 2002 ). In their urbilaterian ancestor (Bailly et al. 2013 ; but Trematoda there are usually three pairs of longitu- see Semmler et al. 2010 ). dinal cords (dorsal, lateral, and ventral); the ven- tral cords are usually the largest (and thus constitute Neuronal Diversity and Molecular the so-called main cords), and they are often inter- Complexity connected at the caudal tip (Bullock and Horridge In contrast to other invertebrates, platyhelminths 1965; Leksomboon et al. 2012 ). In the Cestoda, a have a large variety of neural cells, including variable number of longitudinal cords may occur, uni-, bi-, and multipolar neurons (Rieger et al. from one pair to up to 38 or 60 cords in other spe- 1991 ). A characteristic of fl atworm neurons is cies; the lateral cords are often the main cords their high content of vesicles, which are thought (Bullock and Horridge 1965 ; Joffe and Reuter to be secretory. Examples include small clear 1993 ). vesicles (often regarded as cholinergic), dense- Several evolutionarily trends can be observed core vesicles (aminergic), and large dense vesi- in the orthogon and main cords. Firstly, there cles (peptidergic) (Reuter and Gustafsson seems to be a reduction in the number of 1995 ). The neurons of some catenulids and 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 51

macrostomids seem to be structurally least dif- mones are often enriched in specifi c cell types ferentiated, whereas the vesicles of others show or regions within the CNS (Collins et al. 2010 ), a greater heterogeneity (Reuter and Gustafsson indicating a high neuronal diversity. 1995 ). The complexity of the fl atworm nervous sys- A large number of aminergic and peptidergic tem at the cellular and molecular levels is further neuroactive substances have been detected in supported by analysis of the expression of neural most fl atworm taxa, both in free-living and par- genes. Until now, the expression of dozens of asitic species (Fairweather and Halton 1991 ; neural genes have been reported and are used to Reuter and Gustafsson 1995 ; Gustafsson et al. identify distinct neuronal populations (Mineta 2002 ; Ribeiro et al. 2005 ; Cebrià 2008 ; et al. 2003 ; Nakazawa et al. 2003 ; Cebrià 2007 ; McVeigh et al. 2011 ). These markers have been Cebrià et al. 2010 ). Most of these studies have used to describe the architecture of the nervous been done in triclads such as Schmidtea mediter- system (Fig. 4.5 ) and have revealed the high ranea or Dugesia japonica and revealed how neuronal diversity in fl atworms (see Table 4.1 their brains, although quite simple at the morpho- for a list of Platyhelminthes in which specifi c logical level, can be subdivided into different neuronal populations have been identifi ed). As molecular compartments. Thus, for example, pla- Platyhelminthes do not possess true endocrine narian homologs of the otx / otp homeobox genes glands and a circulatory system, the nervous defi ne different domains along the medio-lateral system can be seen as a neuroendocrine system axis of the cephalic ganglia, as otxA is expressed that controls processes such as growth, devel- in the most medial region, otxB in the central opment, and regeneration (Fairweather and neuropil, and otp in the brain lateral branches Halton 1991 ). Immunohistochemistry for many (Umesono et al. 1999 ). Smed - netrinA is also of these neuroactive substances has shown that mainly expressed in the medial region of the they are most often expressed in distinct, cephalic ganglia (Cebrià and Newmark 2005 ). nonoverlapping neuronal populations. More Other genes specifi cally expressed in the lateral recently, 51 prohormone genes and 142 pep- branches are Djnlg (noggin-like gene, Ogawa tides have been identifi ed and characterised et al. 2002a ; Molina et al. 2009 ), 1008HH (a glu- biochemically in the triclad Schmidtea mediter- tamate receptor homolog, Cebrià et al. 2002a ), ranea. Of these 51 genes, 85 % are expressed in Gtsix3 ( Six / sine oculis homolog, Pineda and Saló the CNS and, remarkably, different prohor- 2002), and 1791HH (a G protein alpha subunit,

ABC

Fig. 4.5 Neuronal populations in the platyhelminth interconnect the ventral cords. ( B ) Double immunostain- Schmidtea mediterranea . (A ) Ventral nerve cords (in ing with anti-5-HT (in red ) and anti-allatostatin (in green ) green after immunostaining with anti-synapsin) sur- along the ventral nerve cords. ( C ) Double immunostain- rounded by serotonergic neurons (in red after immunos- ing with anti-5-HT (in red ) and anti-GYRFamide taining with anti-5-HT (serotonin)). Bipolar serotonergic (in green ) along the ventral nerve cords. Scale bars: neurons are also seen in the transverse commissures that ( A ) 100 μm; ( B , C ) 50 μm 52 F. Cebrià et al.

Table 4.1 Summary of distinct neuronal populations in Platyhelminthes Neuronal population Species Order Reference Serotonin (5-HT) Schmidtea mediterranea Tricladida Cebrià (2008 ), März et al. (2013 ), Currie and Pearson (2013 ) Dugesia japonica Tricladida Nishimura et al. (2007a ) Microstomum lineare Macrostomida Reuter et al. (1986 ) Girardia tigrina Tricladida Reuter et al. (1995b ) Polycelis tenuis Tricladida Reuter et al. (1996a ) Fasciola hepatica Echinostomida Gustafsson et al. (2001 ) Diphyllobothrium dendriticum Pseudophyllidea Lindholm et al. (1998 ) Castrella truncata Rhabdocoela Kotikova et al. (2002 ) Bothriomolus balticus Seriata Joffe and Reuter (1993 ) Stenostomum leucops Catenulida (class) Reuter et al. (2001 ) Echinococcus multilocularis Cyclophyllidea Koziol et al. (2013 ) Cryptocotyle lingua Plagiorchiida Pan et al. (1994 ) Schistosoma mansoni Strigeidida Gustafsson (1987 ) Macrostomum h. marinum Macrostomida Ladurner et al. (1997 ) Dendrocoelum lacteum Tricladida Reuter et al. (1996a ) Substance P Microstomum lineare Macrostomida Reuter (1994 ) Stenostomum leucops Catenulida Reuter (1994 ) Schistosoma mansoni Strigeidida Gustafsson (1987 ) Diphyllobothrium dendriticum Pseudophyllidea Gustafsson et al. (1993 ) Neuropeptide F Moniezia expansa Cyclophyllidea Maule et al. (1992 ) Microstomum lineare Macrostomida Reuter et al. (1995c ) Procerodes littoralis Tricladida Reuter et al. (1995a ) Girardia tigrina Tricladida Reuter et al. (1995b ) Archilopsis unipunctata Seriata Reuter et al. (1995c ) Promonotus schultzei Seriata Reuter et al. (1995c ) Stenostomum leucops Catenulida Reuter et al. (1995c ) Schmidtea mediterranea Tricladida Cebrià (2008 ) RFamide Polycelis tenuis Tricladida Reuter et al. (1996a ) Archilopsis unipunctata Seriata Reuter et al. (1995c ) Promonotus schultzei Seriata Reuter et al. (1995c ) Stenostomum leucops Catenulida Reuter et al. (1995c ) Bothriomolus balticus Seriata Joffe and Reuter (1993 ) Dendrocoelum lacteum Tricladida Reuter et al. (1996a ) FMRFamide Girardia tigrina Tricladida Reuter et al. (1996b ) Diphyllobothrium dendriticum Pseudophyllidea Lindholm et al. (1998 ) Castrella truncata Rhabdocoela Kotikova et al. (2002 ) Stenostomum leucops Catenulida Reuter et al. (2001 ) Schmidtea mediterranea Tricladida Fraguas et al. (2012 ) Echinococcus multilocularis Cyclophyllidea Koziol et al. (2013 ) Cryptocotyle lingua Trematoda (class) Pan et al. (1994 ) GYRFamide Girardia tigrina Tricladida Kreshchenko et al. (1999 ) Fasciola hepatica Echinostomida Gustafsson et al. (2001 ) Schmidtea mediterranea Tricladida Cebrià (2008 ) Dopamine Schmidtea mediterranea Tricladida Fraguas et al. (2012 ) Dugesia japonica Tricladida Nishimura et al. (2007b ) 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 53

Table 4.1 (continued) Neuronal population Species Order Reference Octopamine Schmidtea mediterranea Tricladida Fraguas et al. (2012 ) Dugesia japonica Tricladida Nishimura et al. (2008a ) GABA Dugesia japonica Tricladida Nishimura et al. (2008b ) Girardia tigrina Tricladida Eriksson and Panula (1994 )

Cebrià et al. 2002a ). Moreover, some genes fl atworm and acoelomorph taxa will be required defi ne different domains along the anterior- to determine the extent to which the CNS of posterior axis; thus, for example, DjWntA (a Wnt Platyhelminthes would resemble the urbilate- homolog) is mainly expressed in the posterior rian nervous system. half of the brain, whereas DjfzA (a frizzled homo- log) is expressed in the anterior half (Kobayashi Regeneration: Process and Genes et al. 2007 ). Among the genes expressed in the The amazing regenerative capabilities of some triclad CNS, there are homologs of FGF and Platyhelminthes include the ability to regrow a EGFR receptors (Cebrià et al. 2002c ; Ogawa complete and functional CNS de novo (Fig. 4.6 ). et al. 2002b ; Fraguas et al. 2011 , 2014 ); Pax6 This remarkable feature has been best described (Pineda et al. 2002 ); neural cell adhesion mole- in triclads (Cebrià 2007 ; Agata and Umesono cules (Fusaoka et al. 2006 ); nicotinic acetylcho- 2008 ; Umesono et al. 2011 ; Fraguas et al. 2012 ). line receptors (Mineta et al. 2003 ); roundabout Table 4.2 summarises some of the genes that play (Cebrià and Newmark 2007 ); mushashi (Higuchi relevant roles in planarian CNS regeneration. et al. 2008 ); FoxG1 (Brain factor-1) (Koinuma Upon head amputation, new cephalic ganglia et al. 2003 ); SNF2 - like (Rossi et al. 2003 ); eye develop within the blastema at the same time that absent (Mannini et al. 2004 ); innexins (Nogi and the truncated ventral nerve cords grow to re- Levin 2005 ); Smad (Molina et al. 2007 ); mem- establish connections with the forming brain. The bers of the SET1/MLL family of histone methyl- whole process can be divided into three main transferases (Hubert et al. 2013 ); components of stages: (1) brain primordia formation and pat- the Wnt/β-catenin signalling pathway such as terning, (2) re-establishment of connectivity, and β - catenin , apc , and dishevelled (Gurley et al. (3) functional recovery. After 1–2 days of regen- 2008 ; Iglesias et al. 2008 ; Petersen and Reddien eration, two small bilateral clusters of cells cor- 2008 ); hedgehog (Yazawa et al. 2009 ); and sev- responding to the primordia of the new cephalic eral RNA-binding proteins (Guo et al. 2006 ; ganglia are observed (Cebrià et al. 2002b ; Rouhana et al. 2010 ; Wagner et al. 2012 ). Kobayashi et al. 2007). The appearance of the In 2003, Mineta et al. published the only brain primordia depends on the neoblasts. Until high- throughput study so far of planarian neural recently, it was not clear whether neural progeni- genes from a comparative evolutionary perspec- tors exist in planarians. However, recent studies tive. From 3,101 nonredundant ESTs of a pla- have reported that some neoblasts already express narian head library, they identifi ed 116 genes distinct transcription factors that defi ne different that showed clear homology to genes related neuronal populations. Remarkably, these tran- to the nervous system in other animals. scription factors correspond to conserved genes Remarkably, more than 95 % of these genes had that have similar functions in neurogenesis of homologs in Drosophila , Caenorhabditis ele- other invertebrate and vertebrate species. For gans , and humans, indicating a high degree of example, a pitx homolog defi nes the serotonergic conservation. However, a broad systematic lineage during cell renewal and regeneration comparison of the neural genes of different (Currie and Pearson 2013 ; März et al. 2013 ). 54 F. Cebrià et al.

AB C

D EF

G

Fig. 4.6 Posterior regeneration of the CNS in Schmidtea formed (white arrowheads ). Arrows point to thin sprouts mediterranea. All images show anti-neuropeptide F near the posterior tip. Yellow arrowheads point to the new immunostaining in head fragments regenerating a new ganglia-like knots of the regenerated ventral nerve cords. posterior region. ( A ) Arrows point to thin fi bres sprouting ( G ) Schematic drawings of the posterior regeneration of from the truncated nerve cords. ( B ) Arrowhead points to a the ventral nerve cords. Cephalic ganglia are in red and transverse commissure connecting left and right nerve ventral nerve cords in green . Days of regeneration are cords. ( C ) Arrows point to fi bres sprouting posteriorly. indicated in ( A – F ). Asterisks mark the newly regenerated ( D) Regenerating ventral nerve cords keep growing poste- pharynx. Anterior to the top left in ( A – F ). Anterior to the riorly ( arrows ). (E ) Newly formed transverse commis- top in ( G ). Scale bar, 200 μm (Reproduced with permis- sures ( arrowhead) from sprouting fi bres ( arrows ). (F ) As sion from The International Journal of Developmental regeneration proceeds, new transverse commissures are Biology (Int. J. Dev. Biol. 2012 vol. 56:143–153)) 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 55

Table 4.2 Selection of genes required for planarian CNS regeneration Planarian gene Homolog RNAi phenotype Reference nou - darake Fibroblast growth Ectopic brain differentiation in Cebrià et al. (2002c ) factor receptor - like posterior regions Smed - netrin2 Netrin Disorganised meshwork of Cebrià and Newmark axonal projections (2005 ) Mis-targeting of visual axons Smed - netR Netrin receptor Disorganised meshwork of Cebrià and Newmark axonal projections ( 2005 ) Mis-targeting of visual axons DjCAM N - CAM (neural cell Defasciculation of brain axonal Fusaoka et al. (2006 ) adhesion molecule) bundles DjDSCAM DSCAM (Down Neuropil disorganised and Fusaoka et al. (2006 ) syndrome cell adhesion reduced number of brain lateral molecule) branches Smed - slit Slit CNS collapsed at the midline Cebrià et al. (2007 ) Smed - roboA Roundabout Abnormal reconnection of the Cebrià et al. (2007 ) cephalic ganglia and the ventral nerve cords Mis-targeting of visual axons DjCHC Clathrin heavy chain Failed neurite extension and Inoue et al. (2007 ) maintenance DjWntA Wnt Brain expansion towards Kobayashi et al. (2007 ) posterior regions Smed - GSK3sa Glycogen synthase Smaller and disconnected Adell et al. (2008 ) kinase - 3 cephalic ganglia Mis-targeting of visual axons Smed - evi / Wntless Evenness interrupted Disconnected and laterally Adell et al. (2009 ) ( Wntless / Sprinter ) displaced cephalic ganglia Smed - Wnt - 5 Wnt Disconnected and laterally Adell et al. (2009 ), displaced cephalic ganglia Almuedo-Castillo et al. Mis-targeting of visual axons (2011 ), Gurley et al. (2010 ) Smed - dvl1 / 2 Dishevelled Disconnected and laterally Almuedo-Castillo et al. displaced cephalic ganglia (2011 ) Mis-targeting of visual axons Smed - lhx1 / 5 - 1 Lhx Loss of serotonergic neurons Currie and Pearson (2013 ) Smed - pitx Paired class Loss of serotonergic neurons Currie and Pearson homeobox / pituitary (2013 ), März et al. homeobox (2013 ) Smed - coe Collier / olfactory - Smaller and disconnected Cowles et al. (2013 ) 1 / early B - cell factor cephalic ganglia Smed - sim Single - minded Reduced neuropil density Cowles et al. (2013 ) Smed - egr - 4 Early growth Failed differentiation of the Fraguas et al. (2014 ) response brain primordia Dj Dugesia japonica , Smed Schmidtea mediterranea a For GSK3s, drug-mediated inhibition was performed instead of RNAi-mediated gene silencing 56 F. Cebrià et al.

More recently, Cowles et al. ( 2013 ) identifi ed 44 of Platyhelminthes (Younossi-Hartenstein and genes predicted to code for a bHLH (basic helix- Hartenstein 2000 ; Younossi-Hartenstein et al. loop- helix) domain, of which 12 are expressed in 2000 ; Hartenstein and Jones 2003 ; Cardona the CNS and neoblasts. Some of these genes are et al. 2005; Rawlinson 2010; Bailly et al. 2013 ), co-expressed with different markers of specifi c much less is known about the genes expressed neuronal populations: cholinergic, GABAergic, at the different embryonic stages and their puta- octopaminergic, dopaminergic, and serotonergic. tive functions during neurogenesis. It would be More importantly, two of these genes, coe interesting to characterise how neurogenesis ( collier / olfactory - 1 / early B - cell factor ) and sim occurs at the molecular level in developing ( single - minded) are co-expressed with proliferat- embryos and compare it with the regenerative ing neoblasts and contribute to the regenerative process. blastema, further supporting the existence of neural progenitor cells in planarians. The Role of the CNS in Regeneration The second stage in CNS regeneration It has been suggested that the nervous system involves growth of the brain primordia into might play an important role during regeneration mature cephalic ganglia and the extension of in several models (Kumar and Brockes 2012 ). the amputated ventral cords into the blastema. For example, it is well known that the amphibian This leads to the rewiring of the regenerating limb cannot regenerate if previously denervated nervous system. Several families of axon guid- (Singer and Craven 1948). Also, head regenera- ance cues are evolutionarily and functionally tion in Hydra depends on de novo neurogenesis conserved and have been shown to play impor- (Miljkovic-Licina et al. 2007 ). However, fi nding tant roles in the wiring of the developing ner- the exact nature of such neurally derived mole- vous system. Among these cues, netrin and slit cules has been very elusive. In newts, a molecule occupy a prominent position (Guan and Rao called nAG appears to be responsible for the 2003 ; O’Donnell et al. 2009 ). Planarians have nerve dependence of limb regeneration (Kumar allowed analysing the function of such cues et al. 2007 ). However, in most cases the molecu- during the rewiring of the regenerative nervous lar basis for nerve-dependent regeneration system. Interestingly, these cues are not only remains to be characterised. In fl atworms, sev- found in planarians but are also required for eral lines of evidence support a role for the ner- proper CNS regeneration (Cebrià and Newmark vous system in regeneration (Cebrià 2007 ). This 2005 ; Cebrià et al. 2007 ). For example, silenc- neural infl uence could be in regulating different ing of the homologs of either netrin or a netrin events, such as cell proliferation, differentiation, receptor disrupts neural architecture in both and migration, as well as patterning. Some clas- intact and regenerating animals. The ventral sical studies have reported that regeneration is nerve cords regenerate as a completely disor- inhibited by removal of the nervous system in ganised meshwork of neural processes instead planarians such as Leptoplana , Dendrocoelum , of the normal two parallel cords of compacted and Bdellocephala (Child 1904a , b; Morgan axonal processes. Also, the regenerated neuro- 1905 ). In Dugesia dorotocephala, lateral pieces pil appears loosely packed after silencing of with no ventral nerve cord give rise to ‘head- any of these genes. In addition, these genes are bump’ regenerates: an abnormally large head required for the proper targeting of the visual with no differentiation of post-cephalic struc- axons (Cebrià and Newmark 2005 ). These tures (Sperry et al. 1973). However, in other spe- results exemplify the value of the fl atworm cies, lateral pieces with no ventral nerve cord CNS as a tool to investigate how the poor regen- can regenerate a head (Morgan 1898 , 1900 ). erative capabilities of mammals could be Also, it has been suggested that the brain is enhanced. required to induce eye regeneration through an Although the development of the nervous unknown factor of neuro-humoral nature (Wolff system has been described in different groups and Lender 1950; Lender 1955 ). Similarly, 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 57

putative neurohormones would control neoblast morphogenesis defects observed (Cebrià and migration towards the wound (Stéphan-Dubois Newmark 2007). Other studies have suggested and Lender 1956 ). that ventral nerve cords could provide some long- Other studies have analysed the role of neu- range cue to control polarity during regeneration rotransmitters in regeneration, mainly through (Oviedo et al. 2010 ). A recent report suggested indirect pharmacological approaches. For that the progression of head regeneration is example, dopamine and norepinephrine inhibi- blocked in the absence of a proper brain primor- tors reduce the rate of regeneration (Franquinet dium. The proposed model hypothesises that the 1979 ), somatostatin might inhibit proliferation initial brain primordium produces a signal that (Bautz and Schilt 1986 ), substance P appears induces neoblast proliferation and/or differentia- to stimulate cell proliferation and differentia- tion or migration to allow blastema growth tion in intact and regenerating planarians (Fraguas et al. 2014 ). (Baguñà et al. 1989b ), and neuropeptide F and FMRFamide stimulate pharyngeal regenera- tion (Sheiman et al. 2004). Serotonin also Eye Specifi cation and Regeneration accelerates regeneration, whereas anti-seroto- nergic drugs delay it (Franquinet et al. 1978 ). Due to its remarkable precision and complexity, More recently, Ladurner et al. (1997 ) proposed the eye has attracted the interest of many evolu- that the inability of Macrostomum to regener- tionary and developmental biologists. The animal ate when amputated postpharyngeally could be kingdom contains a great variety of eyes, with correlated with the lack of serotonergic neu- few representative basic types. The wide varia- rons in the posterior region. In fact, the pres- tion in morphology and organisation of the eye, ence of serotonergic neurons along the main as well as the differences in embryological devel- nerve cords has also been associated with asex- opment of morphologically similar eyes, sug- ual reproduction by paratomy in Microstomum gests that eyes have evolved independently a lineare . In these animals, the fi rst cells of the large number of times in animals (Salvini-Plawen new brain differentiate in close contact with and Mayr 1977 ). However, despite their morpho- the existing nerve cords, suggesting a neural logical diversity, eyes share many molecular sim- effect on neoblast proliferation and/or differ- ilarities, supporting the existence of conserved entiation (Reuter and Palmberg 1989; Reuter biochemical and developmental pathways and Gustafsson 1996 ). (Gehring and Ikeo 1999 ; Pichaud et al. 2001 ). At the molecular level, the silencing of neural The presence of a common eye morphogenetic genes such as nou - darake (Cebrià et al. 2002c ) programme supports a monophyletic origin of an and roboA (Cebrià and Newmark 2007 ) results in ancestral light sensory system. To reconcile such patterning and morphogenesis defects during disparate observations, it has been suggested that regeneration. Thus, after nou - darake RNAi, brain the common ancestor had a prototypical light tissues ectopically differentiate along the body, sensory organ. ‘Eyes’ then, as organs, evolved indicating that this gene is necessary to restrict independently, in some cases by convergence, the differentiation of brain tissues to the head such as the camera eyes of vertebrates and cepha- region. In contrast, after silencing roboA , the lopods or the various eyes found in different mol- newly differentiated cephalic ganglia appear dis- luscan taxa. connected from the truncated ventral nerve cords, A key question is how to defi ne the proto- which do not regenerate normally. Ectopic pha- typical eye (Gehring 2002 ). Planarian eyes, con- rynges and dorsal outgrowths develop around sisting of a combination of photoreceptor cell these regions of failed connection, suggesting and pigmented cell (Fig. 4.7 ; Hesse 1897 ), are that, in the absence of a proper connected CNS, considered good representatives of such a prim- some neurally derived signal could alter the itive visual system. This structure collects direc- behaviour of the neoblasts nearby and induce the tional information by using the pigment to block 58 F. Cebrià et al.

A1 A2 B1 B2

A3

B3

C

Fig. 4.7 Different types of simple eyes in Platyhelminthes. ceptor cell and one pigment cell ( B2 ). (B3 ) Lateral view Tricladida pigment cup ocelli: (A1 ) Two eye spots located of a polycelis planarian showing long rows of eyes along anteriorly, dorsal to the cephalic ganglia with the exten- the dorsoventral border. (C ) Lecithoepitheliata epidermal sions of bipolar photoreceptors connecting to the cephalic eyes represent one of the simplest eyes. They are pro- ganglia with a partial optic chiasma (A2 ). (A3 ) The eye- duced by a single cell and consist of ciliated sensors accu- cup is produced by the opposition of several pigment cells mulated in a pocket that is produced by cell membrane and, inside, the dendritic extremities of photoreceptor invagination and surrounded by a pigmented area. cb pho- cells have a rhabdomeric structure in which opsin accu- toreceptor cell bodies, p parenchyma, pg pigment cells, rh mulates. ( B1 ) Dorsolateral view of a planarian head with rhabdomeric structures. Scale bars in ( A1) 0.5 mm; (A2 , multiple eye spots formed by assembly of one photore- B1 , B3 , and C ) 0.25 mm; ( B2 and A3 ) 0.04 mm the light coming from specifi c directions. The a rhabdomeric structure, a regularly ordered majority of eyes among Platyhelminthes are microvilli assembly where opsin accumulates pigment cup ocelli situated close to the dorsal (Orii et al. 1998 ). The pigmented cells group to basal membrane, although special epidermal form an eyecup that surrounds the rhabdomeres eyes or rhabdomeric receptors without pig- (Fig. 4.7 ), removing the damaged photorecep- mented cells occur in some taxa. Such simple tive membranes and orientating the direction of ocelli are in close association with the brain and light (Tamamaki 1990 ; Rieger et al. 1991 ). In present a large variety of shapes and morpho- planarians, some muscle fi bres insert into the logical strategies. Since photoreceptors are eyecup, suggesting that the diameter of the eye bipolar neurons, their axons extend towards the opening can be adapted to varying light intensi- dorsomedian side of the cephalic ganglia ties and that ocelli can rotate towards incident and form a partial optic chiasma, which light direction. Finally, various light-refractive integrates photosensory inputs on both sides or lenticular structures have been described in (Fig. 4.7 ). Dendritic extremities generally have planarians in the area of the eyecup opening, for 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 59 example, cell extensions from the pigmented have been described in Stenostomum cell covering the eye opening (MacRae 1964 ; (Catenulida). These are single nerve cells with a Carpenter et al. 1974 ) or accumulation of mito- giant light-refracting mitochondrion (Ruppert chondria in pigment cell extensions in front of and Schreiner 1980), which, since these animals the photosensitive cells to form ‘mitochondrial are not phototactic, are thought to have some lenses’ (Sopott-Ehlers and Ehlers 2003 ). Such function related to photoperiodicity. Finally, in apparent sophistication of structures is only the larval stages of parasitic taxa, such as the enough to confer phototactic behaviour with free-swimming cercariae of trematodes, simple perception of variation in light intensity but not paired pigment cup ocelli contain a single pig- image formation (Rieger et al. 1991 ). mented cell and a photoreceptor rhabdomeric The second type of photoreceptive mem- cell located in front of the posterior pole of the brane amplifi cations are ciliary elaborations, cephalic ganglion. In addition, three unpig- which have only been found in some species of mented rhabdomeric photoreceptors in front of the orders Prolecithophora, Proseriata, and the pigmented cup ocelli have been described. Macrostomida. It is also found in the larvae of The pigmented ocelli serve to detect the direc- polyclads, whereas the adults have rhabdomeric tion of incoming light and to control the direc- structures. This duality in photoreceptive mem- tion of swimming in relation to the light brane has been found in few groups and may direction, whereas the unpigmented photorecep- indicate that this type of specialisation evolved tors serve as monitors for light intensity (Sopott- only once. Commonly, the rhabdomeric eyes Ehlers et al. 2001 ). In summary, the random consist of one or a few covering cells, most distribution of eye spots or ocelli with different often pigmented, and one or a few specialised complexities and cell number suggests that the bipolar nerve cells whose dendritic parts project various types of eyes evolved multiple times into the covering. These eyes, described in many independently within the Platyhelminthes. species among polyclads and macrostomids, The conserved eye genetic toolkit found in measure 30–50 μm in diameter. Triclads only Drosophila and confi rmed for different model have a single pair of eyes (between 50 and systems (Gehring 2002 ) is also present in 100 μm) containing between 35 and 70 photore- Platyhelminthes, at least in triclads (Saló et al. ceptive cells in a cup made of many cells 2002). Co-expression of planarian pax6 , six1 , (Fig. 4.7 ). In other species, the number of pig- and eya in eye cells appears to explain a lack of ment cup ocelli varies. Most have from one to requirement for the master control gene pax6 several pairs. The fi rst pair is dorsally located in (Pineda et al. 2000 ), which may be functionally front of the brain and the next pair occupies a substituted during planarian eye regeneration more lateral position. In some triclads, such as (Saló and Batistoni 2008 ). Table 4.3 summarises Polycelis, numerous small ocelli are scattered the most important genes expressed in the planar- over the head (Fig. 4.7 ) or from the front end of ian eye; an exhaustive eye transcriptome was the body and in rows along the lateral body mar- described in Lapan and Reddien 2012 . gin, even to the posterior pole of the body During eye regeneration, a stripe of migrating (Fig. 4.7 ). eye precursor cells appears once the cephalic The simplest platyhelminth eyes have been brain primordium is determined. These cells observed in the lecithoepitheliate Gnosonesima express the conserved transcription factors ovo , brattstroemi . It has a single epithelial eye cell six1 , and eya , which continue to be expressed in (Karling 1968 ) with ciliary pits inside the cell the differentiated eye cells. Ovo is a zinc fi nger membrane invagination which is surrounded by domain-containing protein with a conserved role pigmented vacuoles (Fig. 4.7 ). Proseriates also in determination of eye fate (Mackay et al. have cerebral rhabdomeric ocelli with intracel- 2006 ). The additional expression of o txA speci- lular modifi ed cilia in the same ocelli (Sopott- fi es the photoreceptor cells, whereas expression Ehlers 1982 ). Cerebral light-refracting bodies of sp6- 9 (a zinc fi nger-containing gene) and dlx 60 F. Cebrià et al.

Table 4.3 Summary of genes expressed in planarian eyes and phenotypes observed after their silencing by RNAi Planarian gene Homolog RNAi phenotype Reference Smed - otxA Crx / otd Absence of Lapan and Reddien (2011 ) photo receptor cells Dj / Gt / Smed - Pax6A Pax6 / eyeless / twin Not reported Callaerts et al. (1999 ), Rossi et al. Dj / Gt / Smed - Pax6B of eyeless ( 2001 ), Pineda et al. (2001 , 2002 ) Dj / Gt / Smedsix - 1 Six1 - 2 / Sine oculis No eyes Pineda et al. (2000 , 2001 ), Mannini et al. (2004 ) Dj / Smed - eya Eye absent 1 - 4 No eyes Mannini et al. (2004 ) Dj / Gt / Smed - opsin Opsin Loss of negative Sanchez-Alvarado and Newmark phototaxis (1999 ), Pineda et al. (2000 , 2001 ), Saló et al. (2002 ) Smednos Nanos Not reported Handberg-Thorsager and Saló (2007 ) Smed - netrin2 Netrin and netrin Defects in visual axon Cebrià and Newmark (2005 ) Smed - netR receptor targeting and abnormal phototaxis Dj b - arrestin Arrestin Not reported Agata et al. (1998 ), Nakazawa et al. (2003 ) DjTPH Tryptophan Not reported Nishimura et al. (2007a ) hydroxylase Smed - Ovo ovol1 - 3 / shavenbaby No eyes Lapan and Reddien (2012 ) zinc fi nger Smed - soxB Transcription Smaller eyes Lapan and Reddien (2012 ) Smed - foxQ2 factors Smed - klf Smed - meis Smed - SP6 - 9 Sp6 - 9 zinc fi nger Absence of eye Lapan and Reddien (2011 ) gene family progenitor cells Smed - dlx Distal - less Absence of eye Lapan and Reddien (2011 ) progenitor cells Djeye53 Unknown Impaired negative Inoue et al. (2004 ), Collins et al. Smed - eye53 - 1 prohormone gene phototaxis (2010 ) Smed - eye53 - 2 Smed - npp - 12 Neuropeptide Not reported Collins et al. (2010 ) precursor 12 Smed - mpl - 2 Myomodulin Not reported Collins et al. (2010 ) prohormone - like - 2 Smed - smad6 / 7 - 2 Smad Reduced eyes, Gonzalez-Sastre et al. (2012 ) phototaxis and lack of anterior photoreceptor cells Smed - BMP BMP Elongated eyes with Gonzalez-Sastre et al. (2012 ) expanded anterior photoreceptor cells Dj Dugesia japonica, Gt Girardia tigrina , Smed Schmidtea mediterranea

(a homeobox- containing gene) specifi es the pig- two bilaterally symmetric visual cell clusters in ment cell lineage. Finally, the transcription fac- the dorsal blastema. The pigment cells form an tors meis , klf , foxQ2 , and soxB regulate eye eyecup that surrounds the rhabdomeres. A series differentiation and morphogenesis (Lapan and of prohormone genes revealed the existence of Reddien 2011, 2012 ). The eye cells aggregate in three different subpopulations of photoreceptor 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 61 cells, located in the anterior, dorsal posterior, the re-establishment of polarity in the missing and posterior regions of the eyecup (Collins parts. According to this hypothesis, the appear- et al. 2010 ). The proper photoreceptor subpopu- ance of Janus heads would indicate that a mini- lation distribution inside the eyecup is regulated mal AP distance is required in a region of tissue by the BMP pathway (González-Sastre et al. in order to be interpreted as a gradient and to 2012 ). Interestingly, analysis of the expression specify polarity (Morgan 1905 ; Child 1911 ). The pattern of eye-specifi c transcription factors in study of planarian regeneration was one of the the planarian species Schmidtea polychroa mainstays of the ‘morphogenetic gradient’ and revealed extensive similarities between embry- ‘organising centre’ theories (Lewis et al. 1977 ; onic development and regeneration of the eye Meinhardt 1978), which continue to be central to (Martín-Durán et al. 2012 ). developmental biology today. In the last 10 years, with the sequencing of the genome of the planarian Schmidtea mediterranea SIGNALLING PATHWAYS and the introduction of RNAi techniques, the IN REGENERATION, GROWTH, molecular nature of the gradients that govern AND HOMEOSTASIS axial patterning has been elucidated (reviewed in Reddien 2011 ; Almuedo-Castillo et al. 2012 ). The Establishment of the Body Axis Thus, it was demonstrated that Wnts and BMPs are the morphogens that specify AP and DV axial Platyhelminthes show bilateral symmetry, with an identities, respectively, in adult planarians. The anterior-posterior (AP) axis (in which the anterior following section discusses the evidence for a pole corresponds to the brain region) and a very conserved role of these signalling pathways in short dorsoventral (DV) axis. Almost nothing is planarians and across metazoans. known about the molecular mechanisms that specify axial identity during embryogenesis in The Wnt/β-catenin Pathway Establishes any platyhelminth (Chapter 3 ). However, the the Planarian AP Axis molecular mechanisms of axial re-establishment The Wnt/β-catenin signalling pathway has an during regeneration have been studied in detail in evolutionarily conserved role in establishing the planarian species Schmidtea mediterranea polarity during embryonic development. It speci- and, to a lesser extent, Dugesia japonica . fi es the main axis in cnidarians (Wikramanayake Re-establishment of the body axis after ampu- et al. 2003 ) and echinoderms (Logan et al. 1999 ) tation has fascinated biologists since the nine- and the AP axis in most bilaterians (Holland teenth century, and planarians have remained a 2002; Croce and McClay 2006; Petersen and favourite model system in which to investigate it Reddien 2009 ). The binding of Wnts, the secreted (Morgan 1904 ). Classical amputation experi- elements of the pathway, to the receptors frizzled ments in different planarian species demonstrated and coreceptors LRP leads to the disruption of that planarians are able to regenerate the missing the β-catenin ‘degradation complex’, composed organs with the proper polarity in relation to the of axin, glycogen synthase kinase 3 (GSK3), pre-existing tissues after amputation. However, it casein kinase 1 (CKI), and adenomatous polypo- was observed that very narrow bipolar regenerat- sis coli (APC). Afterwards, β-catenin, the key ing pieces occasionally generated bi-headed pla- intracellular element of the pathway, accumulates narians (‘Janus heads’) (Morgan 1898 ). Although in the cytoplasm, enters the nucleus, and activates the molecules involved in this process were com- TCF transcription factors, which regulate the pletely unknown at that time, these experiments expression of multiple genes. led to the concept of ‘gradient activity’. It was Several elements of the Wnt/β-catenin signal- hypothesised that adult planarians should have an ling pathway that have been characterised in intrinsic gradient that, when broken through any Schmidtea mediterranea reveal a functional amputation, tends to be restored, thus inducing conservation of this pathway in the specifi cation 62 F. Cebrià et al. of the AP axial identities (reviewed in Almuedo- onstrating that adult animals require a continu- Castillo et al. 2012 ). Thus, silencing of Smed - ously active signalling pathway to maintain the βcatenin1 leads to an extreme anteriorised correct axial organisation (Iglesias et al. 2008 ). phenotype in which ‘radial-like hypercephalised’ The anteriorisation of the RNAi animals can be planarians develop large circular cephalic ganglia followed by the appearance of external morpho- together with several ectopic eyes all around the logical traits, such as the eyes, as well as by the planarian body (Fig. 4.8 ; Iglesias et al. 2008 ). In ectopic appearance of anterior (brain and eyes) amputated planarians that must regenerate the markers and the disappearance of posterior ones head and the tail, inhibition of Smed -β catenin1 (posterior Hox genes) (Gurley et al. 2008 ; Iglesias induces the regeneration of bi-headed planarians et al. 2008 ; Petersen and Reddien 2008 ). In the that gradually anteriorise completely. Interestingly, most extreme phenotypes, not only are posterior inhibition of Smed -β catenin1 in intact planarians regions absent, but the pharynx, which is located also leads to anteriorisation of the animals, dem- in the central region, also eventually disappears

ABC

D E

F

Fig. 4.8 Phenotypes generated after silencing Wnt or staining of the planarian central nervous system showing BMP signalling pathways. ( A ) In vivo images of planari- its complete anteriorisation after βcatenin1 silencing (B , ans in which βcatenin1 or BMP has been silenced by C ), its duplication after Smad1 inhibition (E ) (note the RNAi. Moderate silencing of βcatenin1 induces the appearance of an ectopic nervous system in the dorsal side regeneration of a head in the posterior part (bi-headed pla- of planarians), and its expansion along the medio-lateral narians) and higher inhibition leads to a fully anteriorised axis after the inhibition of Wnt5 ( F ) ( white line indicates phenotype (radial-like planarians; note the appearance of the midline), compared to controls ( D ). Anterior is to the ectopic eyes around the animal). Silencing of BMP leads left (A , C ) or the top left (B , D – F). Scale bars: (A ) 1 mm; to a ventralisation and duplication of the dorsoventral ( B – C ) 400 μm; ( D – F ) 200 μm border (Siamese-like planarians). (B – F ) Anti-synapsin 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 63

(Iglesias et al. 2008). The timing of appearance Interestingly, this situation resembles the one and disappearance of these molecular markers described for non-bilaterians, since the nested demonstrates that anteriorisation is a gradual and expression of Wnts in the aboral region and Wnt dose-dependent process. This is consistent with a inhibitors (dkk ) in the oral region defi nes the main model in which a morphogenetic gradient activity body axis of cnidarians (Fig. 4.9 ). Thus, β-catenin from a posterior organiser is responsible for pat- signalling appears as a general mechanism to terning the different identities along the AP axis defi ne axial polarity. (Adell et al. 2010 ). The involvement of the It is still not known whether the same signal- βcatenin- dependent Wnt signalling in this process ling networks pattern the embryonic body axis in is further supported by the fi nding that RNAi inhi- planarians. The core components of the canonical bition of elements of the β-catenin destruction Wnt pathway are expressed in planarian embryos complex ( Smed- APC and Smed - axin) leads to the at the stage where the yolk has been completely opposite phenotype, that is, posteriorisation of the ingested, that is, when the embryo differentiates animals (Gurley et al. 2008 ; Iglesias et al. 2011 ). the defi nitive structures that will replace the In agreement with the existence of a posterior embryonic ones (Martín-Durán et al. 2010 ; see organising region as a source of morphogens, four Chapter 3 ). The expression pattern resembles that of the nine Wnts that comprise the Wnt family in of adults, suggesting that the Wnt signalling planarians (Smed -Wnt1 /11 -1 / 11- 2/ 11- 5) are pathway also controls the establishment of defi n- expressed in nested domains in the most posterior itive axial identities during embryogenesis. part of planarians, and RNAi silencing of one of However, the molecular control of early polarity them (Smed -Wnt1 ) generates bi-headed planari- requires further attention. Smed - βcatenin1 is ans (Adell et al. 2009 ). Moreover, a Wnt inhibitor expressed earlier, mostly associated with the (Smed -notum ) is expressed in the most anterior development of the transient embryonic pharynx. tip, and RNAi for Smed -notum leads to the oppo- At this stage, genes involved in gastrulation site phenotype, that is, to ‘bi-tailed’ planarians ( snail ) and germ layer determination (foxA and (Petersen and Reddien 2011 ). Taken together, twist ) are specifi cally expressed in migrating these results strongly support a conserved role for blastomeres giving rise to the temporary phar- Wnt signalling in the specifi cation of the AP axis. ynx. Thus, β-catenin could have a primary role in Furthermore, the role seems also to be conserved endomesoderm specifi cation, which seems to be in different developmental contexts such as an ancestral role of the Wnt signalling, linked to embryonic development and adult regeneration. its function in the specifi cation of the main body

wnts

wnt11-5 V SFRPs wnt11-2 dkk Notum wnt1 D BMP BMP

Noggin BMP V D Wnt Wnt Wnt

AAPPAboral Oral Planarians Vertebrates Cnidarians

Fig. 4.9 Evolutionary conservation of the Wnt and BMP genesis, although in vertebrates a shift in the DV axis has signalling pathways in metazoan anterior-posterior and occurred. In prebilaterian animals such as cnidarians, Wnt dorsoventral axis establishment. In adult planarians, the signalling also patterns the main (oral-aboral) axis. The Wnt and the BMP signals pattern the anterior-posterior expression of Wnts (in blue ) and Wnt inhibitors (in red : (AP) and dorsoventral (DV) axes both during regeneration sFRP , notum and dkk) in opposite poles is shown in pla- and homeostasis. Patterning is similar to Xenopus embryo- narians and cnidarians 64 F. Cebrià et al. axis (Martín-Durán and Romero 2011 ). the shift in the positioning of the nervous sys- Moreover, expression analysis indicates that tem in chordates (De Robertis and Kuroda β-catenin could be a maternal gene (Martín- 2004 ). During Xenopus development, both dor- Durán et al. 2010), as found in several animals sal and ventral signalling centres serve as across the phylogenetic tree. sources of BMPs and their modulators (De From an evolutionary point of view, it is also Robertis and Kuroda 2004; De Robertis 2009 ). interesting to note that a duplication and func- BMPs and BMP antagonists are secreted ven- tional specialisation of β-catenin has been trally and ADMP (anti-dorsalising morphoge- reported in Schmidtea mediterranea. In most netic protein, a member of the BMP family) and animals, β-catenin is a bifunctional protein that other BMP antagonists (chordin and noggin) are has a transcriptional role when activated by the secreted by the dorsal centre. They together interaction of Wnts with their receptors and also confi gure a complex self-regulatory circuit that a role in cell adhesion, as a component of adher- fi nally restricts BMP signalling to the ventral ens junctions (Schneider et al. 2003). In planari- region of the embryo. ans, while Smed- βcatenin1 is exclusively Homologs of BMPs and their inhibitors have involved in transducing Wnt signalling, a second been identifi ed in Schmidtea mediterranea . β -catenin (Smed -β catenin2) is specifi cally Their expression patterns and functional charac- involved in cell adhesion (Chai et al. 2010 ). terisation by RNAi support the existence of an Duplications of β-catenin have also been equivalent self-regulatory circuit in adult pla- described in the nematode Caenorhabditis narians that patterns their DV axis during regen- elegans and in vertebrates, since plakoglobin is a eration and also during normal homeostasis duplication of β-catenin. However, phylogenetic (Molina et al. 2011a ). RNAi silencing of planar- analyses demonstrate that they originated from ian BMP , ADMP , and Smads , which are the independent duplication events (Korswagen intracellular effectors of BMP signalling, results et al. 2000). Interestingly, two β-catenins, which in animals in which the dorsal side is trans- probably share an evolutionary origin with the formed into a ventral one (Molina et al. 2007 ; planarian ones, are also found in Schistosoma 2011b; Orii and Watanabe 2007; Reddien et al. (Chai et al. 2010 ). Studies on representatives 2007 ; Gaviño and Reddien 2011 ). This ventrali- from other platyhelminth subclades will clarify sation is shown by the disappearance of dorsal the evolutionary relationship of these duplica- molecular markers, together with the ectopic tions and to which extent the separation of the appearance of ventral ones on the dorsal sides two roles could have implications for the plastic- and the differentiation of ectopic structures ity (regeneration and growth/degrowth) found in (Fig. 4.8 ; Molina et al. 2007 ; Reddien et al. the phylum. 2007 ). In severe phenotypes, there is a duplica- tion of the body margin and an almost complete The BMP Pathway Establishes ectopic CNS develops on the ventralised dorsal the Planarian DV Axis side, resulting in ‘Siamese twin-like’ planarians BMP proteins (bone morphogenetic proteins; (Molina et al. 2007 ). As in Xenopus , planarian Decapentaplegic [Dpp] in Drosophila ) comprise BMP and ADMP show complementary expres- a subfamily within the TGF beta superfamily of sion patterns along the dorsal and ventral mid- extracellular ligands, which have essential roles lines, respectively (Molina et al. 2007 , 2011b ; in key developmental processes in all metazo- Reddien et al. 2007 ; Gaviño and Reddien 2011 ), ans. Specifi cation of the DV axis is an evolu- and in accordance with the DV inversion that tionarily conserved role of the BMP pathway occurred in chordates. Furthermore, ADMP pro- (De Robertis and Kuroda 2004 ; Ashe and motes BMP expression and BMP inhibits ADMP Briscoe 2006 ), although in vertebrates BMP expression, in agreement with its role as a regu- signalling promotes ventral fates and in latory circuit that buffers against perturbations invertebrates it promotes dorsal fates, linked to of BMP signalling. Moreover, the planarian 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 65

BMP /ADMP circuit seems to be regulated by DV identities. In planarians, which are the only canonical antagonists of the noggin family, Platyhelminthes in which this issue has been since RNAi silencing of planarian noggin genes studied, two signals are known to be imple- ( noggin1 -2 ) produces a dorsalisation of the ani- mented, the Wnt5- and the Slit-secreted fac- mals (Molina et al. 2011b ). tors. Neither of these signalling inputs controls Very recently, an expanded noggin family the cellular transcriptional status, as with Wnt (up to ten members) was found to be present in or BMP signals in AP and DV patterning. planarians. Interestingly, its characterisation Instead, they control the assembly of the cel- allowed the discovery of a new type of noggin lular cytoskeleton in neural cells, acting as genes that were called noggin -like genes (nlg ), evolutionarily conserved axon-guidance cues which carry an insertion within the noggin (Ciani and Salinas 2005 ). domain (Molina et al. 2011b). Functional analy- The Wnt5 homologs of the planarian species sis of these genes in planarians shows that Schmidtea mediterranea and Girardia tigrina Smed -nlg -8 does not function as a canonical have been identifi ed (Smed - Wnt5 and Gt - Wnt5 ) BMP inhibitor. Instead, silencing induces ven- (Marsal et al. 2003 ; Adell et al. 2009 ). Smed - tralisation and enhances BMP RNAi pheno- Wnt5 is expressed from the most external part of types. Importantly, nlg homologs are found to the CNS to the lateral edges and RNAi leads to a be present in the genome of all metazoans, from lateral displacement of the CNS (Fig. 4.8 ; Adell sponges to chordates (Molina et al. 2011b ). The et al. 2009; Gurley et al. 2010). This phenotype unexpected activity of this family of novel regu- suggests that Smed - Wnt5 functions in restricting latory elements adds a new step in the complex the location of the CNS along the ML axis, pos- regulation of DV axis establishment that should sibly through its role as a repulsive cue for grow- be further investigated in different developmen- ing axons. This would be consistent with the role tal models. of Wnt5 in other models, such as Drosophila and To conclude, despite the shift in the posi- vertebrates, where Wnt5 acts through Slit or ROR tioning of the nervous system, the comparative receptors to control the decision of growing results available demonstrate conservation of axons to cross the midline or remain on the same BMP/ADMP signalling in establishment of the side (Yoshikawa et al. 2003 ; Ypsilanti et al. DV axis in protostomes and deuterostomes. 2010 ). Interestingly, the planarian homolog of They also support the notion of an ancestral slit ( Smed - slit), which is also a conserved signal role for the pathway in the specifi cation of neu- involved in repelling growing axons, shows a ral fates within ectodermal derivatives, since complementary expression pattern with respect neural territories are specifi ed in the region of to Smed - Wnt5, since it is expressed from the lowest BMP signalling (Reversade and De internal part of the CNS to the midline (Cebriá Robertis 2005 ). et al. 2007 ). Moreover, silencing of Smed - slit in planarians leads to collapse of the CNS at the Wnt5 and Slit Establish the Planarian midline (Cebrià et al. 2007 ), a phenotype that Medio-lateral Axis could be considered opposite to the one gener- Three axes are required to defi ne the positional ated after silencing Smed - Wnt5 , at least in rela- identity of a three-dimensional body. Thus, tion to the ML positioning of the nervous system. besides the AP and the DV axis, a third perpen- All this may indicate that Smed - Wnt5 and Smed - dicular axis from the midline to the edge of the slit could establish a signalling network that animal, the medio-lateral (M-L) axis, is defi ned restricts the positioning of the nervous tissues in bilaterians. This axis can be extended to the along the ML axis (Gurley et al. 2010 ; Almuedo- left-right axis, when left-right asymmetries are Castillo et al. 2011 ). Although a cooperative role observed. The molecular signals that pattern of these systems has not been described before, this axis seem not to be so general and broadly their role as repulsive cues for axons is conserved conserved as the ones that pattern the AP and among metazoans. However, two immediate 66 F. Cebrià et al. questions related to the role of Wnt5 and slit in it appears completely conserved with respect to planarians remain open, that is, the specifi c Schmidtea mediterranea (Yazawa et al. 2009 ). As molecular relationship between Smed - Wnt5 and mentioned above, RNAi for β - catenin in planar- Smed - slit in the control of CNS positioning and ian species with restricted regenerative abilities the nature of their receptors. From data in other ( Dendrocoelum lacteum , Procotyla fl uviatilis , models, a Derailed or ROR homolog could be the and Phagocata kawakatsui ) restores their ability Smed - Wnt5 receptors, and a Robo homolog to regenerate the head. This demonstrates the should be the Smed - slit receptor. In fact, a Smed - conservation of Wnt signalling in specifying AP RoboA gene has been identifi ed, but its RNAi axis identities. Moreover, it highlights the essen- phenotype suggests that it is not the Smed - slit tial link between the activation of the intercellu- receptor (Cebrià and Newmark 2007 ). lar signalling pathway and the ability to Altogether, the results found in planarians regenerate. These experiments highlight the rele- demonstrate that the Wnt and the BMP signalling vance of increasing the number of model species pathways are broadly used for the specifi cation of to advance our knowledge on animal develop- the AP and DV axis across metazoans and in all ment, regeneration, and evolution. developmental contexts, from embryonic devel- opment to the re-establishment of the axial iden- tities during adult regeneration and also for its Growth Control in Platyhelminthes maintenance during homeostasis. Remarkably, very recently it has been shown that in the acoel A long-standing question in developmental biol- Hofstenia miamia , which also shows whole-body ogy is how the size of an organ or a whole organism regeneration properties based on adult stem cells, is determined. Classical embryological studies the Wnt/β-catenin and BMP/ADMP signalling indicate that developing organs possess intrinsic pathways control the re-establishment of AP and information about their fi nal size. In the same man- DV axes, respectively, during regeneration. These ner, regenerating tissues also have mechanisms to results suggest that animals such as acoels and control their growth and reach the correct size. For planarians, separated by more than 550 million example, when a piece of a liver is removed, the years of independent evolution, would share sim- remaining part regenerates and stops growing ilar molecular regeneration mechanisms when the liver reaches the original size. In the same (Srivastava et al. 2014 ). manner, when fi rst-instar larvae or amputated ima- The conservation of these mechanisms during ginal discs are transplanted into adult fl ies, they the process of embryogenesis in planarians and grow or regenerate, respectively, to reach the same Platyhelminthes in general is a question that still fi nal size as unmanipulated ones. The molecular requires further studies, since it would allow for mechanisms that determine this ‘size checkpoint’ direct comparison of the patterning mechanisms in order to stop organ growth at the appropriate in embryonic and adult stages in the same species. point during development or regeneration are just In addition, understanding the mechanisms that beginning to be elucidated, mainly with the discov- control axial polarity during such a divergent ery of the intercellular communication mechanism cleavage process would help to understand this of the Hippo pathway. mode of development, which currently remains very poorly understood. The Hippo Signalling Pathway Regarding the evolutionary relationship of The Hippo cascade appears as an evolutionarily axial specifi cation within Platyhelminthes, the conserved growth suppressive signalling path- existence of Wnt and BMP elements has been way. It was initially discovered in the fruit fl y, reported in the planarian Dugesia japonica (Orii where mutations in components of the pathway et al. 1998 ) and a BMP homolog has also been resulted in dramatic overgrowth of tissues, and found in Schistosoma (Liu et al. 2013 ). Their role later on it was found to be conserved in mammals has only been characterised in D. japonica , where (reviewed in Johnson and Halder 2014 ). The 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 67 main function of the Hippo pathway is to nega- (Fig. 4.10 ). Hippo signalling is currently thought tively regulate the activity of Yorkie (YAP/TAZ to function as a sensor for the physical organisa- in mammals), a transcriptional co-activator that tion of cells in tissues (cell-cell contact, cell is the main downstream mediator of Hippo polarity, cell adhesion, etc.) to control cell prolif- (Fig. 4.10 ). Hippo kinase, the protein that gives eration and cell death. Thus, the pathway would the name to the pathway, is responsible for phos- coordinate the physical cues with the classic phorylating Warts and Mats, which, in turn, growth factor-mediated signalling pathways phosphorylate Yorkie. The phosphorylated form (Gumbiner and Kim 2014 ). Therefore, the Hippo of Yorkie is inactive, since it is retained in the pathway has a fundamental role not only in organ cytoplasm and ubiquitinated. When Hippo is growth control during embryonic development inactive, dephosphorylated Yorkie can enter the but also during regeneration and may be a hub in nucleus and transcriptionally control its targets, the control of stem cell and tumour growth. which generally promote cell proliferation and Hippo signalling has been most extensively inhibit cell death. Although several direct down- studied in mouse liver and heart, and in Drosophila stream target genes of the Hippo pathway have embryos, where YAP /Yorkie overexpression or been identifi ed, including cyclins, growth factors loss of lats /Warts kinase activities increases size and inhibitors of apoptosis, and genes involved in by increasing cell numbers (more proliferation and cell proliferation, cell survival, and stem cell less cell death). In general, the activation of the functions, the mechanism through which Yorkie Hippo pathway limits tissue growth by restricting drives tissue growth is not understood. proliferation and promoting apoptosis, and inhibi- Hippo signalling is similar to other signalling tion of the pathway (by activation of Yorkie ) is networks in that it depends on a cascade of phos- associated with stem cell expansion. Therefore, phorylation events. However, in contrast to other Hippo pathway inhibition is correlated with the pathways, it does not appear to have dedicated ability to regenerate missing tissues. Consistent extracellular signalling peptides and receptors with this, several studies have demonstrated that and is instead regulated by a network of upstream YAP is a crucial regulator of cardiomyocyte prolif- components, many of which are involved in eration and cardiac morphogenesis. Cardiac regulating cell adhesion and cell polarity deletion of YAP impedes neonatal heart

Fig. 4.10 The Hippo signalling pathway. Signals from the components of the cellular membrane modulate the phosphorylat- ing activity of Hippo and warts/lats kinases. When Hippo is active, it phosphorylates Yorkie/ YAP/TAZ inhibiting its entrance to the nucleus. When Hippo is inactive, Yorkie/YAP/TAZ can enter the nucleus and modulate the transcription of target genes, mainly associated with proliferation and apoptotic responses. First the Drosophila and then the mammalian name for each protein are indicated 68 F. Cebrià et al. regeneration, whereas its forced expression in the narians do not regenerate properly, but the cause adult heart stimulates cardiac regeneration after seems not to be reduced proliferation or increased myocardial infarction (Xin et al. 2013). In the apoptosis, as in other systems. Instead, it seems to same manner, during limb bud regeneration in the misregulate patterning, acting through the Wnt amphibian Xenopus laevis , YAP is upregulated, pathway. This result is not surprising, since cross- and inhibition causes limb bud regeneration talk with other intercellular signalling pathways defects (Hayashi et al. 2014 ). However, the rela- emerges as an intrinsic feature of the Hippo path- tionship between Hippo signalling, regeneration, way (Konsavage and Yochum 2013 ). Thus, and organ size is not absolute. The role of the although functional analysis of the cytoplasmic Hippo signalling pathway in the intestine, for Hippo kinases is lacking, Hippo signalling does example, remains unclear, since overexpression of not appear to directly regulate apoptotic and pro- YAP causes an enlargement of the stem cell com- liferative rates in planarians. partment but does not lead to an overall increase in An important consideration regarding the organ size (Li and Clevers 2013 ). differences found between Macrostomum and Schmidtea is the expression pattern of the Hippo Role of the Hippo Pathway elements. In Macrostomum , as in the mammalian in Platyhelminthes heart and liver, Yorkie is expressed in the stem cell Very recently the Hippo pathway has been char- compartment, that is, in neoblasts. In contrast, in acterised in two Platyhelminthes: Macrostomum planarians it is expressed in post-mitotic cells lignano (Macrostomida) and the planarian (Demircan and Berezikov 2013; Lin and Pearson Schmidtea mediterranea (Tricladida) (Demircan 2014 ). The situation in planarians strikingly and Berezikov 2013 ; Lin and Pearson 2014 ). resembles the one found in the mammalian and Knockdown of the Hippo pathway core genes in Drosophila gut system. In the mouse intestine, M. lignano during regeneration causes hyperpro- Yorkie /YAP is found in intestinal stem cells but liferation of neoblasts, which leads to the forma- also in enterocytes. Its overexpression fi rst acti- tion of outgrowths and to the disruption of vates proliferation and suppresses differentiation, allometric scaling, since regenerated parts appear in agreement with its generic growth- promoting bigger than the original ones (Demircan and effect, but it eventually causes the loss of Paneth Berezikov 2013 ). In contrast, Yorkie is essential cells (specialised epithelial cells of the small intes- for neoblast self-renewal, since Yorkie silencing tine located at the base of the crypts) and of the leads to the same ‘tissue regression’ phenotype intestinal crypts. Unexpectedly, inhibition causes observed after the inhibition of neoblast genes an increase in Paneth cells and intestinal stem cells (Reddien et al. 2005 ). Thus, in M. lignano , the (Li and Clevers 2013 ). These observations may be Hippo pathway appears to be functionally con- explained by the functional relationship between served in relation to Drosophila and mammals. the Hippo and the Wnt signalling pathways. Since, Although the homologs of the core Hippo path- in mouse, Paneth cells are an essential source of way elements have been identifi ed in Schmidtea growth factors, such as Wnts, they appear to be mediterranea , only the role of the transcriptional critical for the maintenance of the intestinal stem effector, Yorkie , has been reported (Lin and cell niche (Li and Clevers 2013 ). In planarians, Pearson 2014 ). Inhibition of Smed - Yorkie gener- like in the gut, inactivation of Yorkie also leads to ates a plethora of effects that are diffi cult to associ- activation of proliferation, and moreover, a rela- ate with a single function. However, it seems to be tionship between Hippo and Wnt signalling has directly involved in the maintenance and regenera- been demonstrated. Thus, it could be proposed tion of the planarian excretory system. A role for that the precise role of Yorkie depends on the cel- YAP , the mammalian Yorkie , in nephrogenesis lular composition of the tissues. In stem cell-based during mouse kidney development has also been systems, like planarians and the intestine, which reported (Reginensi et al. 2013 ). Regarding its role continuously undergo a stem cell-driven renewal in growth control, as expected, Yorkie RNAi pla- programme, Hippo and Yorkie elements would not 4 Regeneration and Growth as Modes of Adult Development: The Platyhelminthes as a Case Study 69 be directly required for stem cell proliferation but • Why is the ability to regenerate all missing tis- rather for the maintenance of the stem cell niche. sues restricted to certain Platyhelminthes? Why In other systems, such as the vertebrate liver and can only some triclads regenerate the head? heart, which are composed of mostly quiescent • What is the evolutionary origin of neoblasts? cells and only enter the cell cycle in extreme and • What is the percentage of true totipotent stem rare conditions, Yorkie would directly control cells in Platyhelminthes with high regenera- entry into the stem cell cycle. Supporting this tive capabilities? view, Macrostomum does not show such • How plastic are the progenitor cells to change broad regenerative capability as do planarians. their fate after amputation? Macrostomum normally reproduces sexually, and • Does the maintenance of adult stem cells and when forced, they are only able to regenerate the continuously active intercellular signalling posterior part but never the head region. Thus, play an equal role in regenerative capacity or Yorkie behaviour resembles that described in is one more relevant than the other? mammals and vertebrate heart tissues. • Is there a neoblast niche? The different functions found for Yorkie , for • Are similar molecular mechanisms involved in instance, in Macrostomum versus planarians, the control of embryogenesis and regeneration? highlight the existence of context-specifi c behav- iours that need to be addressed. The study of Acknowledgements We thank Bernhard Egger and Jim metazoan species with different stem cell natures Collins and Phil Newmark for providing the images of (regenerating versus non-regenerating species) Macrostomum lignano and Schistosoma mansoni, respec- tively, used in Fig. 4.4 . We thank Miquel Vila-Farré for will help to clarify the extent to which Yorkie has providing the specimens of Phagocata ullala and a conserved role specifi cally in the stem cell Camerata robusta used for the immunostainings shown in niche. The broader conserved role of Hippo sig- Fig. 4.4. We thank Maria Almuedo-Castillo for providing nalling in sensing cell contacts to control the fi nal planarian images in Fig. 4.3 . We thank Iain Patten for advice on the English. This work was supported by cell density and organ size suggests that its role is grant BFU2012-31701 (Ministerio de Economía y not restricted to the stem cell population but rather Competitividad, Spain) to F.C, grant BFU2008-01544 that it functions as a hub in coupling proliferative (Ministerio de Economía y Competitividad, Spain) to ES, and apoptotic responses. Based on limited data grant 2009SGR1018 (Agència de Gestió d’Ajuts Universitaris i de Recerca) to ES and FC, and grant from unicellular organisms, at least some ele- AIB2010DE-00402 (Ministerio de Economia y ments of the pathway appear to have been present Competitividad Accion Integrada) to ES. before the origin of multicellularity (Sebé-Pedrós et al. 2011 ; Artemenko and Devreotes 2013 ; Rock et al. 2013 ). Functional and comparative studies References in uni- and multicellular models should provide insights into the ancient role of those kinases and Adell T, Marsal M, Saló E (2008) Planarian GSK3s are the evolution of their linkage to extracellular sig- involved in neural regeneration. Dev Genes Evol nals in multicellular organisms. 218:105–106 Adell T, Saló E, Boutros M, Bartscherer K (2009) Smed - Evi / Wntless is required for b -catenin -dependent and -independent processes during planarian regeneration. 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Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger A. Wanninger (*) Department of Integrative Zoology , University of Vienna, Althanstrasse 14 , Vienna 1090 , Austria e-mail: [email protected] R. Neves Biozentrum – Molecular Zoology , University of Basel , Klingelbergstrasse 50 , Basel 4056 , Switzerland

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 79 DOI 10.1007/978-3-7091-1871-9_5, © Springer-Verlag Wien 2015 80 A. Wanninger and R. Neves

INTRODUCTION stage and colonizes a new host (Fig. 5.2G ). It has been interpreted as a derived trochophore-like Cycliophora is an acoelomate, monogeneric phy- larva, although characteristic features such as an lum that includes two described species, Symbion apical organ or a distinct prototroch are lacking pandora and Symbion americanus, but popula- (see Chapters 6 , 7 , and 9 ; Funch and Kristensen tion genetics and phylogeographic studies sug- 1995 ). gest that there are additional species awaiting In the original description of the fi rst cyclioph- formal description (Nedvěd 2004 ; Obst et al. oran species, Symbion pandora , Entoprocta and 2005 , 2006 ; Baker et al. 2007 ; Baker and Giribet Ectoprocta had been considered its closest extant 2007 ). Cycliophorans are found epizoically on relatives, mainly because all three phyla show setae of the mouthparts of lobsters, including clonal reproduction by budding (cf, Chapters 6 the Norway lobster, Nephrops norvegicus , the and 11; Funch and Kristensen 1995 ). This European lobster, Homarus gammarus , and the assumption has received support by some molec- American lobster, Homarus americanus . The ular phylogenetic studies that have united minute animals are characterized by a highly Ectoprocta, Entoprocta, and Cycliophora in a complex metagenetic life cycle that includes an monophyletic clade, the Polyzoa (Hejnol et al. asexual, polyp-like feeding stage that is fi rmly 2009; Paps et al. 2009 ). Other analyses, however, attached to the host lobster (Figs. 5.1 and 5.2A ). have proposed close rotifer/syndermate affi nities The feeding stage is also the easiest recogniz- of Cycliophora (Winnepenninckx et al. 1998 ; able life cycle stage. Its three-partite body is cov- Giribet et al. 2000). Since cycliophoran nucleo- ered by a thick cuticle and is subdivided into a tide sequence data have so far only been rarely buccal funnel that is surrounded by compound included in large-scale phylogenomic studies, the cilia and houses the mouth and esophagus, an defi nite position of Cycliophora within the bilat- oval-shaped trunk with all internal organs, and a erian tree remains contentious. short, acellular stalk with attachment disc (Funch and Kristensen 1995 ). It is the only stage in the cycliophoran life cycle known to feed, and its LIFE CYCLE STAGES buccal funnel and gut are recurrently regenerated AND DEVELOPMENT and replaced by the next generation of the inter- nally preformed corresponding structures. As a Cycliophorans exhibit one of the most complex consequence, older feeding stages have a series animal life cycles known to date (Fig. 5.1 ), which of cuticular wrinkles (scars) on their trunk is characterized by an alternation of asexual and (Fig. 5.2A ). The gut of the feeding stage is sexual reproductive cycles (metagenesis) (Funch U-shaped, with an anal opening located close to and Kristensen 1999 ). With a size of 300–400 μm the transition zone between the trunk and the the so-called feeding stage is the largest and most buccal funnel (Funch and Kristensen 1995 ). prominent stage in the life cycle. It has a simple Feeding stage individuals give rise to a number of muscular body plan that includes a myoepithelial other important stages in the cycliophoran life ring musculature surrounding the mouth, an anal cycle by internal clonal reproduction. These sphincter, and a few longitudinal muscles in the include the so-called Pandora and Prometheus buccal funnel and trunk (Fig. 5.3). Circular body larva and the sexual female (Fig. 5.2B, C, E ). wall muscles are absent, and the stalk and attach- These stages, as well as the male that develops ment disc are devoid of musculature altogether from an encysted Prometheus larva and the sexu- (Neves et al. 2009a , 2010a ). ally produced chordoid larva, are free-swimming The neuroanatomy of the feeding stage is poorly for at least some time and bear characteristic cili- known. Two anterior ganglia, one around the esoph- ary fi elds, usually in anteroventral and posterior agus and one in the buccal cavity, were described position (Fig. 5.2B–D ). The chordoid larva is (Funch and Kristensen 1997 ), but immunocyto- probably the longest-lived cycliophoran larval chemical studies using a suite of commercially 5 Cycliophora 81

MALE ESCAPES 17. FERTILIZATION?

16. 18. 19. ATTACHED PROMETHEUS 15. 20. LARVA ON FEEDING STAGE FEMALE 21. FEMALE SETTLES ESCAPES ON SAME HOST

14. 22.

PROMETHEUS LARVA ESCAPES 12. 23. SEXUAL 13. FEEDING STAGE WITH DEVELOPING FEMALE CHORDOID FEEDING LARVA STAGE WITH ESCAPES DEVELOPING PROMETHEUS 24. LARVA 9.

7. PANDORA LARVA ESCAPES 25. 10. 8.

REPLACEMENT OF BUCCAL FUNNEL

6. PANDORA 11. 1. FEEDING LARVA STAGE ASEXUAL SETTLES WITH ON SAME INNER BUD HOST

3. 2. CHORDOID 5. 4. LARVA SETTLES ON NEW HOST

Fig. 5.1 The hypothetical life cycle of Symbion pandora (From Neves et al. ( 2012 ), modifi ed after Obst and Funch (2003 ) ) 82 A. Wanninger and R. Neves

AB

C

DE

FG

Fig. 5.2 Representative life cycle stages of the cyclioph- pandora shown in A . ( D ) Dwarf male with penis ( double oran species Symbion pandora (A , D – G) and S. america- arrowhead) located inside a pouch-like structure; ventral nus (B , C ). Scanning electron (A – D , G ) and light view. (E ) Free-living female with large oocyte ( oo ). (F ) micrographs ( E , F). Anterior faces left in all aspects Chordoid cyst enclosing a chordoid larva. Note the chor- except for A , in which the buccal funnel faces up. (A ) doid organ (co ). (G ) Chordoid larva with paired dorsal

Feeding stage with two Prometheus larvae attached (apl 1– ciliated organs (arrows ). Abbreviations: ac anterior cili- 2 ). Note the wrinkle in the cuticle of the trunk (arrow- ated fi eld, ad attachment disc, bf buccal funnel, fc frontal head ). (B ) Pandora larva with anterior ciliated fi eld and ciliated fi eld, fe female exuvium, ls lateral sensorial organ, posterior ciliated tuft; ventral view. (C ) Prometheus larva pc posterior ciliated fi eld, pt posterior ciliated tuft, se seta with a posterior pair of toes (to ); lateral view. Note that the of the mouthparts of the host lobster, st stalk, tr trunk, vs toes are absent in the attached Prometheus larvae of S. ventral sensory organ (After Neves et al. (2010a , b ; 2012 ) ) 5 Cycliophora 83

A B

C D

Fig. 5.3 Feeding stage individual of Symbion sp. and shown in C . Abbreviations: bf buccal funnel, bfm settled Pandora larva with developing feeding stage musculature of the buccal funnel, cu cuticle, nbfm inside. Distal is up in all aspects. (A , C ) are light musculature of the new buccal funnel, gm musculature micrographs, (B , D ) show musculature visualized by of the gut, obfm musculature of the old buccal funnel, F-actin staining using fl uorochrome-coupled phalloidin rlm remnants of the musculature of the Pandora larva, and confocal microscopy. (A ) Feeding stage where the rm 1 ring muscle of the old buccal funnel, rm 2 ring new buccal funnel (nbf ) emerges and is about to replace muscle of the new buccal funnel, se seta of the mouth- the old buccal funnel (obf ). ( B ) Musculature of the parts of the host lobster, tlm longitudinal muscles of the feeding stage shown in A . ( C ) Settled Pandora larva trunk, tr trunk, double-headed arrows F-actin signal of with feeding stage developing inside. (D ) Musculature degenerating musculature of the Pandora larva of the feeding stage developing inside the Pandora larva 84 A. Wanninger and R. Neves available antibodies directed against a number of maternal feeding individual (Funch and neural markers (e.g., serotonin) only showed scat- Kristensen 1995 ). The cerebral ganglion of the tered and weak signal in the buccal funnel of Pandora larva and all other free-swimming stages Symbion pandora and did not recognize any gangli- degenerates during settlement. onic structures. Transmission electron microscopy In order to enter the sexual reproduction phase, analyses identifi ed nervous tissue in the buccal fun- feeding stages may either develop, one at a time, a nel, but whether this is ganglionic in nature could female or a Prometheus larva, both internally not be clarifi ed (Neves et al. 2010b ). (Figs. 5.1 and 5.2C, E). While still within the feed- According to the data available, all other life ing stage, the female develops one single oocyte cycle stages are at least for some time (Fig. 5.2E). The Prometheus larva settles on the free- swimming (Wanninger 2005 ; Neves et al. trunk of a feeding individual after release 2010b , c ). They have varying sets of circular, lon- (Fig. 5.2A ), encysts, and generates internally one gitudinal, dorsoventral, and other muscles that to three mature dwarf males with a cuticular penile support the body (Wanninger 2005; Neves et al. structure (Fig. 5.2D ; Neves et al. 2010c ). Electron 2009a , 2010a ). In these stages, the nervous sys- microscopy studies showed that younger males tem comprises a relatively large, bilobed cerebral are larger and possess more nucleated somatic ganglion and two ventral longitudinal nerves. cells (aproximately 200) than mature individuals, These elements contain serotonin (Wanninger who only have around 50 nucleated cells and 2005 ; Neves et al. 2010b , d ). The chordoid larva nuclei-free muscles and epidermis (Neves and alone has four ventral neurite bundles which fuse Reichert 2015 ). With 30–40 μm in length these in the posterior region (Wanninger 2005 ). In animals range among the smallest free-living, sex- addition to the serotonergic signal, these nerves ually mature metazoans. Despite their small size, showed synapsin as well as FMRFamide immu- cycliophoran dwarf males have a distinct body noreactivity, with the latter being confi ned to the wall musculature, a brain that occupies the ante- two outer neurites (Neves et al. 2010d ). rior third of the body, as well as two ventral nerve Posterolateral sensory organs (“lateral ciliated cords (Obst and Funch 2003 ; Neves et al. 2010b ). pits”) and a dorsoanterior ciliated sensory organ Feeding stages with encysted Prometheus lar- are present in the chordoid larva (Funch 1996 ), vae often bear females inside the trunk. The male but the latter most likely does not correspond to probably hatches from the cyst, and the mature the apical organ of other lophotrochozoan larvae female is released from the feeding stage. (Neves et al. 2010d). Distinct neural subsets Fertilization of the oocyte is internal, but whether underlying (and potentially innervating) the cili- it occurs while the female still resides within the ary fi elds, as present in most lophotrochozoan feeding stage (through the wall of the male cyst, larvae, were not found. Protonephridia are only the cuticle of the feeding stage, and the body wall known from the chordoid larva. The name-giving of the female!) or during the free-living stage “chordoid organ” of this larva is composed of a remains speculative. In any case, the female with series of muscular subunits that form a bow-like the fertilized oocyte settles with the anterior pole structure that extends from the posterior pole in on the mouthparts of the same host, degenerates, ventral direction from where it continues into the and forms the so-called chordoid cyst (Fig. 5.2F ). anterior region (Wanninger 2005 ). Light sensory Therein, the sexually produced chordoid larva organs are absent in all stages. develops from the fertilized oocyte. As part of the asexual life cycle, the Pandora Isolated observations found a female with an larva emerges from the feeding stage and, after a uncleaved oocyte inside a feeding stage and sev- supposedly brief free-swimming stage, settles on eral free-swimming oocyte-bearing females. This setae of the mouth parts of the same host lobster. may indicate that fertilization takes place after It attaches with the apical region and gives rise to release of the female from the feeding stage. a new feeding stage, whose anlagen are already Indeed, the only account on embryogenesis stems formed in Pandora larvae liberated from the from settled females (Neves et al. 2012 ). Cleavage 5 Cycliophora 85

AB

C

Fig. 5.4 Symbion pandora . ( A ) Light micrograph of a specimen shown in ( B). Abbreviations: ac auxiliary cells, feeding stage with a developing female inside. Note the ad attachment disc, dbf degenerated buccal funnel, sd large oocyte ( oo ). (B ) Light micrograph of a settled female settlement disc, se seta of the mouthparts of the host lob- with an embryo (em ) at the eight cell stage. Anterior faces ster, st stalk, tr trunk (After Neves et al. (2010b , 2012 ) ) down. ( C ) Line drawing of a settled female based on the appears to be holoblastic with four micromeres Crucial events such as fertilization, settlement of and four macromeres at the eight cell stage, but the chordoid larva, and details of the embryology blastomere arrangement bears no similarities to a have never been directly observed, and details on spiral cleavage pattern (Fig. 5.4B, C ). Polar bod- the development of the various asexually pro- ies were not observed. Hatched chordoid larvae duced stages are unknown. Needless to say that probably seek a new lobster specimen, settle, no data on gene expression patterns are currently encyst, and metamorphose into new feeding indi- available, leaving the fi eld wide open for future viduals. Accordingly, the chordoid larva is studies into literally all directions of cycliophoran regarded as the cycliophoran dispersal stage. developmental biology. While this reads like a well-founded recon- A recent observation found cycliophoran indi- struction of the cycliophoran life cycle, most of its viduals on copepods that live on the mouthparts of dynamics are only inferred from studies on the the European lobster, casting doubt on the pro- morphology of the various stages and their distri- posed cycliophoran host specifi city being confi ned bution on the host animals (Obst and Funch 2006 ). to lobster crustaceans (Neves et al. 2014 ). This 86 A. Wanninger and R. Neves

fi nding may be of utmost importance for the under- invertebrates, vol 13, Lophophorates, Entoprocta and stating of the cycliophoran life cycle and questions Cycliophora. Wiley-Liss, New York, pp 409–474 Funch P, Kristensen RM (1999) Cycliophora. In: Knobil concerned with the dispersal and colonization of E, Neill JD (eds) Encyclopaedia of reproduction, vol new host lobsters, illustrating that we have only 1. Academic, New York, pp 800–808 begun to understand the basic mechanisms that Giribet G, Distel DL, Polz M, Sterrer W, Wheeler WC underlie the biology of this enigmatic phylum. (2000) Triploblastic relationships with emphasis on the acoelomates and the position of Gnathostomulida, Cycliophora, Plathelminthes, and Chaetognatha: a combined approach of 18S rDNA sequences and mor- OPEN QUESTIONS phology. Syst Biol 49:539–562 Hejnol A, Obst M, Stamatakis A, Ott M, Rouse GW, Edgecombe GD, Martinez P, Baguñà J, Bailly X, • Virtually all aspects of cycliophoran develop- Jondelius U, Wiens M, Müller WEG, Seaver E, ment, including fertilization, embryology, Wheeler WC, Martindale MQ, Giribet G, Dunn CW cleavage, organogenesis, and gene expression. (2009) Assessing the root of bilaterian animals with • Emergence of the asexually produced life scalable phylogenomic methods. Proc R Soc B 276:4261–4270 cycle stages after settlement and encystation, Nedvěd O (2004) Occurrence of the phylum Cycliophora in particular: Does the chordoid larva indeed in the Mediterranean. Mar Ecol Prog Ser metamorphose into a feeding stage? 277:297–299 • Is the chordoid larva a modifi ed trochophore? Neves RC, Kristensen RM, Wanninger A (2009a) Three- dimensional reconstruction of the musculature of vari- • What induces sexual reproduction in the ous life cycle stages of the cycliophoran Symbion Symbion life cycle? americanus . J Morphol 270:257–270 Neves RC, Sørensen KJK, Kristensen RM, Wanninger A (2009b) Cycliophoran dwarf males break the rule: Acknowledgements AW thanks the Faculty of Life high complexity with low cell-numbers. Biol Bull Sciences, University of Vienna, for generous support 217:2–5 while establishing his group during his fi rst years in Neves RC, Cunha MR, Kristensen RM, Wanninger A Vienna. The Danish Research Agency (FNU) and the (2010a) Comparative myoanatomy of cycliophoran Carlsberg Foundation as well as the European Commission life cycle stages. J Morphol 271:596–611 are thanked for the support of his research during the Neves RC, Kristensen RM, Wanninger A (2010b) Serotonin Copenhagen years. Marion Hüffel provided invaluable immunoreactivity in the nervous system of the Pandora support during this entire project and beyond. AW partic- larva, the Prometheus larva and the dwarf male of ularly thanks his coauthor, Ricardo Neves, for utmost pro- Symbion americanus (Cycliophora). Zool Anz 249:1–12 ductive years of cooperation into cycliophoran research. Neves RC, Cunha MR, Funch P, Wanninger A, Kristensen RM (2010c) External morphology of the cycliophoran dwarf male: a comparative study of Symbion pandora and S. americanus . Helgol Mar Res 64:257–262 References Neves RC, Cunha MR, Kristensen RM, Wanninger A (2010d) Expression of synapsin and co-localization Baker JM, Giribet G (2007) A molecular phylogenetic with serotonin and RFamide-like immunoreactivity in approach to the phylum Cycliophora provides further the nervous system of the chordoid larva of Symbion evidence for cryptic speciation in Symbion america- pandora (Cycliophora). Invertebr Biol 129:17–26 nus . Zool Scr 36:353–359 Neves RC, Kristensen RM, Funch P (2012) Ultrastructure Baker JM, Funch P, Giribet G (2007) Cryptic speciation in and morphology of the cycliophoran female. J the recently discovered American cycliophoran Morphol 273:850–869 Symbion americanus; genetic structure and population Neves RC, Bailly X, Reichert R (2014) Are copepods sec- expansion. Mar Biol 151:2183–2193 ondary hosts of Cycliophora? Org Divers Evol Funch P (1996) The chordoid larva of Symbion pandora 14:363–367 (Cycliophora) is a modifi ed trochophore. J Morphol Neves RC, Reichert H (2015) Microanatomy and develop- 230:231–263 ment of the dwarf male of Symbion pandora (Phylum Funch P, Kristensen RM (1995) Cycliophora is a new Cycliophora): new insights from ultrastructural inves- phylum with affi nities to Entoprocta and Ectoprocta. tigation based on serial section electron microscopy. Nature 378:711–714 PLoS ONE 10: e0122364 Funch P, Kristensen RM (1997) Cycliophora. In: Harrison Obst M, Funch P (2003) Dwarf male of Symbion pandora FW, Woollacott RM (eds) Microscopic anatomy of (Cycliophora). J Morphol 255:261–278 5 Cycliophora 87

Obst M, Funch P, Kristensen RM (2006) A new species of Paps J, Baguña J, Riutort M (2009) Lophotrochozoan Cycliophora from the mouthparts of the American internal phylogeny: new insights from an up-to-date lobster, Homarus americanus (Nephropidae, Decap- analysis of nuclear ribosomal genes. Proc R Soc B oda). Org Div Evol 6:83–97 276:1245–1254 Obst M, Funch P (2006) The microhabitat of Symbion Wanninger A (2005) Immunocytochemistry of the pandora (Cycliophora) on the mouthparts of its host nervous system and the musculature of the chordoid Nephrops norvegicus (Decapoda: Nephropidae). Mar larva of Symbion pandora (Cycliophora). J Morphol Biol 148:945–951 265:237–243 Obst M, Funch P, Giribet G (2005) Hidden diversity and Winnepenninckx B, Backeljau T, Kristensen RM (1998) host specifi city in cycliophorans: a phylogeographic Relations of the new phylum Cycliophora. Nature analysis along the North Atlantic and Mediterranean 393:636–638 Sea. Mol Ecol 14:4427–4440 E n t o p r o c t a 6 Andreas Wanninger

Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger. A. Wanninger Department of Integrative Zoology , University of Vienna, Althanstrasse 14 , Vienna 1090 , Austria e-mail: [email protected]

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 89 DOI 10.1007/978-3-7091-1871-9_6, © Springer-Verlag Wien 2015 90 A. Wanninger

INTRODUCTION side of the atrium, from which nerves emanate and project into the stalk and tentacles (Fuchs Entoprocts or kamptozoans are solitary or colo- et al. 2006 ; Schwaha et al. 2010 ). Intermediate nial aquatic fi lter feeders with a characteristic tentacle ganglia, as found in ectoprocts, are ciliated tentacle apparatus. They are acoelomate, lacking. Ciliated sensory organs are often found but non-mesothelially lined lacunae form along the stalk and calyx (Fuchs et al. 2006 ). schizocoelic spaces that pervade the animals. These may serve as geosensory organs, but About 200 recent species have been described, detailed studies are lacking. Light sensory with a total body size roughly ranging between organs have so far not been found in adult ento- 100 μm and several millimeters. The phylum is procts. The U-shaped digestive tract lies within largely confi ned to marine (including brackish) the calyx and consists of a short esophagus and habitats, with only two freshwater species hindgut and a relatively prominent stomach. known to date. Usually one pair of protonephridia is found in The entoproct body consists of a slender, adults as well as larvae (Nielsen and Jespersen muscular stalk with terminal secretory cells that 1997 ); a complex arrangement of numerous pro- allow for reversible attachment to the substrate. tonephridia, probably serving osmoregulatory An adhesive disc is often present in solitary spe- functions, has been found in the two freshwater cies. The stalk is continuous with the calyx, species, Urnatella gracilis and Loxosomatoides which constitutes the major body region sirindhornae (Emschermann 1965 ; Schwaha (Fig. 6.1 ). It terminates with the atrium, which et al. 2010 ). For further details on the morphol- houses the slit-like mouth opening and the ele- ogy of adult entoprocts, see Emschermann vated anus sitting on the anal cone. Accordingly, (1982 ), Nielsen and Jespersen (1997 ), Fuchs the “upper” side of the animal marks its morpho- et al. (2006 ), and Nielsen (2012 ). logical ventral side. The entire atrial region is Morphological and molecular analyses agree surrounded by the tentacle crown (Fig. 6.1 ). The that colonial and solitary entoprocts each form monolayered epidermis is covered by a cuticle monophyletic assemblages, thus subdividing which may be reduced or entirely lacking on the Entoprota into the Coloniales (comprising the inner side of the tentacles and the atrium; these Barentsiidae, Pedicellinidae, and Loxocalypodidae) areas have been proposed to be the major sites of and the Solitaria (with Loxosomatidae as the sole gas exchange (specifi c respiratory organs are subclade) (Emschermann 1972 ; Fuchs et al. 2010 ). generally absent in entoprocts). Within the calyx It is generally assumed that the (solitary) loxoso- reside the major inner organs including nerves, matids have largely retained major basal features of muscles, gonads, brood pouches, protone- Entoprocta, including a lecithotrophic, creeping- phridia, and digestive tract (Nielsen and type larva (see below). Jespersen 1997 ). The stalk (including attach- Traditionally, Entoprocta has been thought to ment disc, if present) is highly muscular and cluster with the Ectoprocta to form the phylum functions as an internal skeleton, thereby allow- Bryozoa (Nielsen 1971), but this view is chal- ing for fl exible body movements (Wanninger lenged by differences in adult morphology (e.g., 2004 ). The musculature of the calyx is only coelom in ectoprocts versus acoelomate condi- weakly developed. Its most prominent feature is tion in entoprocts) and developmental characters a ring muscle at the base of the tentacles that such as mode of cleavage (spiral in entoprocts allows for closure of the atrial region and protec- versus radial in ectoprocts; see Chapter 11 ). tion of the tentacles upon contraction. A three- Instead, similarities in the non-coelomic, lacuna- layered body wall musculature comprising ring, like circulatory system and in a number of larval diagonal, and longitudinal muscles, typical for features including the presence of a ciliated foot, many (vermiform) lophotrochozoans, is lacking. the morphology of the apical organ, and a The nervous system is simple and includes a tetraneurous nervous system suggest entoproct- dumbbell-shaped cerebral ganglion at the oral molluscan monophyly (Sinusoida, Lacunifera, or 6 Entoprocta 91

Fig. 6.1 Gross morphol- ogy of a solitary entoproct, the loxosomatid Loxosoma nielseni , with major body plan features indicated. Scanning electron micrograph, the total size of the animal from foot plate to anus is 175 μm

Tetraneuralia concept; Chapter 7 ) (Bartolomaeus EARLY DEVELOPMENT 1993 ; Ax 1999 ; Wanninger 2009 ). Some molecu- lar phylogenies recovered a superclade Polyzoa, Many solitary entoprocts are protandric her- with Entoprocta and Cycliophora monophyletic maphrodites. From the colonial genus Barentsia , and the sister to Ectoprocta (e.g., Hejnol et al. species are known that form either male or 2009), but this was heavily refuted by others, female colonies only or have male and female who suggested that this clade resulted from a sys- zooids in a single colony, while other species tematic bias (Nesnidal et al. 2013 ). Additional have hermaphroditic zooids, thus indicating the alternative scenarios have been proposed, includ- high plasticity of entoproct reproductive modes ing one that found Entoprocta in a basal position (Wasson 1997; see Nielsen 2012). The male of a lophotrochozoan subclade that also includes gametes are released into the water column. the mollusks, annelids, nemerteans, brachiopods, Fertilization is internal and the embryos are and phoronids, but notably not the ectoprocts maintained in the calyx. Matrotrophy has been (Dunn et al. 2008 ). The ongoing vivid discus- described for some species (Harmer 1885 ; sions demonstrate that the entoprocts have not Nickerson 1901 ). yet found their fi nal resting place within the Accounts on entoproct early embryology lophotrochozoan tree of life. and cleavage are scarce and not very detailed. For a long time, the main information had been 92 A. Wanninger on two colonial representatives, Pedicellina spiralian cross-like patterns resembling those of (Fig. 6.2 ; Hatschek 1877; Marcus 1939 ) and annelids and mollusks at around 43 cells in Barentsia (Malakhov 1990 ), supplemented by a Loxosomella (Fig. 6.5; Merkel et al. 2012 ). classical work on solitary loxosomatids, mostly Gastrulation is preceded by fl attening of the Loxosomella leptoclini (Harmer 1885 ). Recently, embryo, occurs when the embryo has slightly new data have become available on a yet unde- more than 100 cells, and eventually results in a scribed Loxosomella species employing fl uores- coeloblastula (Merkel et al. 2012 ). This is in cence staining of nuclei and 3D reconstruction of accordance with the data provided by Marcus embryogenesis (Figs. 6.3 and 6.4 ; Merkel et al. ( 1939) on Pedicillina cernua, who found the 2012 ). Most authors noted a holoblastic, more or embryos to gastrulate at around 120 cells. less equal, spiral cleavage pattern, comparable For Loxosomella leptoclini , Harmer (1885 ) to that of other Spiralia. For Pedicillina , syn- mentioned subsequent closure of the blastopore chronous as well as asynchronous cell divisions and the formation of two lateral mesodermal have been reported during early cleavage cycles bands that give rise to the future mesoderm, as (Hatschek 1877 ; Marcus 1939 ). well as de novo generation of the mouth and In Loxosomella sp., cleavage is highly asyn- emergence of the anus in the region of the former chronous. An apical rosette was found in blastopore, but these data need careful reinvesti- Pedicillina and Loxosomella, together with two gation using modern methods.

1a111 1d111

1a112 1d121 1a121 1d112

1a22 1a122 22 1d122 1d21 1d 1c21

12 2d 2d21 11 12 2a21 11 2a 2a 2a22 2d 2d22

3d

4d 3a 4a 3c

5c 5d 5D

5A 5C

Fig. 6.2 Cell lineage of the colonial entoproct Pedicillina blue. The mesoderm founder cell (mesendoblast) 4d is in cernua. Cells of the apical rosette are in yellow , red and blastomeres that give rise to endodermal cells are trochoblasts are in dark blue , and the remaining in magenta . Polar bodies are depicted in gray . (Redrawn blastomeres that contribute to the ectoderm are in light and modifi ed after Marcus (1939 ) ) 6 Entoprocta 93

AB C

D EF

G HI

JKL

MNO

Fig. 6.3 Early entoproct embryology of Loxosomella sp. color in the 3D reconstructions. Polar bodies are in gray in showing spiral cleavage. Images in the left column are the reconstructions and indicated by arrowheads in the confocal micrographs of nuclei stainings; middle and confocal images. Note the almost similar size of macro- right columns are 3D reconstructions. Middle column , meres (Q ) and micromeres (q ). Scale bars represent 10 μm animal view; right column, lateral view. Cells resulting (From Merkel et al. (2012 ) ) from a distinct cleavage cycle are labeled with the same 94 A. Wanninger

ABC

DEF

GHI

JKL

Fig. 6.4 Confocal micrographs and 3D reconstructions animal view. ( C , I , L ) Lateral view. Red : “macromere” of blastula and gastrula stages of Loxosomella sp. “Apical quartet (i.e., vegetal) cell nuclei; green , (derivatives of) cross patterns” are indicated by white and red lines in D , apical rosette cell nuclei; purple , other cell nuclei. (A – C ) G . Apical rosette cell nuclei are marked with asterisks in 36-cell stage. ( D – F) 43-cell stage. Cells surrounding the D , E , G , J and vegetal “macromere” quartet cells by dou- apical rosette ( asterisks ) show both a molluscan- and an ble arrowheads in F . Scale bars represent 10 μm. ( A , D , annelid-like cross pattern (D ). (G – I ) 51-cell stage. ( J – L ) G ) Nucleic acid staining (blue ). ( E , F ) Nucleic acid (blue ) Gastrula stage (107 cells). Derivatives of apical rosette and F-actin staining ( red ). ( J) Nucleic acid staining shown cells lie in a lower plane than the surrounding cells ( J ). as depth-coded confocal projection. ( B , C , H , I , K , L ) 3D ( K , L ) Gastrulation. Vegetal cells (red ), blastocoel (arrow ) reconstructions. ( A , F ) Vegetal view. (B , D , E , G , H , J , K ) (From Merkel et al. ( 2012 ) ) 6 Entoprocta 95

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GH I

Fig. 6.5 Spiralian apical cross cell patterns based on stage of the nemertean Carinoma tremaphoros ; after several authors. Dark gray , apical rosette cells; middle Maslakova et al. ( 2004 ). (F ) Approximately 64-cell stage of gray, periphere rosette/“annelid cross” cells; light gray , the polyclad fl atworm Maritigrella crozieri; after Rawlinson “molluscan cross” cells. (A ) 43-cell stage of Loxosomella (2010 ). ( G) 64-cell stage of the echiurid Urechis caupo ; sp. ( B ) Approximately 64-cell stage of the aplacophoran after Newby (1932 ). ( H ) 48-cell stage of the sipunculan mollusk Epimenia sp.; after Baba (1951 ). ( C ) Golfi ngia vulgaris (= Phascolosoma vulgare ); after Approximately 62-cell stage of the polyplacophoran mol- Gerould (1903 ). (I ) Approximately 58-cell stage of the lusk Stenoplax heathiana (= Ischnochiton magdalenensis ); polychaete Nereis sp.; after Wilson (1892 ) (From Merkel after Heath (1898 ). (D ) 58-cell stage of the gastropod mol- et al. (2012 )) lusk Patella caerulea; after Wilson ( 1904). ( E ) 64-cell

Following cell fates is particularly diffi cult in for Pedicellina , where progenies of the 1a-d lin- entoprocts due to the brooded embryos and eages form the apical rosette and the trochoblasts. because the micromeres and macromeres are of Second quartet micromeres, as, e.g., in mollusks, almost the same size (Merkel et al. 2012 ). The were not identifi ed to contribute to the prototroch only data on entoproct cell lineage are available (Fig. 6.2; Marcus 1939). As in other spiralians (in 96 A. Wanninger particular mollusks; see Chapter 7 : Table 1), the mother animal as (almost?) metamorphic com- second and third quartet micromeres contribute petent individuals. Although they do have a to ectodermal domains, the 4a–c and 5a–d lin- mouth and gut, they do not seem to feed. They eages to the endoderm, and the mesendoblast 4d immediately start a semi-benthic life during was found to constitute the endomesoderm which they creep over the substrate with their mother cell (Fig. 6.2 ; Marcus 1939 ), but more distinct foot. This behavior is only briefl y detailed data using modern methods are highly interrupted by short excursions into the water desired to further assess entoproct cell genealogy. column. As in the swimming-type larvae, the Data on organogenesis and the precise contribu- foot is situated posttrochally but, in contrast to tion of individual blastomeres to larval organ sys- the former, is signifi cantly pronounced (Figs. 6.6 tems are virtually unknown. and 6.7C ). In the creeping-type larva, both swimming and creeping is directed with the frontal organ forward (not with the apical organ LATE DEVELOPMENT as in other lophotrochozoans) and thus refl ects a 90° shift of the (functional) anterior-posterior Larval Morphology axis of the animal compared to that of all other lophotrochozoan larvae. Attachment prior to Entoprocts exhibit a wide variety of larvae metamorphosis is either with the frontal organ which can be subdivided into two major catego- or the foot facing the substrate, similar to the ries, a swimming- and a creeping-type larva, swimming-type larvae. with the latter most likely constituting the basal The entoproct creeping-type larva exhibits a type (Figs. 6.6 and 6.7 ; Nielsen 1971 ; Fuchs number of morphological features that have et al. 2010 ). The swimming-type larva has a pre- been homologized with corresponding struc- dominant episphere. The hyposphere is often tures of mollusks and thus entoproct-mollusk reduced and obscured by the compound cilia of monophyly has been proposed (see above). the prototroch (Fig. 6.7A ), although a short, Shared characters include the ciliated creeping ciliated protrusion, the larval foot, is at foot with distinct foot glands and compound least rudimentarily present in most species. cirri in the anterior region of the foot, which Swimming-type larvae often have a long plank- have been homologized with those of adult neo- tonic and planktotrophic phase which can last meniomorph mollusks (Fig. 6.6 ; Haszprunar for several weeks. The apical organ is simple and Wanninger 2008 ). Both the creeping-type and comprises a ciliated tuft and three or four larva and polyplacophoran larvae have an apical serotonin-positive fl ask-shaped cells (Fig. 6.7D, organ with a complex arrangement of eight to G; Fuchs and Wanninger 2008 ). From there, a ten serotonin-containing fl ask-shaped (ampul- paired nerve projects to the ganglion of the fron- lary) and surrounding, non-fl ask-shaped, periph- tal organ, a sensory organ characteristic for ento- eral cells (Fig. 6.7E, F, H; Wanninger et al. proct larvae, and further to the prototroch nerve 2007 ). Further distinct features shared by the ring (Fig. 6.7G ). The musculature of the swim- entoproct creeping larva and adult mollusks are ming-type larva is fairly complex. It includes a a tetraneurous nervous system comprising two dense set of ring muscles that engulf the episphere ventral (pedal) nerve cords (interconnected by including the apical organ and a prominent ring commissures) and two lateral, more dorsally muscle underlying the prototroch, as well as positioned (visceral) cords (Fig. 6.7E, F ), as numerous supporting muscles. Several apical well as a ventrally intercrossing dorsoventral organ retractor muscles project from the episphere musculature (Haszprunar and Wanninger 2008 ). toward the prototroch (Fuchs and Wanninger Additional neural features of the creeping-type 2008 ). larva include a buccal system and iterated sets In contrast to the swimming-type larvae, of perikarya associated with the pedal nerve creeping-type larvae are released from the cords (Wanninger et al. 2007 ). 6 Entoprocta 97

Fig. 6.6 Schematic representation of the creeping-type trochozoan larvae. Scale bar is 50 μm. The prototroch ( pt ) larva of the solitary entoproct Loxosomella murmanica . encircles the larval foot ( ft) with characteristic anterior Swimming/creeping direction is with the frontal organ cirri (cr ) (From Haszprunar and Wanninger (2008 ), based ( f1 ) forward and not with the apical organ (ao ) as in other on the original drawing by Nielsen ( 1971 ) )

Entoproct creeping-type larvae are highly data available indicate a high degree of variabil- muscular and exhibit an extremely complex ity in the dynamics of this process. In many arrangement of distinct muscle units that is Loxosomella species, the metamorphic compe- unmatched among lophotrochozoan larvae tent larva attaches to the substrate with the fron- (Merkel and Wanninger, unpublished). These tal organ (Nielsen 1971 ). In other entoprocts, include ring muscles underlying the prototroch, e.g., Pedicellina and Barentsia , the frontal organ the frontal and the apical organ, as well as vari- is used for probing the substrate, while fi xation ous longitudinal muscles including frontal to the substrate takes place by secretions of organ and apical organ retractors, body wall specifi c gland cells of the foot (Fig. 6.8 ). muscles, and dorsoventral muscles, allowing for Subsequently, the larval retractors and the the high fl exibility of the body observed in live prototroch ring muscle contract, thus enclosing larvae. the entire larval body. As typical for lophotro- Entoproct larvae have one pair of protone- chozoan larvae, the apical organ and the pro- phridia and eyes are common. They are usually totroch are lost. This is also true for the frontal situated in the region of the frontal organ and organ, although the gland cells used for adhe- consist of a photoreceptor cell, a pigment cell, sion to the substrate in some Loxosomella spe- and a lens cell; they are lost at metamorphosis cies may persist in the adult as a distinct foot (Woollacott and Eakin 1973 ). gland (Nielsen 1971 ). The fate of larval key fea- tures such as neural elements, muscles, protone- phridia, and the subsequent ontogeny of the Metamorphosis corresponding adult systems remain unknown, although loss of major parts of the larval body Our current knowledge on entoproct metamor- plan features is commonly assumed, with de phosis largely relies on a few decades- to novo generation of adult organ systems century- old works (e.g., Harmer 1885 ; Marcus (catastrophic metamorphosis). 1939 ; Nielsen 1967 , 1971 ) and thus requires Following adhesion of the larva to the reassessment using modern methodology. The substrate, the digestive tract rotates such that the 98 A. Wanninger

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Fig. 6.7 Entoproct larvae and larval nervous systems. arrows ), and paired perikarya (1–4). Note also the split of A – C are scanning electron micrographs; D and E are con- the neurite bundles (nb ) that arise from the apical neuropil focal projections of the serotonin-like nervous system. and give rise to neurites that run toward the pedal neurite Line drawings F – H are based on various confocal analy- bundles ( nbf) and to the prototroch neurites (nbp ). ses of the respective larval types. The apical organ is up in ( F) Schematic representation of the serotonin-like ner- A , D , G , and H . Frontal organ is up in B , C , E , and F . ( A ) vous system of the creeping-type larva depicted in E . Swimming-type larva with apical tuft ( at), prototroch ( pt ), (G ) Schematic representation of the nervous system based and small hyposphere (hy ). (B ) Creeping-type larva with on semithin section ( light gray) data by Nielsen (1971 ) view onto the retracted apical organ (ao ). (C ) Ventral view with the fl ask cells of the apical organ (ao ) included of a creeping-type larva with ciliated frontal organ (fo ), ( black). Other components include the frontal ganglion prototroch ( pt), and foot (ft ). (D ) Three serotonin-positive ( fg ), prototroch ring nerve (pn ), and lateral sense organ fl ask cells in the apical organ (ao ) of a swimming-type ( lso ) with connecting nerve (nlso ). prm , prototroch ring larva, antero-ventral view. (E ) Serotonin-like nervous sys- muscle, lm longitudinal muscle. (H ) Serotonin-like apical tem of a creeping-type larva in ventral view. Note the organ of the creeping-type larva shown in E and F shifted much higher complexity of the nervous system than that by 90° to visualize the eight fl ask cells (black ) and the of the swimming-type larva in D , including a complex surrounding bipolar peripheral cells ( gray ), both con- apical organ (ao ), anterior nerve loop (an ), prototroch nected to the underlying neuropil ( np ). ( B , C , E , F , H neurites ( pn ), pedal neurite bundles (arrows ) with com- modifi ed from Wanninger et al. (2007 ); D , G modifi ed missures ( arrowheads ), lateral neurite bundles (double from Fuchs and Wanninger ( 2008 ) ) 6 Entoprocta 99

ABC

Fig. 6.8 Entoproct metamorphosis. (A ) Metamorphic and attach with the frontal organ forward and not with the competent larva with ciliated prototroch (blue ), contracted foot as indicated here. ( C) Metamorphosis involves a 90° foot (red ), as well as apical organ (orange ), frontal organ rotation of the digestive tract and de novo formation of the ( yellow ), and rudiment of the digestive tract (gray ). (B ) tentacles ( green). The region of the larval foot becomes Settlement and adhesion is followed by loss of the pro- part of the atrium of the adult (Redrawn and modifi ed totroch and subsequent degeneration of the apical and after Marcus (1939 ) ) frontal organ. Note that some entoprocts seem to settle mouth and anal opening eventually come to lie on tive tract and the protonephridia are folded off the opposite side of the substrate (Fig. 6.8A, B ). from the atrium (Emschermann 1972). The The stalk grows and the adult tentacles start to scarce data on the exact mechanisms of the bud- form as epithelial outgrowths, eventually sur- ding process (see Nielsen 1971 for a brief rounding the mouth, the anus, and the newly review of the classical works on the topic) sug- formed atrium (Fig. 6.8C ). Signifi cant deviations gest that no distinct endodermal or mesodermal from this process have been reported in species tissues and only few individual mesodermal where the juvenile emerges from the larval body cells of the mother animal appear to be involved after settlement by budding or is preformed in the in the formation of the new bud. Accordingly, live larva (Nielsen 1971 ). Details on the molecu- muscles, gonads, and other organs are most lar, cellular, and morphogenetic mechanisms likely formed from immigrating (mesenchymal? underlying these processes are unknown. stem?) cells, but these processes are not yet understood.

Asexual Reproduction GENE EXPRESSION Asexual reproduction is common in colonial and solitary entoprocts. Buds form as ectoder- No gene expression data on any entoproct species mal protrusions, usually on the latero-oral side exist until today. The few transcriptomes that of the calyx in solitary species. In colonial rep- have been generated have not yet resulted in pub- resentatives, they form at the base of the stalk lished sequences of important developmental and on the stolons that interconnect individual patterning genes such Hox or other homeobox zooids (Emschermann 1972 ; Emschermann and genes, but are likely to be underway. Accordingly, Wanninger 2013 ). The atrium is formed by developmental genetics of Entoprocta is still a invagination from the juvenile bud, which is fi eld wide open to the ambitious developmental followed by growth of the tentacles. The diges- biologist, but methods for culturing and yield of 100 A. Wanninger signifi cant amounts of embryonic, larval, and Emschermann P (1965) Das Protonephridiensystem metamorphosing stages remain to be von Urnatella gracilis Leidy (Kamptozoa). Bau, Entwicklung und Funktion. Z Morphol Ökol Tiere established. 55:859–914 Emschermann P (1972) Loxokalypus socialis gen. et spec. nov. (Kamptozoa, Loxokalypodidae fam. OPEN QUESTIONS nov.), ein neuer Kamptozoentyp aus dem nördlichen Pazifi schen Ozean. Ein Vorschlag zur Neufassung der Kamptozoensystematik. Mar Biol 12:237–254 • Early developmental events following gastru- Emschermann P (1982) Les Kamptozoaires. État actuel lation, including gene expression, cell lineage, de nos connaissances sur leur anatomie, leur dével- and organogenesis. oppement, leur biologie et leur position phylogéné- tique. Bull Soc Zool Fr 107:317–344 • Developmental gene expression (especially Emschermann P, Wanninger A (2013) Kamptozoa. In: Hox, ParaHox genes) during larval develop- Rieger G, Westheide W (eds) Spezielle Zoologie. ment and metamorphosis. Springer Spektrum, Heidelberg • Gene expression profi les of the prototroch and Fuchs J, Wanninger A (2008) Reconstruction of the neuromuscular system of the swimming-type larva the apical organ. of Loxosomella atkinsae (Entoprocta) as inferred by • Body plan remodeling during metamorphosis: fl uorescence labelling and confocal microscopy. Org Which, if any, larval elements contribute to Divers Evol 8:325–335 the adult body plan? Fuchs J, Bright M, Funch P, Wanninger A (2006) Immunocytochemistry of the neuromuscular sys- • Fate of the apical organ after metamorphosis. tems of Loxosomella vivipara and L. parguerensis • Cellular and molecular mechanisms of clonal (Entoprocta: Loxosomatidae). J Morphol 267:866–883 reproduction (budding). Fuchs J, Iseto T, Hirose M, Sundberg P, Obst M (2010) The fi rst internal molecular phylogeny of the animal phylum Entoprocta (Kamptozoa). Mol Phylogenet Acknowledgments I thank Marion Hüffel for help with Evol 56:370–379 the graphical representations used in this chapter as well Gerould JH (1903) Studies on the embryology of the as for invaluable support on many aspects connected to Sipunculidae. I. The embryonal envelope and its this treatise project. The University of Vienna, especially homologue. Mark Anniversary Volume. pp 437–452 the Faculty of Life Sciences, is thanked for the generous Harmer SF (1885) On the structure and life history of support in establishing the Wanninger lab during the past Loxosoma . Q J Microsc Sci 25:261–337, pls. 19–21 years. I am also grateful for the previous support from the Haszprunar G, Wanninger A (2008) On the fi ne structure Danish Research Council (FNU), the Carlsberg of the creeping larva of Loxosomella murmanica : addi- Foundation, and the European Commission for funding tional evidence for a clade of Kamptozoa (Entoprocta) my research during my years in Copenhagen. and Mollusca. Acta Zool (Stockholm) 89:137–148 Hatschek B (1877) Embryonalentwicklung und Knospung der Pedicellina echinata . Z Wiss Zool 29:502–549 References Heath H (1898) The development of Ischnochiton . Zool Jahrb Abt Anat Ontog Tiere 12:567–656 Hejnol A, Obst M, Stamatakis A, Ott M, Rouse GW, Ax P (1999) Das System der Metazoa, 2nd edn. Gustav Edgecombe GD, Martinez P, Baguñà J, Bailly X, Fischer, Stuttgart Jondelius U, Wiens M, Müller WEG, Seaver E, Baba K (1951) General sketch of the development in a Wheeler WC, Martindale MQ, Giribet G, Dunn CW solenogastre, Epimenia verrucosa (Nierstrasz). Misc (2009) Assessing the root of bilaterian animals with Rep Res Inst Nat Res (Tokyo) 19–21:38–46 scalable phylogenomic methods. Proc R Soc Lond B Bartolomaeus T (1993) Die Leibeshöhlenverhältnisse 276:4261–4270 und Nephridialorgane der Bilateria – Ultrastruktur, Malakhov VV (1990) Description of the development of Entwicklung und Evolution. University of Göttingen, Ascopodaria discreta (Coloniales, Barentsiidae) and Göttingen discussion of the Kamptozoa status in the animal king- Dunn CW, Hejnol A, Matus DQ, Pang K, Browne WE, dom. Zool Zh 69:20–30 Smith SA, Seaver E, Rouse GW, Obst M, Edgecombe Marcus E (1939) Bryozoários marinhos brasileiros III. Bol GD, Sørensen MV, Haddock SHD, Schmidt-Rhaesa Fac Fil Ciên Letr Univ S Paulo, XIII. Zoologica A, Okusu A, Kristensen RM, Wheeler WC, Martindale 3:111–354 MQ, Giribet G (2008) Broad phylogenomic sampling Maslakova SA, Martindale MQ, Norenburg JL (2004) improves resolution of the animal tree of life. Nature Fundamental properties of the spiralian developmen- 452:745–749 tal program are displayed by the basal nemertean 6 Entoprocta 101

Carinoma tremaphoros (Palaeonemertea, Nemertea). crozieri; implications for the evolution of spiralian life Dev Biol 267:342–360 history traits. Front Zool 7:12 Merkel J, Wollesen T, Lieb B, Wanninger A (2012) Spiral Schwaha T, Wood TS, Wanninger A (2010) Trapped cleavage and early embryology of a loxosomatid ento- in freshwater: the internal anatomy of the entoproct proct and the usefulness of spiralian apical cross pat- Loxosomatoides sirindhornae . Front Zool 7:7 terns for phylogenetic inferences. BMC Dev Biol 12:11 Wanninger A (2004) Myo-anatomy of juvenile and adult Nesnidal MP, Helmkampf M, Meyer A, Witek A, loxosomatid Entoprocta and the use of muscular Bruchhaus I, Ebersberger I, Hankeln T, Lieb B, body plans for phylogenetic inferences. J Morphol Struck TH, Hausdorf B (2013) New phylogenomic 261:249–257 data support the monophyly of Lophophorata and an Wanninger A (2009) Shaping the things to come: ectoproct-phoronid clade and indicate that Polyzoa ontogeny of lophotrochozoan neuromuscular sys- and Kryptrochozoa are caused by systematic bias. tems and the Tetraneuralia concept. Biol Bull 216: BMC Evol Biol 13:253 293–306 Newby WW (1932) The early embryology of the echi- Wanninger A, Fuchs J, Haszprunar G (2007) The anatomy uroid, Urechis . Biol Bull 63:387–399 of the serotonergic nervous system of an entoproct Nickerson WS (1901) On Loxosoma davenporti sp. nov. creeping-type larva and its phylogenetic implications. an endoproct from the New England coast. J Morphol Invertebr Biol 126:268–278 17:351–380, pls. 32–33 Wasson K (1997) Sexual modes in the colonial Nielsen C (1967) Metamorphosis of the larva of kamptozoan genus Barentsia . Biol Bull 193: Loxosomella murmanica (Nilus) (Entoprocta). Ophelia 163–170 4:85–89 Wilson EB (1892) The cell-lineage of Nereis . A contribu- Nielsen C (1971) Entoproct life-cycles and the entoproct/ tion to the cytogeny of the annelid body. J Morphol ectoproct relationship. Ophelia 9:209–341 6:361–480 Nielsen C (2012) Animal evolution: interrelationships of Wilson EB (1904) Experimental studies in germinal the living phyla. Oxford University Press, Oxford localization. II. Experiments on the cleavage- Nielsen C, Jespersen A (1997) Entoprocta. In: Harrison mosaic in Patella and Dentalium. J Exp Zool FW (ed) Microscopic anatomy of invertebrates, vol 1:197–268 13. Wiley-Liss, New York Woollacott RM, Eakin RM (1973) Ultrastructure of a Rawlinson KA (2010) Embryonic and post-embryonic potential photoreceptoral organ in the larva of an ento- development of the polyclad fl atworm Maritigrella proct. J Ultrastruct Res 43:412–425 Mollusca 7 Andreas Wanninger and Tim Wollesen

Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger. A. Wanninger (*) • T. Wollesen Department of Integrative Zoology , University of Vienna , Althanstrasse 14 , Vienna 1090 , Austria e-mail: [email protected]

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 103 DOI 10.1007/978-3-7091-1871-9_7, © Springer-Verlag Wien 2015 104 A. Wanninger and T. Wollesen

INTRODUCTION two- valved fi lter feeders (Bivalvia such as mus- sels and clams), as well as the single-shelled With probably around 200,000 extant species, Monoplacophora (Tryblidia), Gastropoda Mollusca is the second-most speciose phylum (snails, slugs), Scaphopoda (tusk shells), and after Hexapoda. However, what makes mol- Cephalopoda (octopuses, squids, nautiluses) lusks particularly interesting from an evolution- (Fig. 7.1 ; for a comprehensive recent account on ary perspective is not their richness in species various aspects on molluscan phylogeny and evo- as such, but rather the huge variety of body plan lution, see Ponder and Lindberg 2008 ). phenotypes exhibited by its representatives. With such a diverse morphological toolkit at These include cylindrical, shell-less, spicule- hand, mollusks have managed to conquer almost bearing, wormlike, crawling, and burrowing crea- all natural aquatic and terrestrial realms, and the tures (Neomeniomorpha or Solenogastres and spectacular “fl ying squid” (Todarodes pacifi cus ) Chaetodermomorpha or Caudofoveata), eight- is even able to get partly airborne by using its shelled grazers (Polyplacophora or chitons), fi ns, arms, and tentacles as wings. The origin of

Fig. 7.1 Molluscan diversity. Anterior faces to the right in all aspects. Polyplacophora ( Acanthochitona crinita , dorsal view). Neomeniomorpha ( Helluoherpia aegiri , lateral view; courtesy of Maik Scherholz). Chaetodermomorpha ( Falcidens cros- sotus , lateral view). Monoplacophora ( Laevipilina theresae , dorsal view; © Michael Schrödl, 2015. All Rights Reserved). Bivalvia ( Nucula tumidula , lateral view). Gastropoda (Patella vulgata , dorsal view). Scaphopoda (Antalis entalis , lateral view). Cephalopoda (Idiosepius notoides , lateral view). Images not to scale 7 Mollusca 105 the phylum dates back to at least the Early signifi cant impact on the plankton communities Cambrian some 540 million years ago, but is in the respective habitats, for example, in fresh- most likely rooted deep in the Precambrian water streams and lakes throughout Europe and (Vendian) era (Parkhaev 2008 ). North America), in combination with fast distri- bution rates by free- swimming veliger larvae that result from seasonal mass spawning events. Other Man and Mollusk important invasive species include the terrestrial pulmonate gastropod Arion vulgaris (Spanish Humans have had long-standing ambivalent rela- slug) that has spread throughout Europe during tionships with mollusks. Oysters, scallops, cock- the last half-century, probably from Portugal or les, mussels, octopods, cuttlefi sh, and certain France. Originally constrained in reproductive snails provide a welcome protein source, and gas- success by dry climates, the moister and cooler tropod, bivalve, and scaphopod shells have been conditions in Central Europe have led to regional widely used for decorative and monetary pur- explosions in individual numbers of this species, poses for centuries (including, e.g., the commer- thereby causing severe horticultural and eco- cial production of pearls). On the other hand, the nomic damage. attractive cone snails ( Conus ) cause regular, fatal These examples illustrate that molluscan evo- accidents among shell collectors, but at the same lutionary and ecological success is not merely time high hopes are put in research on the power- historical, but instead an ongoing process ful neurotoxins produced in their salivary glands that is intimately linked to the reproductive for pain relief and the treatment of chronic neural and developmental biology of the phylum’s diseases including Alzheimer’s, Parkinson’s, or protagonists. Considering the numerous aspects depression (Anderson and Bokor 2012 ). These in which molluscan developmental biology plays current approaches may be seen as a continuation a vital role, ranging from applied approaches of a long tradition of mollusks as contributors to such as pest control and protein production to neurobiological research which, for example, has evolutionary- oriented research, the importance led to detailed reconstructions of the immensely of comparative developmental data becomes complex cephalopod brain (reviewed in Nixon immediately obvious. This potential has, how- and Young 2003 ) and the discovery and distribu- ever, only been scarcely exploited to date, which tion of neuroactive substances (Messenger 1996 ). is at least partly due to the lack of a solid pool A number of mollusks are viewed as notori- of model system species from the various class- ous pest species, such as several freshwater gas- level taxa from which ontogenetic material tropods that house intermediate developmental can readily be obtained (see boxed text). Thus, stages of important human pathogenic parasites, today’s comparative developmental malacolo- including fl ukes and nematodes, which may cause gists still need to travel the world to collect adult severe medical problems, particularly in sub- specimens from natural habitats and experiment Saharan Africa and Southeast Asia. Semisessile with rearing conditions including induction of bivalves such as the blue mussel (Mytilus ) or the spawning and (in many aquatic species) meta- zebra mussel (Dreissena polymorpha ) may infl ict morphosis, often with erratic success. In times economic damage as biofouling organisms that of increased pressure on high publication output, attach to plumbing and ship bodies by their bys- these obstacles still render mollusks relatively sus threads. Dreissena represents one of the sev- unattractive models for many developmental eral highly competitive neozoan mollusks with biologists. Thus, what we do know on the various a particularly broad ecological tolerance that aspects of molluscan comparative evolutionary has resulted in rapid colonialization of formerly developmental biology sadly lags behind of what non-native habitats, with in parts dramatic con- we should know, especially given the above- sequences for the native fauna. This is due to its mentioned considerably strong bonds between high effi ciency as adult fi lter feeder (which has humans and mollusks on various grounds. 106 A. Wanninger and T. Wollesen

Potential Molluscan Models in EvoDevo although it is abundant and fulfi lls At present, hardly any molluscan species is numerous criteria that render it suitable available that can be reliably reared in high as an organism for aquaculture. In Korea, numbers and over many generations year oogenesis is initiated in April when the round under controlled laboratory conditions water temperature reaches 16.5 °C. First from spawning through the various life cycle spawning occurs in July at around 27 °C, stages to sexual maturity. Sole exceptions are and mature adults go into resting stage in a few direct developing and phylogenetically November. Eggs are relatively small and derived gastropods with internal fertilization measure 45–50 μm in diameter. Zygotes such as the great pond snail, Lymnaea stag- undergo fi rst cleavage after 1.2 h postfer- nalis . As a consequence, research techniques tilization (hpf), and the blastula stage is that involve transgenesis or reverse genetics reached at approximately 5 hpf at (RNAi, morpholinos) can still not be routinely 24 °C. Gastrulation occurs at 7 hpf, the performed on any molluscan model. Given trochophore stage is reached by 9.5 hpf, their cryptic lifestyle and their often long and D-shaped larvae develop after 20–25 period until sexual maturity, there is little hope hpf. Larvae measure approximately that this will change anytime soon for most 40 μm in length and are thus in principal molluscan clades. well-suited for whole-mount in situ Luckily enough, however, some bivalves hybridization or immunochemical experi- and gastropods can readily be spawned, ments, but their small size renders data yielding thousands of embryos at a time. interpretation (e.g., localization of gene Thus, although adult specimens still need transcripts in relation to developing to be collected from the wild, a few re- organs) often diffi cult, even if the speci- presentatives have the potential to develop mens are sectioned. Due to their largely into models for which advanced molecular asymmetric shape as adults, oysters are methods may be established in the future. well-suited for studies concerned with the However, long planktonic phases including ontogenetic and molecular bases of left- larval planktotrophy usually require addi- right asymmetry. tional facilities to culture the respective algal B. Gastropoda: Ilyanassa obsoleta diet. Here, a few examples are mentioned for The Eastern mudsnail belongs to the which signifi cant EvoDevo data have been Caenogastropoda and occurs on mud published, with the focus on indirect devel- and sand fl ats as well as in estuaries oping species in the gastropods and bivalves, along the Eastern and Western coastline although other candidates exist and are of the USA. Its whorled conical shell is explored in various labs (e.g., the freshwa- dark brownish to black and 1.5–3 cm in ter pond snail Lymnaea or the Pacifi c oyster, length. I. obsoleta may also host a range Crassostrea gigas, for which the genome has of trematode species including those recently been sequenced; see http://gigadb. that cause “swimmer’s itch.” The East- org/dataset/100030 ). ern mudsnail is a facultative scavenger A. Bivalvia: Saccostrea kegaki and feeds on detritus. Animals can also The Japanese spiny oyster Saccostrea be purchased from the Marine Resources kegaki occurs along Japanese, Korean, Center of the Marine Biological Labora- and Taiwanese shorelines. It is an encrust- tory in Woods Hole, MA, USA ( http:// ing oyster which may easily be collected www.mbl.edu). Adults are easily kept in rocky intertidal zones. Due to its small in the laboratory, fertilization is inter- size, it is of no commercial interest nal, and large numbers of fertilized eggs 7 Mollusca 107

are laid throughout the year, although Embryos may also be separated from the their natural spawning season is spring mother animal, allowing for clutches to be to early summer. Development takes reared individually (preferably with the 7–10 days from egg laying to hatching addition of antibiotics to avoid fungal and of the planktotrophic veliger larvae and bacterial growth). Fertilized eggs measure another 2–2.5 weeks to metamorphic about 180 μm, the fi rst cleavage occurs competence at 23–25 °C. Embryos have a after 7.3 h, and the 25-cell stage is reached size of 200–300 μm. They are right sized at 20 h after oviposition at 21–22 °C. for whole-mount in situ hybridization or C. fornicata produces nurse eggs in the immunochemical experiments. The adult capsules. The species does not exhibit a and embryonic morphology of I. obsoleta distinct trochophore stage, and the fi rst is well-studied and serves as an excel- veliger larvae may develop after 7 days. lent reference for future developmental Hatching occurs after 11–12 days, and the analyses. I. obsoleta adheres to the typi- veliger larvae metamorphose after several cal spiralian developmental program, and weeks. Cell lineage studies have been its relatively large and transparent blasto- carried out on C. fornicata, and its meres facilitate manipulation of embryos embryogenesis is well documented. (electroporation, cell ablation, dye injec- Manipulation of embryos may be per- tion). Accordingly, I. obsoleta has served formed, and several experimental proto- as a model for developmental studies cols for, e.g., microinjection of blastomeres concerned with early cleavage, the asym- with fl uorescent dyes, are available. metry of cell division, and gene expres- D . Cephalopoda: Sepia offi cinalis sion, and still continues to do so. The European cuttlefi sh Sepia offi cina- C. Gastropoda: Crepidula fornicata lis occurs in the Mediterranean, the Atlan- Also a caenogastropod, the common tic Ocean, the North Sea, and parts of the slipper shell lives intertidally along the Baltic Sea. Due to its edibility, its adequate East Coast of the USA and has been intro- size, and its mass spawning events in duced to several other parts of the world. spring, it is of importance for local fi shery Adults are suspension feeders, long-lived industries and a promising candidate for with 6–11 years, are easy to keep in future aquaculture enterprises. Its diet is seatables with running seawater, and toler- variable and includes crustaceans, fi sh, ate fl uctuations in salinity. As a protandric and other cuttlefi sh. The dorsal mantle hermaphrodite, C. fornicata changes its length may reach more than 40 cm; how- sex from male to female during adulthood. ever, in captivity, depending on aquaria Adult individuals are usually attached to size and diet, adults may only grow up to each other in stacks with the oldest and 10 cm. S. offi cinalis was one of the fi rst largest animal located at the base of the cephalopods for which the life cycle was stack. Fertilization is internal, and gravid closed in captivity. In their natural adults produce numerous eggs. Egg environment egg clutches are attached to laying may occur spontaneously or may submersed hard substrates during spring be induced by chilling and subsequent and early summer which can readily be warming of the seawater in which the collected. In captivity, eggs can also be adults are kept (e.g., from 14 to 25 °C). obtained from adults which may lay eggs Until hatching, brood care takes place, and for a period of up to 7 months per year. the 1,000–50,000 embryos per female The gelatinous envelopes of freshly laid hatch as planktotrophic veliger larvae. eggs of S. offi cinalis are approximately 108 A. Wanninger and T. Wollesen

1 cm in size and black in color due to the than 2.5 cm in size and endure rapidly addition of ink during egg laying. They changing environmental conditions such become transparent with time. The opaque as fl uctuations in temperature, oxygen, envelopes are easily removed, and early and salinity levels. Due to their minute developmental stages can be observed size, pygmy squids are of no commer- through the transparent chorion. From fer- cial value. In captivity, adults feed on tilization to hatching, it takes less than small crustaceans and fi sh and appear 50 days at 20 °C or less than 100 days at not to be cannibalistic. They attach egg 15 °C. After hatching, individuals are easy clutches to glass surfaces or other items to rear to sexual maturity, but tend to be provided. At 23–25 °C water tempera- cannibalistic. Research areas concerned ture, development takes approximately with camoufl age, communication, and 10 days from egg laying to the hatching related behavioral patterns including their squid. The fi rst days after hatching are molecular underpinnings greatly profi t crucial for rearing Idiosepius notoides , from S. offi cinalis as a model. Moreover, and hatchlings have not been carried over various studies on brain development have this stage and eventually die of unknown been carried out. The large-sized embryos reasons. With approximately 0.5–1 mm are, however, not ideal for whole-mount in size, early developmental stages are immunochemical or in situ hybridization small compared to those of other cepha- experiments. First estimates consider the lopods. Being rather transparent, they are genome of S. offi cinalis with 4.5 Gb as ideally suited for immunochemical and rather large among cephalopods. in situ hybridization experiments. First E . Cephalopoda: Idiosepius notoides estimates suggest that I. notoides has a The southern pygmy squid Idiose- relatively small genome (2.1 Gb), which pius notoides lives in seagrass beds off renders it an ideal candidate for sequenc- Eastern Australia. Adults reach no more ing efforts.

7 Mollusca 109

Molluscan Origin and Phylogeny etal elements in the polyplacophoran body plan are interpreted as remnants of ancestral segmenta- The undisputed monophyly of Mollusca implies tion. However, an annelid-like posterior growth that the wide morphological diversity exhib- zone is absent in mollusks, and detailed compara- ited by their recent (and extinct) representa- tive analyses of polychaete and aculiferan neuro- tives originated from a common body plan of an muscular development and skeletogenesis have early ancestor, often conceptually, but neverthe- uniformly rejected the idea of a segmented mol- less oddly, referred to as “hypothetical ancestral luscan ancestor (Friedrich et al. 2002 ; Wanninger mollusk (HAM).” Obviously, assessing the mor- and Haszprunar 2002a ; Nielsen et al. 2007 ; phological radiation that emerged from that very Wanninger 2009 ; Haszprunar and Wanninger real last common molluscan ancestor, together 2012 ; Scherholz et al. 2013 ). Morphologically, by with its evolutionary and developmental driving far the strongest support is provided for a scenario forces, fi rstly calls for a reconstruction of the that suggests a mollusk- entoproct clade based on ancestor itself. However, with a solid agreement numerous synapomorphies shared by larval ento- on molluscan inter- as well as intrarelationships procts and larval and adult aplacophorans and still largely lacking, little progress has hitherto polyplacophorans. This includes a creeping foot, been made to this end. This may, however, be anterior sensory cirri, a highly complex larval api- about to change. Several independent phyloge- cal organ, and a tetraneurous nervous system netic analyses employing large sequence datasets (hence Tetraneuralia, formerly Lacunifera or have confi rmed a classical hypothesis that had Sinusoida concept; see Bartolomaeus 1993 ; Ax been largely dormant for the last 15 or 20 years, 1999 ; Wanninger et al. 2007 ; Haszprunar and namely, that the shell-less, wormy aplacophoran Wanninger 2008 ; Wanninger 2009 ). taxa Neomeniomorpha and Chaetodermomorpha, With (molecular) phylogeny still painting a together with the Polyplacophora, form a mono- rather blurred picture of molluscan affi nities and phyletic clade, the Aculifera (Fig. 7.2 ). This early evolution, developmental studies may pro- assemblage is currently believed to form the vide the key to clarify important issues. Recent sister taxon to all remaining mollusks – the pri- work on the hitherto largely neglected aplacoph- marily single-shelled Conchifera (Kocot et al. orans has shown that muscle development in the 2011 ; Smith et al. 2011 ; Vinther et al. 2012 ). neomeniomorph Wirenia argentea undergoes a Such a dichotomy at the base of the molluscan series of polyplacophoran-like stages with dis- tree, however, still leaves the question open as to tinct muscular systems being confi ned to both which of the aculiferan or conchiferan taxa bears lineages alone (Scherholz et al. 2013 ; see below). most resemblance to their last common ancestor. This strongly argues for a polyplacophoran-like Accordingly, questions such as Did the mollusks ancestor of neomeniomorphs and thus probably derive from an aplacophoran - like forefather ? Or, the entire Aculifera – which provides us with the rather, from a chiton - like creature ? Was it a uni - basal condition for at least one of the two early shelled , conchiferan - like species , maybe resem- molluscan branches. bling a monoplacophoran ? remain unanswered. In any case, and despite the contentious Usually, outgroup comparisons are an impor- phylogenetic issues, the unmatched morphologi- tant means for assessing ancestral character states cal plasticity of their body plan renders mollusks of a given taxon. Potential molluscan sister groups prime organisms for the study of the evolution of among the Lophotrochozoa abound, but the nature morphological novelties and their underlying of the closest molluscan ally remains controver- molecular, cellular, and morphogenetic mecha- sial. Various scenarios have been proposed, includ- nisms. As a consequence of this wide phenotypic ing the reoccurring annelid-mollusk hypothesis, variation, clear-cut autapomorphies for the often favoring a segmented ancestral mollusk, phylum are scarce, since all potential characters whereby the repetitive muscular, neural, and skel- have undergone signifi cant remodeling from their 110 A. Wanninger and T. Wollesen

Fig. 7.2 Competing hypotheses on the interrelationships ( 2011), based on phylogenomic data. Monoplacophoran of molluscan class-level taxa. ( A ) After Waller (1998 ), image © Michael Schrödl, 2015. All Rights Reserved; based on morphological data. ( B) After Haszprunar neomeniomorph image courtesy of Maik Scherholz ( 2000 ), based on morphological data. (C ) After Smith et al. 7 Mollusca 111 ancestral state. Even the most commonly prevalent, with one pair of longitudinal nerve cords recognized molluscan feature, the radula (rasping usually running ventrally and a second pair of nerve tongue) has been lost in an entire class-level lin- cords situated more dorsolaterally to it. eage, the Bivalvia, and complete reorganization of Although we are only beginning to understand the body architecture has taken place within the the molecular, cellular, and morphogenetic heterobranch gastropods and the bivalves, often mechanisms that underlie the establishment of resulting in secondary fl atworm-like (e.g., marine molluscan key features and evolutionary novel- nudibranchs) or cylindrical vermiform organisms ties, important recent progress has been achieved. (e.g., interstitial slugs such as Rhodope and Thus, in the wake of modern-day EvoDevo, Helminthope or shipworms such as Teredo). mollusks re-enter the limelight of comparative Again, the molecular and developmental base that developmental research. It is thus about time to underlies these drastic phenotypic deviations from summarize the landmarks of our current know- a common ground plan remains unexplored. ledge on molluscan evolutionary developmental On the cellular level, the free-fl oating rhogocytes biology and point towards the numerous (pore cells), hemocoelic cells that have been linked unclarifi ed issues associated with this fascinating to hemocyanin synthesis, excretion, and various phylum that deserves our special attention. metabolic processes, are thought to be confi ned to mollusks alone and may constitute the only defi nite feature exhibited by all recent mollusks (Haszprunar EARLY DEVELOPMENT 1996; Stewart et al. 2014 ). On organ system level, a closer look reveals that almost all taxa exhibit a Reproduction and Cleavage foot of some sort, which may serve as creeping, bur- rowing, swimming, sensory, or grooming device. Many mollusks, including the chaetodermo- A dorsoventral musculature that often attaches to morphs, polyplacophorans, bivalves, scaphopods, the shell(s) (therefore often referred to as “shell and basal gastropods, are dioecious broadcast musculature” in conchiferans) and intercrosses spawners with external fertilization and indirect above the foot sole is present in all class-level taxa development via trochophore and, in most gastro- (Haszprunar and Wanninger 2000 ; Wanninger pods and bivalves, veliger-type larvae (Figs. 7.3 2009 ). In addition, a tetraneurous nervous system is and 7.4 ). Neomeniomorphs are hermaphroditic

Fig. 7.3 Gross morphology of a molluscan trochophore- like larva. Here, an early scaphopod larva is depicted by scanning electron microscopy, with typical partitioning of the body into pretrochal episphere (including the apical organ), large, ciliated trochoblasts forming the prototroch, and the posttrochal hyposphere. Anterior is up and total size of the larva is 300 μm 112 A. Wanninger and T. Wollesen

Fig. 7.4 Diversity of molluscan larvae. All images are (D ) Late-stage polyplacophoran larva (Mopalia muscosa ) scanning electron micrographs with anterior facing up in lateral view with long cilia of the apical tuft (at ), pro- except for (E ) where anterior is to the left. Scale bars are totroch ( pt), and developing shell plates (arrows ). The 25 μm in (A – C) and 50 μm in (D – F). ( A) Ventral view of anlagen of the foot (ft ) and mantle fold (girdle, gi ) are vis- an early trochophore larva of a basal gastropod, the patel- ible as distinct, ciliated regions. (E ) Lateral left view of a logastropod Lottia , showing apical tuft (at ), prototroch bivalve pediveliger larva of the shipworm Lyrodus pedicel- (pt ), and mouth (mo ). Note the individual cells in the larval latus with larval shell (protoconch II, pc2 ), velum ( ve), and hyposphere. (B ) Ventroposterior view of a later stage larva reduced apical tuft (at ) (image courtesy of Reuben of the neomeniomorph Wirenia argentea showing cells of Shipway). (F ) Late-stage larva close to metamorphic com- the apical cap (ca ), ciliary band of the developing foot (ft ), petence of the scaphopod Antalis entalis in ventral view and telotroch ( tt) (Modifi ed from Todt and Wanninger with elongated embryonic shell (protoconch I, pc1 ). While (2010 )). (C ) Larva of the chaetodermomorph Chaetoderma the apical tuft has been lost, the cilia of the prototroch (pt ) nitidulum with prominent prototroch ( pt) and telotroch ( tt). are still retained

and lay clutches of uncleaved zygotes which like- tropods, sometimes involving brood care. Larval wise develop into planktonic larvae. Both dioecy stages are entirely lacking in the dioecious cepha- and hermaphroditism occur in the monoplaco- lopods, and brood care is particularly pronounced phorans, and indirect development via brooded in the octopods, which may carry the developing larvae is assumed. There is a strong tendency embryos either in the mantle cavity (as, e.g., in towards hermaphroditism and direct develop- the pelagic Argonauta ) or guard and ventilate ment in marine and terrestrial heterobranch gas- them in rock crevices (e.g., Octopus ). 7 Mollusca 113

Some free-spawners may be successfully coidal cleavage, whereby the early embryo forms inseminated artifi cially following dissection of as a monolayered disk of blastomeres (blastodisc) the gonads (e.g., several patellogastropods and on the animal pole of the egg (Fig. 7.5 ). Here, the numerous bivalves), but often the spawning event polar bodies are ejected some hours after fertil- as such is required to yield fertilizable oocytes. ization, and the fi rst cleavage furrow emerges Follicle cells are common and oocytes are brightly close by and determines the prospective longitu- colored in orange, red, yellow, or green in some dinal axis (Naef 1928 ; Boletzky 1989 ; Boletzky species. Polyplacophoran eggs are often covered et al. 2006 ). In this case, cleavage solely involves by a characteristic, richly sculptured, species-spe- the blastodisc, and the second cleavage furrow cifi c hull (Eernisse and Reynolds 1994 ). In the runs perpendicular to the fi rst cleavage plane, polyplacophoran Mopalia muscosa , sperm mito- while the third cleavage plane is located again chondria and centrioles do not penetrate the egg perpendicular to the second one (Fig. 7.5 ). surface, a phenomenon that so far appears to be unique among metazoans (Buckland-Nicks 2013). In most mollusks, meiosis is only com- Cell Lineage and the 4d Cell pleted after fertilization, resulting in shedding of the polar bodies at the animal pole of the zygote While a number of classical accounts on mollus- (Fig. 7.5 ; Longo 1983 ). Accordingly, the animal- can (mostly gastropod) early embryology were vegetal axis of the embryo is discernible from this generated in the late nineteenth and early twenti- stage onwards. The point of fusion of the sperm eth century (Fig. 7.6; Lacaze-Duthiers 1858 ; cell with the oocyte determines the orientation of Kowalevsky 1883a , b ; Lillie 1895 ; Conklin 1897 ; the fi rst cleavage furrow. Cleavage is usually total Heath 1898; Wilson 1898; Meisenheimer 1901 ; (holoblastic) and may be equal or unequal. Robert 1902 ; Casteel 1904 ; Wierzejski 1905 ; Cytoplasmic segregations may result in the for- Clement 1986 ; Van Dam 1986 ), most of our mation of so-called polar lobes as early as prior to detailed knowledge concerning the subsequent fi rst cleavage (Fig. 7.5 ), and several gastropods fates of individual cells produced during early (e.g., Ilyanassa ) and the scaphopod Antalis cleavage cycles stems from a very limited number exhibit a so-called trefoil stage which results from of polyplacophoran (Chaetopleura apiculata ) a polar lobe being formed following fi rst cleavage and gastropod representatives (e.g., the patello- (Fig. 7.5 ; van Dongen and Geilenkirchen 1974 , gastropod limpet Patella vulgata , the slipper shell 1975 , 1976). After second cleavage the macro- Crepidula fornicata, and the mudsnail Ilyanassa meres A, B, C, and D are established and as a con- obsoleta ) (Render 1991 , 1997 ; Henry et al. 2004 ; sequence also the anterior- posterior axis of the Hejnol et al. 2007 ; Goulding 2009 ; see also embryo, which runs through the B–D cells. At Nielsen 2004 , Hejnol 2010 for review). Some this stage the typical spiral cleavage pattern is ini- data on early cell fates are available for a scapho- tiated, whereby the four daughter cells (micro- pod ( Antalis entalis , formerly Dentalium dentale ; meres) come to lie above each cleavage furrow of van Dongen and Geilenkirchen 1974 , 1975 , their mother cells at the animal pole. Each subse- 1976 ). Rudimentary, century-old reports are quent cleavage cycle results in a set of additional available for bivalves, both on freshwater species micromeres that involves such a 45° twist of their (the painter’s mussel Unio and the zebra mitotic spindle axes relative to that of the mother mussel Dreissena polymorpha; Lillie 1895 ; cells, however, with alternating clockwise and Meisenheimer 1901 ), supplemented by few counterclockwise chirality between generations modern accounts (on the Japanese spiny oyster of developing micromeres. As a result, the cells in Saccostrea kegaki and the Japanese purple mussel the cleaving embryo appear spirally arranged if Septifer virgatus), but these largely focus on the viewed from the animal pole (Fig. 7.5). In the cell lineage of the bivalved shell (Kin et al. 2009 ; exceptionally large and yolky cephalopod Kurita et al. 2009 ). No cell genealogies have hith- embryos, the spiral pattern has given way to dis- erto been reconstructed for any neomeniomorph, 114 A. Wanninger and T. Wollesen

1a ectoderm A 2a ectoderm 1A 2A 3a ectomesoderm AB 3A 4a mesoderm 4A endoderm 1b ectoderm B 2b ectoderm 1B 3b ectomesoderm 2B 4b mesoderm zygote 3B 4B endoderm 1c ectoderm C 2c ectoderm 1C 3c ectoderm 2C CD 3C 4c endoderm 4C endoderm 1d ectoderm D 2d ectoderm 1D 3d ectoderm 2D 3D 4d endomesoderm 4D yolk

Fig. 7.5 Cleavage patterns in mollusks. Top : Typical spi- second polar lobe at second cleavage. Small dots indicate ral cleavage pattern of the equal cleaving gastropod polar bodies. Micromeres are color-coded by quadrant in Trochus , fi rst forming four macromeres that defi ne the both embryos to the right. A-quadrant, green ; B-quadrant, four quadrants of the embryo ( A – D ), then the subsequent blue ; C-quadrant, yellow ; D-quadrant, magenta . From sets of micromeres emerge ( A – D ). Daughter cells always Goulding (2009 ). Bottom left: Micromere and macromere come to lie in the cleavage furrows of their mother cells. contributions to the three germ layers in the gastropod Lineages of the four quadrants are color-coded in the large Crepidula. Bottom right: Discoidal cleavage of cephalo- embryo to the right. A-quadrant, green ; B-quadrant, blue ; pods results in a monolayer of blastomeres (blastodisc). C-quadrant, yellow; D-quadrant, magenta. Upper row of Animal view of an embryo after the third ( upper left ), cleaving embryos: animal view; lower row: lateral view fourth ( upper right ), fi fth (lower left), and sixth ( lower (slightly modifi ed from Goulding ( 2009 )). Middle : Spiral right) cleavage. Red numbers indicate the fi rst to fi fth cleavage pattern of the unequal cleaving gastropod cleavage furrow. Blastomeres are located in the center of Ilyanassa. The embryo forms a fi rst polar lobe (PL ) prior the cytoplasmic cap, while blastocones surround the latter to the establishment of the four quadrants (macromeres and are continuous with the yolk syncytium ( stippled A – D ), resulting in an intermediate “trefoil stage” and a lines ) (Redrawn after Boletzky (1989 ) ) 7 Mollusca 115

Fig. 7.6 Polyplacophoran cell lineage and postembry- (1898 ). Primary trochoblasts are in yellow , mesoderm pre- onic development. Camera lucida drawings of selected cursor cells in red developmental stages from the classical work by Heath 116 A. Wanninger and T. Wollesen chaetodermomorph, monoplacophoran, or cepha- bilateral and largely disorganized embryos simi- lopod species. Accordingly, comparative analyses lar to ones raised after polar lobe deletion (van of molluscan cell fates are largely restricted to the den Biggelaar 1977 ; Martindale et al. 1985 ; few taxa summarized in Table 7.1 . As expected Martindale 1986 ; Kühtreiber et al. 1988; Damen by the shared conserved spiral cleavage program, and Dictus 1996 ; Lambert and Nagy 2003). these lineages show a relatively high degree of Chemical inhibition of the 3D cell produced conservation with respect to the contribution of radialized gastropod, polyplacophoran, and, to individual blastomeres to certain germ layers some degree, scaphopod embryos (Gonzales (Fig. 7.5 ). However, differences do occur if mor- et al. 2007 ). There is recent evidence from the phogenesis of entire organ systems is traced back caenogastropods Ilyanassa and Crepidula that to the cellular level, but such detailed investiga- the mesentoblast 4d likewise may function as tions remain restricted to even fewer species an organizing signaling center (Henry and Perry (Table 7.1). 2008 ; Rabinowitz et al. 2008 ). In general, the macromeres (3A–D, 4A–D) In the fi ve mollusks studied in detail, the fi rst and the micromeres 4a–d give rise to endoderm- quartet micromeres (1a–d) are destined to con- derived structures in mollusks (see Nielsen 2004 ). tribute to the formation of ectodermal structures, These may include (parts of the) larval excretory predominantly in pretrochal domains (Table 7.1 ). systems, the heart, parts of the digestive tract This includes the primary and secondary trocho- including the midgut gland, stomach, and style blasts that form the prototroch and velum, the sac, as well as various sets of muscles (Table 7.1 ). major larval swimming devices. The main larval The mesentoblast 4d, a derivative of the mesen- sensory structure, the apical organ, that is typical toblast mother cell 3D, is the spiralian-specifi c for most lophotrochozoan larvae (Figs. 7.7 and mesoderm founder cell from which a pair of meso- 7.8A, F; see Chapters 6 and 9 ), also derives from dermal bands arises (see Lambert 2010 for brief the 1a–d lineages. In Crepidula , where cell lin- review). However, detailed studies on the caeno- eage has been followed until late- stage veliger lar- gastropod Crepidula have shown that 3D may also vae, it could be shown that a type of sensory cell contribute to endoderm-derived features such as characteristic for lophotrochozoan apical organs, the gut as well as to ectodermal subportions of the the fl ask-shaped receptor cells, as well as the anla- larval kidney complex (Hejnol et al. 2007 ; Lyons gen of the adult cerebral ganglion, likewise derive et al. 2012 ). There is indication that 4d may also from these micromeres (Hejnol et al. 2007 ). The give rise to cells of the early germ line, both in larval eyes form from descendants of 1a and 1c Crepidula (Lyons et al. 2012 ) and in the freshwa- in Crepidula and from 1c progenies alone in ter clam Sphaerium (Ziegler 1885 ; Woods 1931 , Ilyanassa , which also give rise to the right tentacle 1932), but these assumptions need detailed studies in this species (Render 1997 ). In Crepidula , parts on gonadogenesis for defi nite proof. of the mantle likewise stem from the 1a–c lineage, A specifi c role in molluscan development and nerves innervating cerebral structures arise is assigned to the 3D macromere, which is from the 1d cell. Polyplacophoran larvae typically induced by the overlying micromeres to become have ectodermal spicules, and the anterior ones an important signaling (“inductive”) center are derived from the 1a–d lineage. In these larvae and developmental “organizer.” This happens alone, the 1b micromere contributes to the mouth. sometime during the 24–63 cell stages and As we have seen, the fi rst quartet micromeres has been described for the gastropods Patella are exclusively confi ned to ectodermal fates. The and Haliotis as well as the polyplacophoran situation is somewhat different for the second Acanthochiton (van den Biggelaar and Guerrier quartet micromeres (2a–2d), which at least in 1979 ; Arnolds et al. 1983 ; Boring 1989; van scaphopods and gastropods appear to be involved den Biggelaar 1996 ; see also Wanninger et al. in the formation of a specifi c portion of the 2008 ; Nielsen 2012 ). In experimental studies, molluscan mesoderm – the ectomesoderm. In inhibition of 3D induction has resulted in non- polyplacophorans, however, these micromeres 7 Mollusca 117 (continued) Velum, mouth, mantle, Velum, foot, statocyst Velum, Velum, esophagus Velum, Velum, esophagus, Velum, apical organ apical organ Velum, Velum, apical organ, apical organ, Velum, tentacle (only right eye, one, left one unknown) Velum, mouth, mantle, Velum, foot retractor (adult?) mantle Velum, apical organ apical organ Velum, Velum, mantle, statocyst, mantle, statocyst, Velum, heart, mouth, foot heart Mantle, foot ask cell) apical organ Velum, ask cells), cerebral cerebral ask cells), eye, ask cells), head nerves Velum, pedal ganglion, statocyst, pedal ganglion, statocyst, Velum, mantle larval retractor, velum muscle ring) velum retractor, larval muscle ring) organ (fl organ ganglion, mantle, eye organ (fl organ ganglion, mantle Velum, mouth, visceral nerve cords, mouth, visceral nerve Velum, ectoderm, mantle fold, postvelar nerves eye Velum, foot, statocyst, larval kidney kidney larval foot, statocyst, foot, statocyst, Velum, Velum, (incl. fl Velum, mantle, statocyst, heart, mantle, statocyst, Velum, osphradium Velum, foot, statocyst, larval kidney kidney larval foot, statocyst, foot, statocyst, Velum, Velum, Mantle (incl. nerves), foot (incl. Mantle (incl. nerves), sensory cells) Posttrochal ectoderm (foot, mantle fold, shell gland), prototroch Pretrochal ectoderm, prototroch apical Pretrochal ectoderm, velum, Pretrochal ectoderm, prototroch apical Pretrochal ectoderm, velum, fold, shell gland), prototroch, mouth cavity), mouth, telotroch cavity), Pretrochal ectoderm, prototroch (fl apical organ Velum, Pretrochal ectoderm, prototroch Pretrochal ectoderm, apical organ Posttrochal ectoderm (foot, mantle fold, shell gland), prototroch Posttrochal ectoderm (mantle mouth, telotroch cavity), Posttrochal ectoderm (foot, mantle fold, shell gland), telotroch Prototroch, ectomesoderm Apical organ, prototroch Apical organ, prototroch Prototroch Posttrochal ectoderm (foot, mantle Ectoderm Posttrochal ectoderm (mantle Apical organ, prototroch Apical organ, prototroch Prototroch, ectomesoderm Posttrochal ectoderm, foot Posttrochal ectoderm (foot, mantle, shell gland) prototroch, spicules, eye prototroch, spicules, eye spicules, prototroch ectoderm, prototroch mouth spicules, shell gland mouth ectoderm, prototroch, spicules prototroch, spicules, eye prototroch, spicules, eye prototroch, spicules, shell gland shell gland 2a Ectoderm, mouth, 1a Pretrochal ectoderm, mere Contribution Contribution Class Genus Blasto - Polyplacophora mere Chaetopleura Scaphopoda Antalis Patella Gastropoda Crepidula Ilyanassa 1c Apical tuft, pretrochal 2b Ectoderm, prototroch, 3a Ectoderm, mouth Ectoderm Ectomesoderm Esophagus, muscles (incl. main 3b Mouth Ectoderm Ectomesoderm Esophagus, muscles (incl. velum 3c Posttrochal ectoderm, 1b Pretrochal ectoderm, 1d Apical tuft, pretrochal 2c Posttrochal ectoderm, 3d Posttrochal ectoderm, 2d Posttrochal ectoderm, Molluscan cell fates Second quartet micromeres First quartet micromeres Third quartet micromeres Table Table 7.1 118 A. Wanninger and T. Wollesen ), 2007 ), Hejnol et al. ( Mesentoblast, main retractors, gut larval (parts), heart, larval kidney retractor larval retractor larval Style sac, midgut gland, retractor main larval Mesentoblast mother cell Endoderm 2004 ), Nielsen ( 2004 ), Henry et al. ( Mesentoblast, gut, larval kidney, kidney, Mesentoblast, gut, larval retractors, muscles incl. larval mesenchyme, heart, foot tissue, germ cells (?) gland (?) mesentoblast, gut, larval kidney, kidney, mesentoblast, gut, larval retractors, muscles incl. larval mesenchyme, heart, foot tissue, germ cells (?), yolk (?) 1997 ), Dictus and Damen ( 1997 , (endomesoderm) Mesentoblast mother cell Mesentoblast mother cell, 1991 ), Render ( 1976 , Mesentoblast mother cell 1975 , 1974 ) 2012 cell, gut, larval muscles cell, gut, larval ), Lyons et al. ( ), Lyons 4a 4b 4c Endoderm 4d Endoderm Endoderm Mesentoblast Endoderm Endoderm Endoderm Mesentoblast Endoderm Endoderm Mesentoblast, 2 mesodermal bands Endoderm Larval kidney Larval kidney Stomach, style sac Endoderm Endoderm Endoderm mere Contribution Contribution Class Genus Blasto - Polyplacophora mere Chaetopleura Scaphopoda Antalis Patella Gastropoda Crepidula Ilyanassa 3B 3C Gut Gut Endoderm Endoderm Endoderm Endoderm Larval kidney, midgut gland Stomach, style sac, salivary Midgut gland, main 3D Mesentoblast mother 4A 4B 4C Endoderm Endoderm Endoderm Endoderm Endoderm Endoderm Endoderm Endoderm Endoderm Midgut gland Midgut gland Stomach, style sac, salivary gland Endoderm Endoderm 4D Endoderm Endoderm Endoderm Yolk Yolk, midgut gland 2009 (continued) Macromeres 3A Gut Endoderm Endoderm Larval kidney, midgut gland Midgut gland, main Fourth Fourth quartet micromeres Data from van Dongen and Geilenkirchen ( Data from van Table 7.1 Table Goulding ( 7 Mollusca 119

A BC

D EF

GHI

Fig. 7.7 Developmental fate of the molluscan apical additional cells of a putative lateral sense organ (lso ) are organ. (A – C) and ( F) are scanning electron micrographs, interconnected with the fl ask cells at their base; dorsal ( D) and ( G – I) are confocal projections of neural subsets view. ( E) Schematic representation of the apical organ stained with an anti-serotonin antibody. All images are of the polyplacophoran Ischnochiton hakodadensis in from the scaphopod Antalis entalis except for ( E ), which dorsal view showing eight fl ask cells ( black ), four periph- is from the polyplacophoran Ischnochiton hakodadensis eral nonsensory cells, and two additional bipolar neurons (= Stenoplax heathiana). Scale bars are 50 μm in ( A – C ) (white cell bodies). ( F ) Detail of the prototroch of Antalis and ( H ), 25 μm in (D ), and 10 μm in (F , G , I ). ( A ) Lateral showing bundles of compound cilia. ( G ) Detail of a fl ask view of the anterior region of a trochophore larva with cell of the apical organ. (H ) Degenerating apical organ. prototroch (pt ) and well-developed apical tuft (at ) that is The fl ask cells have migrated to the area of the future cere- part of the apical organ. (B ) Anterior view of a late larva bral ganglia, i.e., behind the prototroch; the fi rst cells of at the onset of degeneration of the apical tuft. (C ) Anterior the cerebral ganglia have already formed (cg ). ( I ) Detail view of a larva close to metamorphic competence with of ( H) showing the degenerating fl ask cells (asterisks ) of almost fully degenerated apical tuft. (D ) Four fl ask-shaped the apical organ and the fi rst immunoreactive cells of the serotonin-positive cells ( asterisks ) of the apical organ, as cerebral ganglia. In Antalis entalis , the apical organ is lost typical for many molluscan larvae. In Antalis larvae, two prior to metamorphic competence 120 A. Wanninger and T. Wollesen

AB

C D

EF

Fig. 7.8 Molluscan larval neuromuscular anatomy. Cell ( vvr1 , vvr2 ) and an accessory larval retractor (alr ), as well nuclei are stained in blue (HOECHST; lacking in B , D ) as the anlagen of future adult muscles such as the anterior and musculature is labeled green (phalloidin, F-actin). ( aa ) and posterior adductor muscles (pa ), the foot retrac- Serotonin-like immunoreactive (serotonin-LIR) nervous tor (fr ), the digestive system (dm ), and the mantle retrac- elements are stained in red and cilia (acetylated α-tubulin) tors (Image courtesy of Marlene Karelly). ( D ) The are labeled in turquoise in ( F). All images are confocal serotonin-LIR nervous system and the musculature of the projections with anterior facing up (except D where it veliger larva of the gastropod Aplysia californica (dorsal faces down). All scale bars are 50 μm. ( A ) The serotonin- view, anterior faces down). The muscle system comprises LIR nervous system of the late trochophore of the chiton musculature associated with the propodium ( pp ), metapo- Acanthochitona crinita is composed of fl ask-shaped cells dium ( mp), and velum ( v). Further muscles are the mas- ( arrowheads) in the apical organ, additional sensory cells sive larval retractor muscle (lrm ), transversal muscle ( arrows), and various other cell somata which are con- fi bers (tmf ), and an accessory retractor muscle ( arm ). The nected via neurites ( double arrowheads ) to the apical larva exhibits serotonin-LIR innervation of the velum, organ or the cerebral commissure (dorsal view). (B ) propodium, metapodium, and visceral sac ( vs ). Serotonin- Dorsal view of the larval serotonin-LIR nervous system of LIR cells are present in the apical organ and in the anlagen the neomeniomorph Gymnomenia pellucida showing of the cerebral ganglia (cg ). ( E) The complex larval mus- weak staining in the apical organ (ao ) and lateral (lc ) and culature of the patellogastropod Lottia is, among others, pedal nerve cords ( pc), with a serotonin-LIR cluster of composed of a larval retractor muscle ( lrm) that connects cells connected to the latter (arrowheads ). A dense median to the velar ring musculature (vrm ). Lateral left view neural plexus ( mp) is visible along the longitudinal axis (Image courtesy of Alen Kristof). ( F) The apical organ (Image courtesy of Emanuel Redl). ( C) The complex ( ao) of this larva of the patellogastropod Lottia houses musculature of the veliger larva of the protobranch bivalve three serotonin-LIR fl ask-shaped cells. Lateral left view Kurtiella bidentata comprises larval retractor systems (Image courtesy of Alen Kristof). Further abbreviations: including dorsal ( dvr1 , dvr2 ) and ventral velum retractors apical tuft (at ), foot (f ) 7 Mollusca 121 contribute to the (posttrochal) ectoderm. In addi- sodermal regions in Patella and to esophageal tion, 2a, 2b, and 2c form trochoblasts. Spicule- and velar cells (including muscles) in Crepidula forming cells derive from the 2a–c lineage but not and Ilyanassa . 3c and 3d contribute to posttrochal from 2d, which instead contributes to parts of the ectodermal domains including mouth and telo- seven shell fi elds of the larva (Henry et al. 2004 ). troch in Patella and to the velum, foot, and stato- Interestingly, the larval ocelli, which, in contrast cyst in Crepidula and Ilyanassa . In Crepidula , to the pretrochally positioned eyes of other spira- both micromeres also contribute to various cell lian larvae, are situated ventrolaterally behind the types associated with the larval kidney. prototroch in polyplacophorans, derive from the Taken together, the overall cell lineage 2a–c cells. They thus have a different develop- patterns appear to be rather conserved among mental history than other spiralian larval eyes, the various polyplacophoran, scaphopod, and which emerge from the 1a–c lineage (Henry et al. gastropod mollusks investigated, at least with 2004). This different ontogenetic history may respect to their gross morphological formation argue against homology of polyplacophoran and domains. Exceptions do occur, however, as other spiralian larval eyes and should be investi- manifested in the complex genealogy of the gated further by comparative lineage-tracing and prototroch in Patella or the diverging lineage gene expression studies. history of the polyplacophoran larval eyes. For scaphopods, information on the fate of the Unfortunately, detailed cell lineage studies in second quartet micromeres is scarce. The 2a–c particular of neural and other internal features lineages contribute to the three-rowed prototroch such as the various larval and adult muscle sys- and 2a and 2c in addition to the ectomesoderm, tems are still scarce. Even for key gross mor- while 2d forms posttrochal ectodermal structures phological features, such as the various of the foot, mantle, and shell gland. In Patella , prototrochal, metatrochal, and velar structures the 2a–d lineages are rather similar, with a 2d as well as the apical organ, more comparative contribution to the telotroch (Dictus and Damen data are needed and – in particular in combina- 1997 ). The situation in the two other gastropods tion with gene expression studies – should pro- investigated, Crepidula and Ilyanassa , mostly vide a powerful tool for higher-level taxon corresponds to Patella , with a few additional homology assessments across the Mollusca. contribution domains identifi ed in the two caeno- gastropods, such as nervous and sensory struc- tures including the statocyst (2a and 2c in both Gastrulation and Mesoderm species), visceral nerve cords and eye nerves Formation (2b in Crepidula ), sensory cells of the foot (2d in Crepidula), heart (2c in both), osphradium (2c In mollusks with small eggs, gastrulation occurs in Crepidula ), and the foot retractor (i.e., shell by invagination on the vegetal pole and subse- muscle) (2b in Ilyanassa ) (Table 7.1 ). An early quent formation of a fl uid-fi lled coeloblastula. work on bivalve cell fates suggests that cells Species with large, yolk-rich eggs usually exhibit from the 2d lineage also contribute to the shell epiboly and a massive sterroblastula. The macro- fi elds in Dreissena polymorpha (Meisenheimer meres are the fi rst cells to invaginate, followed by 1901 ), a fi nding that was corroborated by a recent the fourth quartet micromeres. During subse- study on the Japanese spiny oyster Saccostrea quent development, the blastopore often narrows kegaki (Kin et al. 2009 ). In the polyplacophoran considerably and may even close in some spe- Chaetopleura and the scaphopod Antalis , the third cies. In most cases, however, it moves in anterior quartet micromeres give rise to various (posttro- direction and fi nally comes to lie behind the pro- chal) ectodermal domains. In Chaetopleura, 3c totroch, where it develops into the defi nite mouth and 3d successors are found in spicule- and shell (Figs. 7.3 and 7.4A ; see Nielsen 2004 , 2012 ). In gland-forming cells. In the gastropods, 3a and 3d the large majority of species, the anal opening contribute to not precisely characterized ectome- forms de novo. However, earlier reports on a 122 A. Wanninger and T. Wollesen

“deuterostomous” condition in the river snail LATE DEVELOPMENT Viviparus, where the blastopore was found to remain in the posterior region where it forms the Diversity of Molluscan Larval anus, while the mouth arises secondarily (Dautert Development 1929 ; Fernando 1931; Fioroni 1979 ), indicate that mouth/anus formation in mollusks may be Mollusks exhibit a variety of postembryonic onto- rather plastic (see Nielsen 2004 ). genetic pathways. Indirect development via a As mentioned above, the molluscan meso- trochophore-like larva is found in all recent classes derm is derived from two embryonic sources, except the cephalopods (potential monoplacopho- the ectomesoderm (in gastropods mostly formed ran larvae remain to be described) and most likely by the third quartet micromeres with minor constitutes the basal condition. Even some terres- contributions from second quartet derivatives; trial gastropods with intracapsular development see Table 7.1 ) and the endomesoderm (derived show rudimentary larvae that undergo metamor- from progenies of the 4d mesentoblast, which phosis (e.g., the mouse ear snail Myosotella ) and divides and forms one left and one right meso- several so-called direct developers show larval dermal band). However, if at all, these bands rudiments such as vestigial prototrochal or velar appear only vaguely visible in most representa- cells (e.g., the great pond snail Lymnaea stagna- tives (Wierzejski 1905; Okada 1939 ; Hinman lis ). In the cephalopods, early blastomere forma- and Degnan 2002 ), and since both ectodermal tion is followed by the embryo overgrowing the and endomesodermal anlagen soon coalesce, large yolk mass, eventually forming a digestive it remains diffi cult to unequivocally assess the tube that is connected to the maternal yolk supply degree of contribution of these individual meso- for nutrient uptake. This leads to continued growth dermal sources to respective organs. Rather, it and differentiation of the embryo which, although appears as if all mesoderm-derived organs have often only millimeter-sized, is equipped with all both ectomesodermal and endomesodermal con- major organs to commence its life as active preda- tributors (Table 7.1 ). tor already at the hatchling stage. In cephalopods, blastomeres and blastocones In indirect developing mollusks, larval life typi- form the cytoplasmic cap of the early discoblas- cally starts with a lecithotrophic (nonfeeding) tula (Boletzky et al. 2006 ). While all blastomeres trochophore-like stage, but some species enter the are located in the center of the cytoplasmic cap, pelagic realm already as ciliated gastrulae. The blastocones surround the latter (Fig. 7.5). trochophore larva is characterized by its name-giv- Blastocones are ray-shaped and part of a ing prototroch and an apical sensory organ that syncytium with the uncleaved portion of the often exhibits a characteristic tuft of long cilia zygote (Boletzky 1989 ). During gastrulation, (Figs. 7.3 and 7.4). The prototroch (formed by shearing movements lead to the migration of the rows of trochoblasts, see above and Table 7.1) con- last two outermost blastomere rows below the stitutes the major swimming device that propels inner blastomeres. These last two outermost the larva through the water column, with the ani- blastomere rows form the mesendoderm, and the mal rotating clockwise around its longitudinal axis. remaining innermost blastomeres of the cyto- It may comprise one to three rows of compound plasmic cap remain as ectoderm. In line with this cilia as well as cells with accessory, smaller cilia. process, the equivalent to the blastopore lip is The prototroch subdivides the molluscan larva into extra-embryonic ectoderm which grows over the the pretrochal episphere and the posttrochal hypo- yolk syncytium in direction of the vegetal pole sphere (Fig. 7.3 ). From the episphere, the anlagen (Boletzky 1989 ). Subsequently, various organ of major adult anterior structures such as the systems such as the anlagen of the ganglia or the cephalic nervous system and various sensory arms become discrete. organs usually form already prior to metamorpho- sis. The mouth usually comes to lie in the anterior- 7 Mollusca 123 most region of the hyposphere, immediately trochophore to early veliger stage (Wanninger adjacent to the prototroch. The hyposphere often et al. 2000 ). elongates during larval development and gives rise Larval settlement is often preceded by active to the visceral body region (Wanninger et al. 1999a ; probing of the substrate with the apical tuft or the Okusu 2002 ; Wanninger and Haszprunar 2002a , b ; foot. Subsequently, metamorphosis is initiated Todt and Wanninger 2010 ). It may develop spic- and involves shedding of the apical cap (neome- ules (in neomeniomorphs, chaetodermomorphs, niomorphs), test cells (protobranch bivalves), and and polyplacophorans), shell plates (polyplacoph- prototroch/velum (polyplacophorans, gastropods, orans), bivalved shells (Bivalvia), or univalved non-protobranch bivalves, scaphopods). The api- shells (monoplacophorans, gastropods, scaph- cal organ and the larval operculum (in gastropods) opods). The larval hyposphere may bear additional are lost, and the adult shell – the teleoconch – starts ciliary rings such as a terminal telotroch (Fig. 7.4B, to form in many conchiferan taxa (see below). At C). The larvae of some gastropods (e.g., Patella ) this stage, the transition from planktonic larval show a transitory tuft of “anal cilia” in the region of life to the juvenile, often benthic, lifestyle takes the anus, and these are sometimes homologized place in most mollusks. During this period larval with the aplacophoran telotroch. Protobranch as well as adult excretory systems may coexist bivalves (e.g., Nucula, Acila) exhibit a so-called alongside each other, and cases where the larval test cell larva, whereby cells with multiple rows of protonephridia are transformed into adult metane- cilia cover the developing juvenile underneath phridia have been described. (Zardus and Morse 1998 ), and neomeniomorph Despite their direct mode of development, larvae produce an “apical cap” or “calymma” that some octopods drift passively in the water col- covers the anterior larval region (Fig. 7.4B ; Okusu umn after hatching and are therefore often 2002 ; Todt and Wanninger 2010 ). referred to as “paralarvae.” These paralarvae Protonephridia have been found in the larvae of often have specifi c, transitory adaptations linked all molluscan classes investigated. Accordingly, to their temporary planktonic life, such as a trans- these excretory organs together with the prototroch parent musculature and a less complex chromato- and apical organ are considered as part of the mol- phore system (Villanueva and Norman 2008 ). In luscan larval ground plan. In larvae of polypla- many periodically planktonic mollusks, settle- cophorans as well as some gastropods and a few ment goes hand in hand with signifi cant behav- bivalves, eyes are common. ioral changes such as loss of positive phototaxis Following the trochophore stage, many or gain of positive geotaxis. gastropods and bivalves develop a secondary larva, the veliger. The name-giving velum is con- sidered an elaborated prototroch and may form Neurogenesis multiple lobes used both for swimming and feed- ing, enabling month-long planktonic life for The development of the gastropod and cepha- numerous species (Fig. 7.4E). In gastropod veli- lopod nervous system has traditionally received gers, an operculum for closure of the embryonic considerable attention. Thus, a signifi cant num- shell is often produced. At this stage, a strong ber of studies, mainly concerned with the analysis tendency towards heterochrony is usually of gangliogenesis based on histological sections, observed, where anlagen of a number of adult exist (e.g., Kölliker 1844 ; Faussek 1901 ; Martin body plan features are already formed in the veli- 1965 ; Raven 1966 ; Meister 1972 ; Demian and ger larva, including the heart, shell muscles, neu- Yousif 1975 ; Kriegstein 1977 ; Marquis 1989 ; romuscular and buccal features, osphradia, Page 1992a , b ; Lin and Leise 1996 ; Page and statocysts, tentacles, and other sensory organs. Parries 2001 ; Shigeno et al. 2001 ). In the gastro- Torsion, the developmental process defi ning the pods, the ganglia form already in the larva from Gastropoda, occurs at the transition of the late invaginating or delaminating ectodermal cells 124 A. Wanninger and T. Wollesen

(Raven 1966 ; Kandel et al. 1981 ; Mescheryakov representatives of all molluscan classes except 1990 ; see Croll 2000 for brief review). Thereby, for Chaetodermomorpha and Monoplacophora. the cerebral and pedal ganglia form prior to the As a common trait, the adult molluscan nervous visceral ganglia (Page 1992a , b; Lin and Leise system starts to form in early developmental 1996), and the buccal ganglia appear to be the stages. The tetraneurous condition can be recog- last set of major ganglia to develop (Cumin nized in larvae of all clades, together with develop- 1972). After establishment of the ganglia, neuro- ing pedal commissures and the anlage of the nal precursor cells proliferate from the region of cerebral commissure or ganglia (where present). gangliogenesis (Jacob 1984 ). Despite the often These neural subsets do not form a strictly linear detailed descriptions of the process of ganglion anterior-posterior formation gradient as in many formation, histology-based investigations largely annelids, thus refl ecting the non-segmental failed to discover details of early molluscan neu- ancestry of Mollusca (Friedrich et al. 2002 ; rodevelopmental processes, for example, the for- Voronezhskaya et al. 2002 ; Wanninger and mation of individual neurons, particularly in early Haszprunar 2003 ). In conchiferans, the anlagen of developmental stages. Accordingly, comparative the pedal, visceral, pleural, and other major gan- molluscan (and generally invertebrate) neurode- glia are usually recognized by immunocytochemi- velopmental studies employing modern optical, cal staining for serotonin and FMRFamide (Marois computational, and imaging tools (immunofl uo- and Carew 1990 , 1997a , b ; Marois and Croll 1992 ; rescence labeling and confocal microscopy in Voronezhskaya and Elekes 1993 , 2003 ; Croll and combination with powerful 3D reconstruction Voronezhskaya 1996 ; Diefenbach et al. 1998 ; software) have signifi cantly altered our under- Croll 2000; Hinman et al. 2003 ; Wanninger and standing concerning the timing and morphoge- Haszprunar 2003 ; Croll and Dickinson 2004 ; netic details underlying neurogenesis. By the Voronezhskaya et al. 2008 ; Wollesen et al. 2009 , use of these techniques, a solid body of data on 2010 , 2012 ; Kristof and Klussmann-Kolb 2010 ). the distribution of neuroactive substances during The molecular mechanisms that govern mol- molluscan neurogenesis has become available. luscan neurogenesis are still poorly understood, However, a pan-neural marker for molluscan (and but the few detailed data on two vetigastropods lophotrochozoan) developing nervous systems ( Haliotis and Gibbula) and one cephalopod (the still awaits discovery; the widely used tubulin decapod squid Euprymna ) demonstrate involve- markers or antibodies directed against horse- ment of a number of Hox and potentially also radish peroxidase, which render comprehensive ParaHox genes in this process, although loss of results for the study of, e.g., arthropods (see Vol. function of individual genes has also been 3, Chapters 3 and 4 ; Vols. 4 and 5), do not work demonstrated (O’Brien and Degnan 2002a , b , well on early developmental stages in mollusks, 2003 ; Hinman et al. 2003 ; Lee et al. 2003 ; Samadi because they often fail to recognize existing neu- and Steiner 2009 , 2010a , b ). Cell lineage studies ral subsets and/or because signifi cant fractions based on blastomere injection have shown that of the developing nervous system are obscured the cerebral ganglia are derived from micromeres by the massive presence of cilia (Fig. 7.8F ). 1a–c, while the pedal ganglia form from the 2a Nevertheless, neurodevelopmental work dur- and the visceral nerve cords from the 2b lineage ing the past two decades has shown that some in the caenogastropod Crepidula (Table 7.1 ; commercially available antibodies, especially Hejnol et al. 2007 ). Due to the lack of compara- against the widely distributed neurotransmitter tive data, it is yet impossible to assess the degree serotonin (5-hydroxytryptamine, 5-HT) as well of conservation of nervous system cell lineage as FMRFamide-related peptides, do label signifi - across the Mollusca. cant portions of the developing molluscan ner- The most striking and characteristic neural vous system and thus provide important insights component in molluscan larvae is the apical into comparative molluscan neurogenesis. As organ, a common feature shared with the vast such, data on neurogenesis are now available for majority of lophotrochozoan species (Figs. 7.7 7 Mollusca 125 and 7.8 ; see also Chapters 6 , 7 , 8 , 9 , 10 , 11 , and 12 ; that drastically deviates from the simple condi- Bonar 1978 ; Wanninger 2009 ). This organ is usu- tion found in prosobranch gastropods, bivalves, ally the fi rst neural structure that develops in mol- and scaphopods (Fig. 7.7 ; Croll and Dickinson luscan larvae. Some terminological confusion 2004 ). Apart from the above-mentioned gastro- exists because this system has also been referred pods, such a complex apical organ is also pres- to as “apical ganglion” (although it does not form ent in polyplacophoran larvae (Friedrich et al. a ganglionic structure). To make matters worse, 2002 ; Voronezhskaya et al. 2002 ) and in the “apical ganglion” has also been applied to the creeping larva of entoprocts (Chapter 6 ; entire anterior complex that comprises the larval Wanninger et al. 2007 ). This and other shared apical organ and the developing adult cerebral features strongly argue in favor of a mollusk- commissure (Nielsen 2012 ). In order to strictly entoproct clade (Tetraneuralia), implying that a separate these ontogenetically and structurally complex apical organ is plesiomorphic for different neural features, “apical organ” should Mollusca. only be applied to the larval anterior sense organ Recent gene expression studies on the vetigas- (cf. Richter et al. 2010 ). tropod Gibbula varia have shown expression of From an immunocytochemical perspective, the Hox genes Lox2 , Lox4, and Lox5 as well as the molluscan apical organ is relatively simple the ParaHox gene Gsx in cells of the apical organ organized, although detailed three-dimensional (Samadi and Steiner 2010a , b ). The cell lineage of reconstructions employing ultrathin serial sec- conchiferan and polyplacophoran apical organs tioning and transmission electron microscopy appears to be conserved to a certain degree, since are lacking. Thus, comparisons largely rely on data on the two caenogastropods Ilyanassa and immunolabeling data alone. These show that the Crepidula as well as the scaphopod Antalis have apical organ is composed of a given number of consistently shown its origin from all four fi rst serotonin-like immunoreactive (serotonin-LIR) quartet micromeres (Table 7.1 ; van Dongen and and sometimes also FMRFamide-like immuno- Geilenkirchen 1974 , 1975 , 1976 ; Hejnol et al. reactive (FMRFa-LIR) fl ask-shaped cells 2007 ; Goulding 2009 ). (sometimes called ampullary cells), from which While immunoreactivity of the cells of the at least some of the compound cilia that form apical organ is lost at metamorphosis, their defi - the apical tuft emerge (Figs. 7.7 and 7.8A ; nite fate remains largely unknown. Most authori- Wanninger 2009 ; Richter et al. 2010 ). The num- ties probably agree that its cells are resorbed. ber of these fl ask-shaped serotonin-LIR cells This has been shown for the gastropod Ilyanassa typically ranges between two and four, and obsoleta by applying markers for apoptotic cells these are often the only immunoreactive com- (Gifondorwa and Leise 2006 ). However, incorpo- pounds that have been identifi ed in the apical ration of at least some individual cells into the organ of basal gastropods as well as bivalves adult nervous or sensory system seems possible. and scaphopods (Page 2002, 2006; Hinman Postmetamorphic individuals of the nudibranch et al. 2003; Wanninger and Haszprunar 2003 ; Phestilla sibogae appear to maintain cells of the Voronezhskaya et al. 2008 ). An apical neuropil apical organ; their eventual fate, however, is is situated immediately posterior to the fl ask unknown (Bonar 1978 ). cells, and at its base the adult cerebral commis- In most indirect developing mollusks, the sure forms (Croll and Dickinson 2004 ). In some cerebral commissure forms at the base of the caeno- and heterobranch gastropods, additional apical organ. This has fueled speculations that sets of (ciliated or non-ciliated) non-fl ask-like the apical organ plays an inductive role in the parampullary (or peripheral) cells are found. formation of the adult central nervous system. These surround the fl ask cells at their base. In Apart from that, detecting (chemical) settle- taxa that show such peripheral cells, the number ment cues has been attributed as a prime role to of fl ask cells is usually increased, resulting in a the apical organ, but experimental data to this complex cellular architecture of the apical organ end are few (Hadfi eld et al. 2000 ). The loss of 126 A. Wanninger and T. Wollesen immunochemical signal in the fl ask cells of lar- and in the juvenile and adult nervous system of vae of the scaphopod Antalis prior to metamor- gastropods and cephalopods (Barlow and phic competence indicates that the apical organ Truman 1992 ; Kempf et al. 1992 ; Kanda and may not be necessary for successful settlement Minakata 2006 ; Ellis and Kempf 2011 ). With and metamorphosis in all mollusks (Wanninger antibodies against these substances available, and Haszprunar 2003 ). and several other neuroactive compounds such Additional larval neural subsets in mollusks as nitric oxide and vasopressin identifi ed include a serotonin-LIR nerve underlying the (Baratte and Bonnaud 2009; Filla et al. 2009 ; prototroch and velum, again a feature shared Bardou et al. 2010 ; Mattiello et al. 2012 ), poten- with other lophotrochozoans (Wanninger 2009 ). tially useful tools are at hand for future compar- The velum is usually heavily innervated by ative evolutionary neurodevelopmental studies serotonin- and FMRFa-LIR neurites that emerge on mollusks. from the apical organ and/or the cerebral com- missure and disappear at metamorphosis (Dickinson et al. 1999 ; Dickinson and Croll Myogenesis 2003 ; Wollesen et al. 2007 ; see Croll 2009 for review). An additional pretrochal sense organ Mollusks are highly muscular animals with comprising two pairs of dorsolateral and two several distinct muscular subsets, including pairs of ventrolateral ampullary cells with a cili- dorso ventral muscles (often called “shell mus- ated lumen that stain positive for FMRFamide cles” or “pedal retractors”), buccal muscles to and, less so, for serotonin has been found in support the radula apparatus, mantle retrac- polyplacophoran trochophores. These cells are tors, as well as more taxon-restricted systems connected to the cerebral commissure. The such as enrolling muscles (in aculiferans), entire ampullary system is lost at metamorpho- head retractors (in scaphopods, gastropods, and sis (Haszprunar et al. 2002 ). cephalopods), a dorsal rectus muscle (in poly- The increased interest in gastropods as neu- placophorans and larval neomeniomorph apla- robiological model species, which peaked in cophorans), and various adductors in bivalved Nobel prize-winning studies on the giant neu- re presentatives (Fig. 7.9 ; reviewed in Haszprunar rons of the sea hare Aplysia (see Kandel 2001 ), and Wanninger 2000 ; see also Scherholz et al. has led to screenings of gastropods and cephalo- 2013 ). The dorsoventral muscles are arranged pods for neuroactive substances. Thereby, a as multiple, serially arranged fi ne bundles along variety of compounds were identifi ed in larval the entire anterior- posterior body axis in neo- stages of the heterobranch gastropod Phestilla meniomorphs. In the chaetodermomorphs, this sibogae , including dopamine, choline acetyl- arrangement is restricted to the anterior region. transferase, and norepinephrine (Kempf et al. Polyplacophorans have eight sets of highly com- 1992), and an even more impressive suite in plicated shell muscles, correlating with the eight cephalopods (Messenger 1996 ). A so-called shell plates. Such a condition is also present in VD1/RPD2 alpha-neuropeptide, isolated from the uni-shelled monoplacophorans, which has led the pulmonate gastropod Lymnaea stagnalis , to heavy speculations concerning a segmented was found to be present in specifi c nerve cells in ancestry of mollusks (Lemche and Wingstrand the central nervous system of this snail as well 1959 ). Everything from one to eight dorsoventral as in other conchiferans (Kerkhoven et al. 1992 , muscle pairs (“foot retractors”) is found among 1993 ; Wollesen et al. 2012 ) and may be con- bivalves, one or two pairs are present in scaph- served among the entire Mollusca. Small cardio- opods, and a single pair is typical for gastropods active peptides (ScPs) are also expressed in (often referred to as “spindle muscle” in snails neural subsets of gastropod and bivalve larvae with a coiled shell) and cephalopods. 7 Mollusca 127

A BC

DE

Fig. 7.9 Neuromuscular anatomy of juvenile scaphopods in ( A ) which highlights the captacula (arrowheads ). (C ) and embryonic and hatched cephalopods. Cell nuclei are This prehatching cephalopod Idiosepius notoides speci- stained in blue (HOECHST) and musculature and neuropil men exhibits muscles of the developing arms (a ), the man- are labeled green (phalloidin, F-actin). Specifi c neural ele- tle ( m ), and the cephalic region, as well as the neuropil of ments are stained red (C –E ) or red and turquoise (D ), the brain (b ) and optic lobes ( ol). Note the FMRFamide- respectively. All images are confocal projections with LIR neurons (arrowheads ) of the posterior subesophageal anterior (i.e., adult ventral) facing up. Scale bars are mass. Anterior view. (D ) Epidermal nerve net (arrow ) in 100 μm in ( A –C ) and 200 μm in ( D, E ). (A ) Lateral right the cephalic and mantle region of the hatching epipelagic view of an early juvenile scaphopod (Antalis entalis). octopod Argonauta hians. Anterior view. Note the sero- Prominent muscles are the cephalic retractor ( cr ), mantle tonin-LIR cell somata in the brain (arrowheads ). (E ) retractor (mr ), and pedal retractor (pr ). Further muscles Horizontal vibratome section through the head of a pre- are muscles of the pedal plexus ( pp), those associated with hatching bobtail squid (Euprymna scolopes ) showing the remaining foot (f ), the mantle fold (mf ), and a hitherto musculature and neuropil (green ) as well as FMRFamide- undescribed muscle inserting close to the pedal retractor LIR elements (red ). Note the prominent musculature of which runs in posterior direction (arrowhead ). (B ) the arms (a ), buccal mass (bm ), mantle (m ), and suckers Magnifi cation of the ventral region of the specimen shown ( s). Further abbreviations: eye (e ), internal yolk duct (iy ) 128 A. Wanninger and T. Wollesen

Myogenesis in Aculiferans chronous appearance of the fi rst (seven) pairs of The aplacophorans exhibit a typical muscular dorsoventral myocytes in neomeniomorphs and tube involving prominent longitudinal muscle polyplacophorans is in stark contrast to the ante- bundles which are overlain by circular muscles. rior-posterior formation gradient in annelids, which Additional interspersed oblique or helical mus- is due to a posterior growth zone (Chapter 9 ). These cles are present in some taxa. Such a muscular fundamental differences during myogenesis cor- body wall is absent in non-vermiform mollusks, roborate all other developmental data and illustrate probably due to the evolution of shells as major the non-segmented ancestry of Mollusca. skeletal elements which, together with a pro- Neomeniomorph evolution from a nounced dorsoventral musculature, provide body polyplacophoran-like ancestor is further supported stability. Interestingly, such a meshwork was also by additional features of neomeniomorph larval found in the anterior region of the polyplacopho- myoanatomy, such as the presence of a rectus ran-like larva and was interpreted as an evolu- muscle as well as a pair of lateral enrolling mus- tionary rudiment of an ancestral body wall cles, typical for adult chitons (Fig. 7.10 ; Scherholz musculature (Fig. 7.10 ; Wanninger and et al. 2013 ). However, and in contrast to the poly- Haszprunar 2002a ). Given the similarities with placophorans, both systems are considerably other worm-like spiralians, the presence of such a remodeled during neomeniomorph metamorpho- muscular tube was thus considered as potentially sis and are incorporated into the longitudinal body ancestral (plesiomorphic) for Mollusca. wall musculature of the juvenile. Accordingly, However, recent comparative developmental neomeniomorph gross anatomy as a simple, cylin- studies on polyplacophoran and neomeniomorph drical, shell-less worm is now considered a sec- myogenesis have rejected this view. The data ondary simplifi cation that evolved from a more obtained demonstrated striking similarities in both complex, polyplacophoran-like ancestor. The clades, revealing muscle systems confi ned to the transversal musculature underlying the polypla- larvae of neomeniomorphs and polyplacophorans cophoran shell plates can be interpreted as concen- alone, including a single ventromedian and a pair trated and modifi ed units of ring muscles that were of ventrolateral muscles (Fig. 7.10 ; Scherholz et al. present in the larva of the last common aculiferan 2013 ). Furthermore, it was shown that the neome- ancestor, similar to the apical muscular grid in the niomorph Wirenia argentea develops seven pairs of chiton larva ( Wanninger and Haszprunar 2002a ). dorsoventral muscles at fi rst. These form simulta- neously and precede the subsequent addition of Myogenesis in Conchiferans multiple muscles along the anterior-posterior axis Myogenesis in the Conchifera follows a much (Scherholz et al. 2013 ). This is particularly interest- simpler pattern than observed in the neomenio- ing because polyplacophorans likewise pass morphs and polyplacophorans. Most adult mus- through a stage with seven pairs of shell muscles cle systems develop more or less directly and which form from multiple sets of serial myocytes without major remodeling from precursors by secondary concentration after metamorphosis already formed in the larva. One of the few mus- (Fig. 7.10A, C ; Wanninger and Haszprunar 2002a ; cles strictly confi ned to the larval stage is the pro- Scherholz et al. 2013 ). Both the additional multiple totroch/velum muscle ring (Figs. 7.8E and 7.10 ). pairs of dorsoventral muscles in the neomenio- This muscle underlies the prototroch/velum and morph and the eighth set in polyplacophorans is present in all molluscan classes (including the develop considerably later. This, together with data aculiferans), except for the scapho-pods and on aculiferan skeletogenesis (see below) and the cephalopods (monoplacophoran condition fi nding of a fossil seven-shelled aplacophoran unknown), and thus most likely belong to the (Sutton et al. 2012 ), argues for a last common molluscan larval ground plan (Wanninger and ancestor of neomeniomorphs and polyplacopho- Haszprunar 2002b ; Nielsen et al. 2007 ; rans with a seven-fold seriality of the dorsoventral Wanninger et al. 2008 ; Dyachuk and Odintsova musculature and, probably, shell plates. The syn- 2009 ; Scherholz et al. 2013 ). 7 Mollusca 129

ADULT LARVA

Polyplacophora A PolyplacophoraB Neomeniomorpha C Polyplacophora D Neomeniomorpha

E F G H

Neomeniomorpha

IJKL

Ventromedian muscle Ventrolateral muscle Dorsoventral muscle Prototroch muscle ring Transversal muscle Ring muscle Mouth Rectus muscle Pedal pit Enrolling muscle

Fig. 7.10 Comparative myogenesis in polyplacophorans cle, advanced sets of dorsoventral muscles, transverse and neomeniomorph aplacophorans. A , B , E , F , I , and muscles, and a lateral enrolling muscle. Note also the J are 3D reconstructions based on confocal projections striking similarity in the larval myoarchitecture of both of specimens stained with phalloidin to label the muscu- taxa as depicted in (A –L ), including numerous subsets lature. ( A –D ) Dorsal muscles seen from ventral. (E –H ) that are lost during neomeniomorph metamorphosis in the Ventral muscles seen from dorsal. (I , J) . Ventrolateral process of secondary simplifi cation, including the rectus, right views. (K , L ) Cross sections. Scale bars are 20 μm. ventromedian, ventrolateral, and enrolling muscle. The Far left column shows adult myoanatomy of polypla- latter two are postmetamorphically remodeled and con- cophorans and the neomeniomorph Wirenia argentea in tribute to the longitudinal body wall musculature of the cross section (top ) and dorsal view (bottom ). Note simple adult. Note also the rudimentary body wall ring muscle in arrangement in the neomenimorph with body wall mus- the anterior region of the polyplacophoran larva (I ). See cles (here only represented by ring muscles) and serial color code for identifi cation of individual muscle sets and dorsoventral musculature, while polyplacophorans have a text for details (From Scherholz et al. (2013 ) ) much more complex musculature involving a rectus mus- 130 A. Wanninger and T. Wollesen

Distinct sets of larval retractor systems occur Myogenesis in bivalves has so far been inves- in gastropods and bivalves. These are resorbed tigated to a surprisingly little extent, but the few during metamorphosis and do not contribute to data available suggest the presence of highly com- the adult dorsoventral musculature. In gastro- plex retractor systems in their veliger larvae, also in pods, usually two asymmetrically positioned lar- certain semi-direct developing (brooding) species val retractors are present (Wanninger et al. 1999a , (Meisenheimer 1901 ; Altnöder and Haszprunar b ; Wollesen et al. 2008 ; Kristof and Klussmann- 2008 ; Wanninger et al. 2008 ; Dyachuk and Kolb 2010 ). The main (or velum) retractor pro- Odintsova 2009 ). An additional ventral larval retrac- jects into the anterior region, where it often inserts tor system is present in Dreissena, Mytilus, Pecten , at the velum muscle ring or other velar tissues. and Lyrodus (shipworm) larvae (Meisenheimer The second larval retractor muscle is considerably 1901 ; Cragg 1985 ; Dyachuk and Odintsova 2009 ; weaker and projects into the mantle. It is there- Wurzinger-Mayer et al. 2014 ), but their almost fore usually termed “mantle retractor” or “acces- opposite projection relative to the gastropod acces- sory larval retractor.” Due to their asymmetrically sory (mantle) retractor argues against homology of positioned insertion sites on the embryonic shell these muscles. Similar to the gastropods, all larval and their different direction of projection (into the retractors are lost prior to or during metamorphosis. mantle versus the prototroch or velum), it has long In bivalves, the anlagen of the various adult shell been speculated that their activity may play a role muscles (the dorsoventral or foot retractors, adduc- in ontogenetic torsion, especially since this gas- tors, and mantle retractors that later form the pallial tropod key invention is completed very quickly in line) already appear functional in the veliger larva basal representatives (for in-depth discussion see and form independently from the larval retractors. Wanninger et al. 2000 ). Detailed investigations of The high complexity of the bivalve larval muscular muscular activity during torsion in larvae of the body plan combined with the next-to-nonexistent limpet Patella strongly suggest that both larval detailed morphological and developmental analy- retractors, aided by hydraulic movements of the ses calls for future studies to fully assess the plastic- foot, are the main driving forces of this process in ity of myogenesis in this molluscan class. patellogastropods (Wanninger et al. 1999b , 2000 ). Scaphopod larvae have individual muscle This was, however, not confi rmed for two vetigas- fi bers that emerge from the anlagen of the adult tropods (Page 2002 ), illustrating that the dynamics foot and head retractors and serve as prototroch of ontogenetic torsion may be rather plastic. retractors. These also disappear at metamorpho- Formation of the adult dorsoventral (i.e., shell sis, but since they lack distinct shell insertion or foot) retractor muscles already starts in the sites, they are probably not homologous to any of veliger stage prior to the loss of the larval retrac- the gastropod and bivalve larval retractor systems tors in both gastropods and bivalves. In gastro- (Wanninger and Haszprunar 2002b ). pods, one pair of adult shell muscle precursors is Muscles form the major fraction of cephalo- formed, even in patellogastropods, which have a pod soft tissue, and although its adult functional multi-bundled horseshoe-shaped shell muscle morphology and physiology has been subject to system as adults. This indicates that one pair of a great body of research (e.g., Kier 1988 , 1991 , adult shell muscles represents the basal gastro- 1996 ; Kier and Thompson 2003 ; Kier and Stella pod condition. Since late gastropod larvae 2007; Kier and Schachat 2008 ), no recent undergo a stage where both the larval and the pre- account on cephalopod myogenesis employing formed adult retractor systems exist alongside fl uorescence labeling, confocal microscopy, and each other, and because the larval retractors are 3D reconstruction techniques exists. The direct usually striated while the adult shell muscles are mode of cephalopod development suggests that smooth, both systems most likely evolved inde- myogenesis proceeds rapidly without “larval” pendently and are thus not ontogenetically components, but the morphogenetic and cellular homologous (Wanninger et al. 1999a ). dynamics of this process are unknown. 7 Mollusca 131

Skeletogenesis

Mollusks have evolved various ectodermal hard parts including spicules, shell plates, and exter- nal as well as internal shells. In aculiferans, the spicules are secreted by individual, invaginated cells. These are distributed across the entire dor- solateral mantle region in aplacophorans. In the eight shell plates-bearing polyplacophorans, spicules, if present, are restricted to the perino- tum (girdle) (Haas 1981 ; Okusu 2002 ; Nielsen et al. 2007; Todt and Wanninger 2010). In many indirect developing conchiferans, the fi rst- formed, unsculptured (smooth) embryonic shell (protoconch I; sometimes termed prodissoconch I in bivalves) is often followed by the adult shell (teleoconch), which may exhibit a variety of growth and/or color patterns and grows until the death of the animal (Fig. 7.11 ). The protoconch I often breaks off in later stages and is therefore usually only identifi able in larvae and early juveniles. Fossil remains of belemnite internal shells confi rm that such an embryonic shell also belongs to the evolutionary history of cephalo- Fig. 7.11 Ontogenetic continuity of molluscan shells. pods (Müller 1994 ). Many indirect developing mollusks, such as the scapho- In addition to these two common shell types, pod depicted here, form a smooth (unsculptured) embry- onic shell, the protoconch I, from a distinct shell fi eld of many planktotrophic caenogastropods and the mantle. In some gastropods and bivalves (but not in bivalves have a third, intermediate, often richly scaphopods), a second shell is formed during larval life, the ornamented shell, the so-called larval shell or often richly ornamented larval shell (protoconch II). After protoconch II (sometimes termed prodissoconch settlement and metamorphosis, the juvenile/adult shell (teleoconch) starts to form from the mantle edge, usually II in bivalves) (Bandel 1975 ). Both the proto- showing characteristic growth patterns. In scaphopods, conch II and the teleoconch, together with the the protoconch I forms from a single dorsal shell fi eld that often multilayered periostracum, are formed suc- extends laterally on both sides and subsequently encloses cessively from cells located in folds along the the entire visceral region, leaving a characteristic ventral fusion line, the suture. Size of this early juvenile Antalis mantle margin (Kniprath 1977 ; Checa 2000 ) and entalis specimen is 750 μm not from a single shell fi eld as the protoconch I. The various layers of molluscan adult shells genesis have been identifi ed. These suggest that may include nacre (mother of pearl) in some gas- the molecular pathways that underlie molluscan tropods (e.g., the tropical abalone Haliotis ) and shell formation evolve considerably fast (Jackson bivalves (e.g., the pearl oyster Pinctada max- et al. 2007 , 2009 ). In a number of gastropods ima), which most likely evolved independently (e.g., marine nudibranchs and other hetero- multiple times (Jackson et al. 2009 ). With a high branchs), the adult shell was secondarily lost, but potential interest for industrial applications (e.g., the protoconch I is still prevalent in the larvae of pearl production), the genetics of molluscan many indirect developing species. Octopods and biomineralization and nacre formation are sub- pelagic decapod squids either have no or noncal- ject of ongoing research, and transcripts of cifi ed remnants of adult shells, as, e.g., exempli- numerous genes involved in molluscan skeleto- fi ed in the chitinous gladius of Loligo . 132 A. Wanninger and T. Wollesen

Red lines ) with data 1981 , which lacks a conchiferan Argonauta eld formation process has been described. eld anlage is instead overgrown by surrounding ectodermal cells. eld anlage is instead overgrown indicate sites of shell secretion (Figure redrawn from Kniprath ( indicate sites of shell secretion (Figure redrawn invagination process may be (partly) prevented by the yolk-rich eggs. Here, the by the yolk-rich eggs. process may be (partly) prevented invagination shell fi in the pelagic octopod even Interestingly, shell, a rudimentary shell fi compiled from various sources cited therein) elds ). The far left ; yellow . In cephalopods, the Limax Comparative conchiferan shell formation. Conchiferan fi Comparative derive from a posttrochal monolayered ectodermal cell layer ( derive Fig. Fig. 7.12 shell- secreting ectodermal regions form by initial invagination, followed by evagi- followed form by initial invagination, secreting ectodermal regions shell- and internaliza- off by budding may be prevented nation in most taxa. Evagination tion of the shell secreting tissue, which subsequently forms a sac in groups with an internal shell, e.g., terrestrial slugs such as 7 Mollusca 133

Ontogeny of the various conchiferan shell subsequent evagination can still be observed, but types follows a considerably complex but rela- shell secretion does not occur (Fig. 7.12 ; Ussow tively conserved pattern (Fig. 7.12). The process 1874 ; Appellöf 1898 ; cf. Kniprath 1981 ). The is highly dynamic and includes invagination and characteristic, fragile brood chamber carried by subsequent evagination of defi ned ectodermal the reproductive female is a secretory product of domains. For a detailed comparative summary the arms and is not related to the conchiferan of data on a number of mollusks, see Fig. 7.12 shells. herein as well as Kniprath ( 1981 ). Accordingly, Early researchers have proposed that shell a so-called embryonic shell field is formed secretion already starts at the stage of invagi- from a posttrochal, thickened, monolayered nation and therefore termed this morphological ectodermal epithelium some time after gastrula- feature “shell gland” (Ganin 1873 ; Lankester tion (Lankester 1873 ; Fol 1875 ; Bütschli 1877 ; 1873 ). However, the onset of shell secretion Kowalevsky 1883b ; Patten 1886 ; Lillie 1895 ; seems to be rather plastic and for numerous mol- Schmidt 1895 ; Meisenheimer 1901 ; Herbers lusks secretion has been claimed to occur only 1913). This embryonic shell fi eld invaginates, after evagination (see Kniprath 1981 for sum- and the organic outer layer of the shell, the peri- mary and discussion). In order to be less defi nite ostracum, starts to form at the margins of the about the exact site and timing of primary shell resulting pore, thus acting as a scaffold for shell secretion, the more neutral term “shell fi eld” secretion (Kniprath 1977 , 1981 ). If an external was subsequently used for the entire epithelial shell is formed (i.e., in scaphopods, bivalves, domain concerned with shell secretion, irre- and many gastropods), the shell fi eld subse- spective of whether in a pre-invaginated, invagi- quently re-fl attens (evaginates), again forming a nated, or evaginated state (Blochmann 1883 ; defi ned region in the posterior portion of the Schmidt 1895 ; Kniprath 1981 ). This is still the mantle from which secretion of the (embryonic) preferred terminology today, although the terms shell starts (Fig. 7.12 ; Kniprath 1981 ). This “shell fi eld” and “shell gland” are often used ectodermal region is the one that is commonly interchangeably. referred to as the molluscan “shell fi eld” (as Comparative cell lineage data show that the opposed to the embryonic shell fi eld, see above). micromeres 2d, 3c, and 3d contribute to the poly- During polyplacophoran shell plate formation, placophoran shell fi elds, while spicules are the invagination-evagination process seems to formed from descendants of the 1a, 2a, 2c, 3c, be lacking. and 3d cell. In the scaphopod Antalis, only 2d In the cephalopods, shell fi eld invagination was identifi ed as shell fi eld precursor, and this appears to be at least partly prevented by the mas- was also proposed for the bivalve Anodonta sive, yolky egg. Contrary to nautiluses, which (Herbers 1913 ) and various gastropods (Conklin retain their external shell, the surrounding epithe- 1897 ; Robert 1902 ; D’Asaro 1966 ), with an addi- lium overgrows the shell fi eld anlage in coleoids. tional contribution of 2c in Ilyanassa (Cather This epithelium is subsequently budded off and 1967 ). In the basal gastropod Patella , all second internalized (Fig. 7.12 ; Lankester 1873 , 1875 ; quartet micromeres appear to be involved in the Appellöf 1898 ; Spiess 1972 ). In terrestrial pulmo- establishment of the shell fi eld (Table 7.1 ). nate slugs an internal shell sac is formed after Accordingly, the cell lineage of the conchiferan invagination of the embryonic shell fi eld. The protoconch I appears to be rather conserved, shell fi eld is then closed, followed by secretion of while cell lineage of the polyplacophoran (adult!) the internal shell (Fig. 7.12 ; Gegenbaur 1852 ; shell plates deviates from this pattern. The gene- Schmidt 1895 ; Meisenheimer 1896 , 1898 ; see alogy of the cells at the mantle edge that secrete Kniprath 1981 ). In the shell-less pelagic octopus the conchiferan adult shell (teleoconch) is Argonauta (paper nautilus), an invagination and unknown. 134 A. Wanninger and T. Wollesen

Contrary to the remaining conchiferans, together with the developing mantle, grows ven- bivalves have a two-partite shell whose valves are trolaterally, before eventually fusing on the ven- separated and interconnected by an organic liga- tral side. There, the suture marks the fusion zone ment. However, the early shell fi eld anlagen arise of the embryonic shell. The tubular teleoconch is from a single ectodermal domain that is subdi- subsequently secreted from cells of the mantle vided only after evagination, during which the edge (Fig. 7.11 ; Wanninger and Haszprunar interconnecting hinge and later the ligament form 2001 ). (Fig. 7.12 ; Ziegler 1885 ; Lillie 1895 ). Cell lin- eage studies have shown that the micromere 2d constitutes the shell-founding cell (often denoted GENE EXPRESSION “X”) in bivalves. This is the fi rst cell in bivalve embryology that divides bilaterally, and its pro- Since experimental protocols involving reverse genitors give rise to the two-partite, symmetrical genetics (e.g., RNA interference) are only begin- early shell. 2d (X) gives rise to its daughter ning to emerge for mollusks (e.g., Rabinowitz micromeres X1 and X2 prior to gastrulation. et al. 2008 ; Hashimoto et al. 2012 ), assessments Accordingly, determination of the lineage pattern of gene functions still largely rely on expression of the bivalve (embryonic) shell is specifi ed studies obtained by in situ hybridization already at this early developmental stage (Kin (Figs. 7.13 and 7.14 ). In gastrulating molluscan et al. 2009 ; Kurita et al. 2009 ). embryos, brachyury is expressed adjacent to the In coleoid cephalopods, the shell fi eld develops blastopore and in cells of the putative ectoderm in a roundish groove (Fig. 7.12). After establish- (Lartillot et al. 2002b; Kin et al. 2009 ). In the ment of the internal shell sac, a secondary subdivi- Japanese spiny oyster, Saccostrea kegaki, it was sion into two halves takes place in octopods also found to be expressed in cells around the (Spiess 1972 ). In decapods (squids and cuttle- developing anus (Kin et al. 2009 ). Together with fi sh), portions of the posterior shell sac detach the mitogen-activated protein kinase (MAPK) bilaterally and form the fi n pockets (Bandel and signaling cascade, brachyury is likely to be Boletzky 1979 ). In shelled non-cephalopod con- involved in establishing correct cell orientation chiferans, the embryonic shell is formed from a during early ontogeny, similar to its role in other single, centrally located secretion center of the bilaterians. The transcription factor snail and the shell fi eld (one center per valve in bivalves). morphogen hedgehog were found to be involved In the polyplacophoran larva the fi rst seven shell in (neuro)ectodermal patterning in the basal gas- plates are formed synchronously and appear to be tropod Patella (Nederbragt et al. 2002c ). Genes of posttrochal origin, although a pretrochal contri- expressed during molluscan mesoderm forma- bution to the fi rst plate is sometimes proposed tion include twist , forkhead, goosecoid, caudal , (Heath 1898 ; Wanninger and Haszprunar 2002a ; and vasa (Table 7.2 ; Lartillot et al. 2002a ; Le Henry et al. 2004 ). The eighth shell plate develops Gouar et al. 2003; Kakoi et al. 2008) as well as considerably after metamorphosis. Since seven- the homeobox gene Mox , which was found in the shelled fossils assigned to either the Polyplacophora paired mesodermal band and cells destined to or Aplacophora are also known from the fossil become adult (but not larval) muscles (Hinman record, this feature is nowadays often attributed to and Degnan 2002 ). In the highly muscular cuttle- the last common aculiferan ancestor – a hypothesis fi sh Sepia offi cinalis , several muscle-specifi c corroborated by developmental data on neomenio- transcription factors, among these Nk4 , MyoD , morph myogenesis (see above). and Myf5 , are expressed during early phases of The protoconch I of scaphopods arises from a myogenesis (Grimaldi et al. 2004 ; Navet et al. single anlage as in the other conchiferans and, 2008 ). 7 Mollusca 135

Velum Mouth Cerebral ganglia mark the Pedal ganglia Mantle fold Hox1, Hox4, Post1 Hox4, Hox1, Foot Apical organ (hatched area) Gibbula varia. Asterisks Gibbula Operculum Coexpression of and Post2 Shell Pleural ganglia Coexpression of & Cdx Gsx, Xlox, Hox5, Esophageal ganglia Post-torsional veliger larva Post-torsional Visceral sac Visceral ganglion A D V Hox genes Hox Hox1 Hox2 Hox3 Hox4 Hox5 (Hox6) Lox5 Antp Lox4 Lox2 Post1 Post2 genes ParaHox Gsx Xlox Cdx P ) ) b , Coexpression of & Lox4 Antp, Lox5, in the velum Apical organ 2010a , Mouth 2009 Operculum with Foot Post2 (hatched area) Post1 Post1 and Cerebral ganglia Pedal ganglia Coexpression of Hox1 Mantle fold Pleural ganglia Mantle Esophageal ganglia Visceral sac Sketch drawings summarizing Hox and ParaHox gene expression in the pre- and posttorsional larva of the vetigastropod of the vetigastropod in the pre- and posttorsional larva gene expression summarizing Hox and ParaHox drawings Sketch Visceral ganglion Shell AB veliger larva Pre-torsional Fig. Fig. 7.13 statocysts (Based on data from Samadi and Steiner ( statocysts 136 A. Wanninger and T. Wollesen

Fig. 7.14 Gene expression in developing scaphopod, gas- Scale bar: 25 μm (Images B and C taken and modifi ed from tropod, and cephalopod mollusks. ( A ) Gsx expression Samadi and Steiner (2010b )). ( D – F) Gene expression in (arrowheads ) in the apical region of an early scaphopod prehatching embryos of the cephalopod Idiosepius notoi- trochophore (Antalis entalis ; Wollesen, unpublished data; des . Ventral faces up. Scale bars: 150 μm. (D ) The ParaHox anterior faces up). Scale bar: 100 μm. (B ) The ParaHox gene Gsx is expressed in the supraesophageal mass (sem ) gene Gsx is expressed in the apical organ (gray arrow- and optic lobes (ol ). (E ) Expression of Pax258 in the supra- head ), in the dorsal median episphere (red arrowheads ), esophageal mass of the central nervous system and in the and around the stomodeum (s) (yellow arrowheads ) in the arms. (F ) POU3 is expressed in the supraesophageal and trochophore larva of the vetigastropod Gibbula varia . posterior subesophageal mass (pem ). Further abbrevia- Anterior view. Scale bar: 20 μm. ( C ) Gsx expression in the tions: apical organ (ao ), arm (a ), digestive gland ( dg ), eye anlagen of the cerebral ganglia (red arrowheads ) and the (e ), foot ( f), mantle ( m), operculum (o ), prototroch (pt ), forming radula sac (r ) ( yellow arrowheads ) of the veliger velum ( v), visceral mass (vm ) larva of the vetigastropod Gibbula varia. Anterior faces up.

Only few early endodermal markers have been Eleven Hox and three ParaHox genes have characterized in mollusks. These include hedge- been identifi ed in mollusks. However, compara- hog, a foregut patterning gene in Sepia , and tive analyses of expression patterns of these and forkhead, which, after expression in the 3A, 3B, other key developmental regulators are only and 3C macromeres, is found in endodermal cells beginning to emerge. Thereby, the works by in Patella (Lartillot et al. 2002a ). Samadi and Steiner (2009 , 2010a , b ) on the caeno- 7 Mollusca 137 ) ) ) ) 2012 ) ) ) (continued) 2015 2000 2008 2009 2009 2009 Fritsch et al. ( Kin et al. ( raHox, and other homeobox genes eld Kin et al. ( eld Hashimoto et al. ( eld Jacobs et al. ( eld Kin et al. ( Collinear expression in ectoderm, endoderm, and mesoderm along the anterior- Collinear expression hyposphere posterior axis of the larval Expression domain during development Expression domain during development Reference Primary trochoblasts, ciliary band, ciliary cells in stomach of D-shaped larva Primary trochoblasts, ciliary band, cells in stomach of D-shaped larva et al. ( Kakoi Shell fi 3-D blastomere during early cleavage, ventral region from blastopore through region ventral 3-D blastomere during early cleavage, anus part, around prospective vegetal-most Pair of 2d descendant cells, cell posterior to blastopore, endodermal cells Pair Larval Larval mesoderm Primary trochoblasts, ciliary bands, telotroch Two-cell stage, four 2d descendant cells Two-cell Shell fi Shell fi Shell fi

Post2 ), Antp (=

Hox7

,

5

3 – tubulin

/

-

Hox1 Hox genes Gene Non-homeobox genes Tektin Dpp Brachyury Beta Frizzled Vasa Arp2 Hox genes Hox1 Homeobox genes Engrailed Homeobox genes Engrailed

, ,

Expression patterns of key developmental genes in Mollusca. Note the overall paucity of available expression data of Hox, Pa expression paucity of available genes in Mollusca. Note the overall developmental Expression patterns of key Species Class/ Acanthochitona crinita Acanthochitona Polyplacophora Transennella tantilla Transennella Saccostrea kegaki Saccostrea Saccostrea kegaki Saccostrea Nipponacmea fuscoviridis Gastropoda Lepidochitona caverna Lepidochitona Bivalvia Bivalvia Transennella tantilla Transennella Saccostrea kegaki Saccostrea throughout the phylum throughout the phylum Table Table 7.2 138 A. Wanninger and T. Wollesen ) 2003 ) 2010a , 2009 ( eld Hinman et al. ( eld eld eld eld eld Samadi and Steiner Trochophore: ventral posttrochal neuroectodermal cell close to “anal marker” posttrochal neuroectodermal cell close to “anal marker” ventral Trochophore: ectodermal cell in posterior foot rudiment, pedal and esophageal ganglia Veliger: Trochophore: semicircle of ectodermal cells around foot anlage Trochophore: pedal and pleural ganglia Veliger: Trochophore: shell fi Trochophore: mantle, shell fi Veliger: Trochophore: ectodermal cells on both sides of foot rudiment (anlagen Trochophore: kidneys?) mantle, cells close to operculum, anlagen of pleural and esophageal Veliger: ganglia Trochophore: ciliated and non-ciliated prototrochal cells Trochophore: visceral ganglion velum, Veliger: ciliary cells of prototroch, apical organ Trochophore: cerebral ganglion, apical ganglion velum, Veliger: of ectodermal cells in apical region 2 rows Trochophore: cerebral ganglia and commissure region, prevelar Veliger: Trochophore: shell fi Trochophore: mantle, shell fi Veliger: Trochophore: ventral posttrochal area on both sides of foot anlage ventral Trochophore: gland visceral mass and digestive mantle covering Veliger: 4 ectodermal cells close to apical organ Trochophore: cerebral ganglia area, apical organ, cells anterior and posterior of velar Veliger: and commissure Expression domain during development Expression domain during development Reference Trochophore and veliger: shell fi and veliger: Trochophore Trochophore: ectodermal cells close to mouth, “anal marker”, and anterior foot ectodermal cells close to mouth, “anal marker”, Trochophore: pedal and esophageal ganglia, ectodermal mesodermal (?) expression Veliger: in head and foot Trochophore: semicircle of ectodermal cells close to foot Trochophore: cord nerve pleural and pedal ganglia, palliovisceral Veliger: Trochophore: ectodermal cells close to foot anlage Trochophore: anlagen of pleural and esophageal ganglia pedal region Veliger: Trochophore: ectodermal cells in antero-ventral mantle, in ectodermal and ectodermal cells in antero-ventral Trochophore: neck and foot of larval mesodermal (?) regions branchial, and esophageal ganglia, in vicinity of pleuropedal ganglia Veliger: Shell fi

Hox2 Hox3 Post2 Hox4 Hox5 Lox4 Lox2 Post1 Lox5 Antp Gene Hox1 Hox2 Hox3 Hox4 Hox5 Hox1

(continued) Species Class/ Haliotis asinina Gibbula varia Gibbula Table 7.2 Table 7 Mollusca 139 ) ) ) ) ) ) ) ) 2009 2002b 2002a 2002a ) ) 2012 2012 ) 2003 ) 2002a (continued) 2000 2008 2007 2003 , b ) , ) ) 2010b 2002a 2003 2002 O’Brien and Degnan O’Brien and Degnan ( Le Gouar et al. ( Nederbragt et al. ( ( ( ( eld Nederbragt et al. ( base of foot eld and in posterior region, Iijima et al. ( eld Hashimoto et al. ( eld, apical region Hashimoto et al. ( eld, precursor cells of eyes, pretrochal ectoderm (apical organ?), and around pretrochal ectoderm (apical organ?), eld, precursor cells of eyes, Shell fi Nervous system, around stomodeum, precursors of primary and accessory Nervous trochoblasts Anterior mesoderm, anterior expression in all three germ layers Anterior mesoderm, anterior expression Lartillot et al. ( Single posterior cell in mantle, ectodermal cells in developing foot, anlagen of Single posterior cell in mantle, ectodermal cells developing and cephalic tentacles, and esophageal ganglia?, anlagen of statocysts eyes ctenidial and osphradial rudiments Early cleavage, dorsolateral pedal region, statocysts, ectoderm, pallial chamber, ectoderm, pallial chamber, statocysts, dorsolateral pedal region, Early cleavage, anlagen of epipodial tentacles Right/left ectoderm of trochophore ( Grande and Patel Expression domain during development Expression domain during development Reference Buccal, branchial, and esophageal ganglia Giusti et al. ( Gastrulation: ectodermal cells of blastopore, paired mesentoblasts giving rise to Gastrulation: ectodermal cells of blastopore, paired mesentoblasts giving posterior mesoderm of trochophore posterior neuroectoderm and parts of mesoderm Trochophore: Shell fi Anlage of apical organ, cells in posterior ectoderm, shell fi Anlage of apical organ, Apical organ, around mouth and foregut around mouth and foregut Apical organ, Dunn et al. ( Shell fi blastopore Cells of shell fi Apical organ, cerebral ganglia, mouth, radula, odontoblasts Apical organ, Samadi and Steiner Nervous system (e.g., apical organ) system (e.g., apical organ) Nervous Nederbragt et al. ( Gut, early neuroectoderm Paraxial mesodermal bands in trochophore, muscle system of veliger mesodermal bands in trochophore, muscle system of veliger Paraxial Hinman and Degnan Nervous system, foot mucus cells, radula sac, statocyst system, foot mucus cells, radula sac, statocyst Nervous O’Brien and Degnan , head - Grainy ),

4 /

CS1

,

Cdx bmp2

,

(

ferritin Non-homeobox genes Dpp Otx Gsc POU4 Pax258 Pitx Gene Hox5 ParaHox ParaHox genes Cdx Engrailed Engrailed Nk2.1 Dpp Engrailed Gsx Xlox Mox Other homeobox genes Otp POU, Pax, and Mox genes POU, Pax, POU3

,

(continued) Species Nipponacmea fuscoviridis Lottia gigantea Biomphalaria glabrata Class/ Haliotis rufescens Patella vulgata Patella Nipponacmea fuscoviridis Patella vulgata Patella Haliotis rufescens Lymnaea stagnalis Lymnaea vulgata Patella Gibbula varia Gibbula Patella vulgata Patella Haliotis asinina Table 7.2 Table 140 A. Wanninger and T. Wollesen ) ) ) ) ) ) ) ) 2009 2002c ) ) ) ) ) ) ), ) 2004 2002 2010 2002a 2002b 2010 2010 2008 2007 2007 2007 2007 2007 2007 ) 2009 ( Kranz et al. ( Lambert et al. ( Jackson et al. ( eld of trochophore et al. ( Koop eld, micro- and macromeres during early cleavage eld, micro- and macromeres during early cleavage et al. ( Koop eld Iijima et al. ( Various domains in competent veliger domains in competent veliger Various and Degnan Williams Nervous system Nervous Le Gouar et al. ( Ectoderm Lespinet et al. ( Expression domain during development Expression domain during development Reference Shell fi Nervous system (ventral midline of trochophore) system (ventral Nervous Nederbragt et al. ( 4 pretrochal cells in trochophore Ventrolateral ectoderm in trochophore Ventrolateral Shell fi Micromeres during early cleavage (maternally expressed), putative mesodermal putative (maternally expressed), Micromeres during early cleavage bands of trochophore Right/left ectoderm of trochophore ( Grande and Patel Around anus Dunn et al. ( Blastopore, ventral midline, ventral edge of shell fi midline, ventral Blastopore, ventral 3d cell during early cleavage, ectoderm, blastopore 3d cell during early cleavage, Lartillot et al. ( Anterior mesoderm, endoderm Lartillot et al. ( Apical organ, posterior and ventral ectodermal cells, paraxial mesodermal bands posterior and ventral Apical organ, Apical ectodermal cells, posttrochal lateral and posterior cells Apical organ Apical organ Dunn et al. ( Ectoderm surrounding mouth and ventrolateral ectoderm, pretrochal region ectoderm, pretrochal region Ectoderm surrounding mouth and ventrolateral et al. ( Koop Ectodermal and subectodermal cells in head and visceral mass, pedal region, Ectodermal and subectodermal cells in head visceral mass, pedal region, close to mouth mantle cavity, 3d cell during early cleavage 3d cell during early cleavage et al. ( Koop ,

Tpmt , MLF1 DAu172c , ,

3 -

13

/

encoding gene - pim

,

Wnt2 DAu322c nanos

, kak , ,

,

catenin - Ther Onecut Soxb Snail Gene Dpp Hh HSP90A Dpp Def Serotonin MAP kinase Vasa nodal FoxA Brachyury Brachyury Fox Coe Elav Tectin3 β

,

(continued) Species Ilyanassa obsoleta Patella vulgata Patella Class/ Haliotis asinina Lymnaea stagnalis Lymnaea Patella vulgata Patella Haliotis asinina Lottia gigantea Biomphalaria glabrata Haliotis rufescens Haliotis asinina Haliotis rufescens Table 7.2 Table 7 Mollusca 141 ) ), ) ), ) ) ) ) ) ) ) ) 2013 2003 2014 2008 2004 ) 2012 2013 2009 2013 2009 2008 2009 2003 herein; 7.12 Grimaldi et al. ( Navet et al. ( Navet Navet et al. ( Navet Wollesen et al. ( Wollesen bers bers : myocytes giving rise to circomyarian helical and cross-striated fi giving : myocytes : myoblasts giving rise to helical smooth-like fi rise to helical smooth-like : myoblasts giving myf5 myoD Nervous Nervous system Fig. Expression domain during development Expression domain during development Reference Palliovisceral and stellate ganglia, arms Palliovisceral Lee et al. ( Nervous system Nervous Buresi et al. ( Palliovisceral and stellate ganglia, arms, funnel Palliovisceral Pedal ganglia, arms Pedal and stellate ganglia, funnel, arms Arms, buccal lappets, light organ lappets, light organ Arms, buccal Pedal ganglia, arms Palliovisceral ganglia, arms, funnel Palliovisceral Arms, funnel Nervous system, eyes system, eyes Nervous Buresi et al. ( Nervous system, sensory organs, eyes eyes system, sensory organs, Nervous et al. ( Farfán Endoderm (esophagus), nervous system, muscle system Endoderm (esophagus), nervous Grimaldi et al. ( Developing iridosomes throughout the body Developing Andouche et al. ( Muscle system et al. ( Navet Eyes, nervous system, sensory organs system, sensory organs Eyes, nervous Hartmann et al. ( Nervous system, lentigenic cells of eyes system, lentigenic cells of eyes Nervous Ogura et al. (

POU6 , like - POU4

myoD , ) 9 and / specifi c transcription c transcription specifi - POU3 ) ,

like

-

6

lhx2

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(

Muscle factors ( Hox genes Gene Hox1 Non-homeobox genes Elav1 Hox3 Hox5 Lox4 Post1 Post2 POU and Pax genes POU and Pax Lox5 Antp POU2 Otx Other homeobox genes Apt ectin genes Refl Hh Nk4 Pax6 Six3 Myf5

,

(continued) Species Class/ Euprymna scolopes Cephalopoda Sepia offi cinalis Sepia offi Idiosepius notoides Sepia offi cinalis Sepia offi Euprymna scolopes Sepia offi cinalis Sepia offi Idiosepius paradoxus Table 7.2 Table 142 A. Wanninger and T. Wollesen gastropod Gibbula varia , by Lee and coworkers while they were recruited secondarily into the for- (2003 ) on the decapod squid Euprymna scolopes , mation of specifi c morphological features in the and by Fritsch et al. ( 2015) on the polyplacopho- conchiferans only (Table 7.2 ). ran Acanthochitona crinita provide the most com- As noted above, some Hox and ParaHox genes prehensive descriptions of Hox (and, in Gibbula are expressed in specifi c larval features in scaph- only, ParaHox) gene expression profi les for mol- opods and gastropods, notably in Gibbula varia . lusks available to date (Table 7.2, Figs. 7.13 and This includes Lox4 and Antp in the prototroch and 7.14B, C ). This is about to change soon, however, Lox2, Lox4, Lox5 , and Gsx in the apical organ with such data being currently generated in the (Table 7.2 , Figs. 7.13 and 7.14A–C ; Samadi and lab of the authors on various molluscan clades, Steiner 2010a). Several other genes have been including scaphopods and bivalves (Fig. 7.14A ; identifi ed to act in gastropod apical organ forma- Wollesen et al. unpublished). Additional data on tion, including collier, nk2.1, tectin3, engrailed , the expression of selected genes of the Hox, POU, and otp (Table 7.2 ; Nederbragt et al. 2002a , b ; and Pax families are available for another Dunn et al. 2007 ; Jackson et al. 2010 ). Although gastropod, Haliotis asinina (O’Brien and Degnan the direct developing cephalopods lack an apical 2002a, b , 2003 ; Hinman et al. 2003 ). The studies organ, preliminary data suggest that they express on the two gastropods showed that Hox and at least some of these genes in the developing ParaHox genes play an important role in the for- (adult!) central nervous system (Fig. 7.14D–F ; mation of the larval apical organ, prototroch, foot, Wollesen, unpublished), raising the question con- shell, statocyst, radula, and anlagen of the adult cerning conserved versus independent recruitment central nervous system (Table 7.2 , Figs. 7.13 and of these genes in larval and/or adult neurogenesis 7.14B, C ). In the cephalopod Euprymna , all but in mollusks. In both gastropods (Haliotis and one Hox gene investigated were found to be Gibbula) and the cephalopod Euprymna, the ante- involved in arm formation. Hox3, Hox5, Lox4 , rior Hox genes Hox1–5 are not expressed in the and Lox5 are additionally expressed in the cerebral ganglia (Table 7.2 ). This resembles the developing funnel (Table 7.2 ; Lee et al. 2003). situation in classical bilaterian models such as This demonstrates that these genes have acquired Drosophila (Vol. 5, Chapter 1 ) or mouse, which novel functions in conchiferans, since Hox genes do not express Hox genes in the proto- and deu- mainly act in neurogenesis and patterning of the tocerebrum or the fore- and midbrain, respec- anterior-posterior axis in the majority of bilateri- tively (Reichert 2005 ). Similar to these model ans (see Jarvis et al. 2012 for review; note, how- species, the anterior-most region of the mollus- ever, that some Hox genes such as Hox1 , Post1 , can central nervous system, i.e., the anlagen of and Post2 in Gibbula as well as Lox5 and Post1 in the cerebral ganglia, expresses genes such as Otx Euprymna have lost their role in central nervous and Pax258 in cephalopods (Fig. 7.14E ; Reichert system formation; see Table 7.2 and Fig. 7.13 ), 2005 ; Buresi et al. 2012 ). Neuroanatomical stud- and ParaHox genes are involved in digestive tract ies on the basal cephalopod Nautilus suggest that formation (Brooke et al. 1998 ; Holland 2001 ). the optic ganglia of coleoids constitute lateral Novel fi ndings on gene expression in polypla- extensions of the cerebral ganglia (Young 1971 ; cophorans corroborate these fi ndings, since there Shigeno et al. 2008 ), a fi nding that is also cor- the Hox genes likewise show a strict anterior- roborated by co-expression of Otx and Pax6 in posterior collinear expression pattern and are not both ganglia (Hartmann et al. 2003; Navet et al. confi ned to any specifi c morphological features 2009 ; Buresi et al. 2012). In the trochophore larva (Fritsch et al. 2015 ). Accordingly, it appears that of the basal gastropod Patella , however, Otx is polyplacophorans (and maybe the aculiferans in neither expressed in the apical organ nor in the general) have retained the original function of developing cerebral ganglia (Nederbragt et al. Hox genes in anterior-posterior body patterning, 2002b). Instead, ring-like arranged cells express- 7 Mollusca 143 ing Otx were found in the pretrochal region, but it apical organs, trochi, and shells, which now can is unclear whether or not they contribute to neural be assessed by comparative gene expression structures (Nederbragt et al. 2002b ). studies. With respect to some key players in bila- Pax258 is expressed in the supraesophageal terian development, the Hox and ParaHox genes, it mass of the pygmy squid Idiosepius notoides seems that these have played an important role in (Fig. 7.14E ; Wollesen, unpublished) and in the the evolution of conchiferan-specifi c morpholog- cerebral ganglia of adult Haliotis , but not in its ical novelties and that they may have been an apical organ (O’Brien and Degnan 2000 , 2003 ). important evolutionary driving force of conchif- Several POU genes are expressed in the central eran or maybe even molluscan diversifi cation and nervous system of Drosophila and mouse (Treacy evolutionary success (see Wanninger et al. 2008 ). and Rosenfeld 1992 ), and this was also found in It is therefore of particular interest to reveal the Idiosepius and Haliotis , indicating an ancestral full potential of Hox and ParaHox gene functions role of these genes in bilaterian neurogenesis by future comparative studies on representatives (Fig. 7.14F ). In any case, some POU genes (e.g., of the various molluscan classes. POU3 and POU4 ) appear to have been recruited into the formation of novel molluscan traits including the statocysts, tentacles, ctenidia, and BREAKING SYMMETRY: EVODEVO osphradia (Table 7.2 ). OF GASTROPOD HANDEDNESS The transcription factor engrailed is consis- tently expressed in cells demarcating the early Asymmetries are widely distributed and obvious shell fi eld(s) from the remaining mantle epithe- in many bilaterian clades, notably in deutero- lium in polyplacophorans, gastropods, scapho- stomes including the echinoderms and verte- pods, bivalves, and cephalopods. Several other brates. Within mollusks, the gastropods are a genes are known to be expressed in shell forma- particularly drastic example of a handed body tion domains including decapentaplegic (Dpp ) in plan, which is manifested by a dextrally (right- gastropods and bivalves, as well as Hox1 , Post1 , handed or clockwise) versus sinistrally (left- and Post2 in the gastropod Gibbula (Table 7.2 , handed or anticlockwise) coiled shell and gut, Fig. 7.13 ). Research into the molecular regulatory together with a respective right or left position of processes of molluscan shell formation has the anal and genital pores. Interestingly, in some recently received considerable attention, espe- pulmonates such as the great pond snail Lymnaea cially with respect to nacre formation in gastro- and the ram’s horn snail Biomphalaria , both dex- pods and bivalves, whereby an impressive genetic trally and sinistrally coiled individuals occur in toolkit has been identifi ed. This suggests indepen- the same population in the wild. Early investiga- dent evolution of the nacreous layer in various gas- tions into the phenomenon as to how handedness tropod and bivalve lineages (Jackson et al. 2010 ). is determined in these species have shown that Although still rather patchy, the data on mol- this break in symmetry is maternally inherited luscan gene expression currently available pro- and that the dextral phenotype is dominant over vide some distinct molecular markers for the sinistral one (Boycott and Diver 1923 ; conchiferan (larval) structures that are of evolu- Sturtevant 1923 ; Boycott et al. 1930 ; Freeman tionary relevance for Mollusca and the entire and Lundelius 1992 ). Although mapping of the Lophotrochozoa. While these are only fi rst steps responsible locus, termed sinistral , has been car- into reconstructing the genetic regulatory ried out, its identity remains unknown (Asami network that underlies the development of these et al. 2008 ; Liu et al. 2013 ; Namigai et al. 2014 ). characters, they provide important reference In the quest to reveal the cytokinetic dynamics points for homology hypotheses of various mol- and the genetic foundations of this phenomenon, luscan and lophotrochozoan features including important progress has recently been achieved 144 A. Wanninger and T. Wollesen

(Shibazaki et al. 2004 ; Grande and Patel 2009 ; “dextralized” and “sinistralized” embryos were Kuroda et al. 2009 ; see Patel 2009 , Lambert 2010 produced (Fig. 7.15 ). In the surviving embryos that for brief summary). Accordingly, the nodal sig- had maintained the artifi cially induced cleavage naling pathway, known to act in the correct estab- pattern, the entire body organization including lishment of left-right asymmetries in shell coiling was reversed. The manipulated deuterostomes, is involved in defi ning handed- (female) snails that made it to fertile adults gave ness in gastropods (Grande and Patel 2009 ). In rise to offspring with “correct” body organization, accordance with the deuterostome models, nodal i.e., the genetically inherited handedness and not and the downstream acting transcription factor the one artifi cially imposed on the mother animal. Pitx are asymmetrically expressed in gastropods, Embryos with altered cleavage directions at the depending on their right or left chirality: In fi rst or second round of cleavage reverted to their embryos of the limpet Lottia gigantea with a dex- inherited cleavage direction at third cleavage, thus tral body plan, both genes are expressed on the providing evidence that the fi rst symmetry-breaking right side of the embryo, while their expression is signaling events take place at the eight-cell stage to the left in sinistral embryos of Biomphalaria and not earlier (Kuroda et al. 2009 ). The two genes glabrata (Grande and Patel 2009). By chemically of the nodal pathway involved in molecular left- inhibiting nodal signaling, it was found that in right patterning, nodal and Pitx , were expressed those Biomphalaria embryos that had survived according to the expected chirality as predicted by the treatment, Pitx expression was lost. The juve- the blastomere arrangement in the eight-cell niles developed tubular rather than the left- embryo, thus demonstrating that the blastomere handed coiled shells of the usual, non-treated confi guration at this stage defi nes the localization of specimens. Interestingly, non-coiled shells were embryonic nodal signaling (Fig. 7.15 ; Kuroda et al. only obtained if nodal signaling was blocked 2009). Whether these cytogenetic and molecular before the blastula stage, while trochophore lar- determinants of left-right asymmetry are conserved vae treated with the drug developed normal shells among Mollusca, Spiralia, or even the entire proto- (Grande and Patel 2009 ). Since asymmetrical stomes and deuterostomes, and thus were present in expression of nodal is already a result of broken the last common ancestor of all Bilateria, remains to symmetry in the gastropod embryo and not its be seen once more comparative studies become cause, the key symmetry-breaking events must available. take place prior to the fi rst signaling events, i.e., Our current knowledge on molluscan develop- before the blastula stage. mental biology as summarized here is testimony In two Lymnaea species, dextral versus sinistral of the numerous pathways evolution has taken in coiling coincides with a clockwise (dextral) versus shaping the wide phenotypic diversity of anticlockwise (sinistral) direction at third cleavage Mollusca. Deciphering the developmental under- (Freeman and Lundelius 1992 ; Shibazaki et al. pinnings of these mechanisms provides an impor- 2004 ). In order to experimentally test whether this tant window into the evolutionary ancestry of this corresponds to the fi rst symmetry- breaking event fascinating phylum. We have only begun to responsible for handedness in these gastropods, scratch the surface of reconstructing molluscan Kuroda et al. (2009 ) manipulated the (genetically evolutionary history, but with the recently estab- determined) direction of third cleavage in both lished and ever advancing molecular, morpho- sinistral and dextral Lymnaea embryos. Thereby, logical, and in silico methods at hand, and more the arrangement of the fi rst quartet micromeres was and more genomes and transcriptomes being altered during their formation at third cleavage by generated, the stage is set for present and future imposing a 90° shift, such that they came to lie in generations of biologists to engage in studies on opposite direction with respect to their genetically the evolutionary developmental biology of this predefi ned position. As a consequence, artifi cial fascinating phylum. 7 Mollusca 145

Sinistral (recessive) Dextral (dominant)

88

One-cell

Two-cell metaphase

Dextralize Sinistralize Four-cell

Apparent SD, Apparent Metaphase Sinistralize dextral Sl sinistral Dextralize Back to Telophase sinistral Back to dextral

Eight-cell

nodal and nodal and LR LR Late Pitx at the Pitx at the trochophore left side Right side

Juvenile snail

Adult snail

Fig. 7.15 Alteration of handedness in the gastropod major genes involved in left (L )-right ( R) patterning in Lymnaea stagnalis by micromanipulation. Experimental many bilaterian animals, are also reversibly expressed in reversion of chirality during the fi rst two cleavage rounds these micromanipulated specimens. Interestingly, the does not result in organisms with reversed chirality. This is dominant dextral snails exhibit spiral deformation ( SD , a because embryos whose cleavages have been manipulated helical deformation of the blastomeres at the third cleav- at such early stages revert at the eight-cell stage to their age metaphase-anaphase directly linked to the handed- original handedness ( gray arrows). However, reversing ness-determining genes) and spindle inclination (SI , a chirality during third cleavage results in “inverted juve- spiral orientation of the four spindles, as a consequence of niles” and eventually in fertile adults ( red and green spiral deformation, before cleavage furrow ingression), arrows ). Note oppositely coiled shells (photographs) in while the recessive sinistral snails do not show them. these individuals (the entire digestive tract likewise shows However, dextralized snails can be produced from sinistral a mirror image in these healthy snails with respect to non- ones without spiral deformation (Figure reproduced from manipulated ones; not depicted). Nodal and Pitx , two Kuroda et al. (2009 ) with permission from the publisher) 146 A. Wanninger and T. Wollesen

OPEN QUESTIONS (Copenhagen), Pedro Martinez (Barcelona), Christiane Todt (Bergen), his coauthor of this paper and longtime colleague Tim Wollesen (Vienna), and many others, including the pres- • Neuromuscular development in ent and past students and postdocs in his labs in Copenhagen Chaetodermomorpha: How does it conform to and Vienna. AW is also grateful for the generous support of the “aculiferan pattern” as exemplifi ed by the Faculty of Life Sciences, University of Vienna, during the Polyplacophora and Neomeniomorpha? past four years as well as the Danish Science Foundation (FNU) and the Carlsberg Foundation for previous support • Neuromuscular development in protobranch during his Copenhagen years. He also warmly acknowledges bivalves: How do developmental patterns of funding of our Early Stage Research Training Network test cell larvae compare to direct versus indi- MOLMORPH during the years 2005–2009 by the European rect development of other mollusks? Commission. The Austrian Science Fund (FWF) is thanked for current support of a project on aplacophoran EvoDevo • Development in Monoplacophora: How is the (grant number P24276-B22). TW thanks Sonia Victoria seriality of neuromuscular and excretory Rodríguez Monje (Vienna) and all members of the Wanninger structures achieved? lab for help and discussions as well as the crews of the • Comparative tempo-spatial expression Néomysis (Roscoff) and the RV Hans Brattström (Bergen) for assistance with the collection of animals. TW kindly patterns of Hox, ParaHox, and other key thanks Andreas Wanninger, coauthor of this book chapter, developmental genes in Neomeniomorpha, and Bernie Degnan (Brisbane) for their great support during Chaetodermomorpha, Monoplacophora, the last years. The authors thank Jonathan Henry (Urbana), Polyplacophora, Bivalvia, and Scaphopoda: Reuben Shipway (Nahant), Hiroshi Wada (Tsukuba), Michael Schrödl (Munich), Emanuel Redl, Maik Scherholz, What are the functions of individual genes, Alen Kristof, Marlene Karelly, and Marion Hüffel (all and how do they govern development of spe- Vienna) for providing images used in this chapter. cifi c morphological features? • Cell lineages and gene expression profi les of test cells and velar and trochal structures: How References conserved or plastic are they among mollus- can (and lophotrochozoan) larvae? Altnöder A, Haszprunar G (2008) Larval morphology of the brooding clam Lasaea adansonii (Gmelin, 1791) • Cell lineages, gene expression profi les, and (Bivalvia, Heterodonta, Galeommatoidea). J Morphol submicroscopic 3D architecture of apical 269:762–774 organs: How (dis)similar are they among Anderson PD, Bokor G (2012) Conotoxins: potential Mollusca, other Lophotrochozoa, and more weapons from the sea. J Bioterr Biodef 3:120 Andouche A, Bassaglia Y, Baratte S, Bonnaud L (2013) distant taxa, e.g., Cnidaria? Refl ectin genes and development of iridophore pat- • What are the functional roles of apical organs? terns in Sepia offi cinalis embryos (Mollusca, • Cephalopod neuro- and myogenesis: Are there Cephalopoda). Dev Dyn 242:560–571 “larval” remnants? Appellöf A (1898) Über das Vorkommen innerer Schalen bei den achtarmigen Cephalopoden (Octopoda). • Genetic signatures of cephalic appendages of Bergens Mus Arb 12:1–15 scaphopods, gastropods, and cephalopods: Arnolds WJA, van den Biggelaar JAM, Verdonk NH Are they homologous? (1983) Spatial aspects of cell interactions involved in • Gene expression profi les of embryonic and the determination of dorsoventral polarity in equally cleaving gastropods and regulative abilities of their adult conchiferan shells and aculiferan spic- embryos, as studied by micromere deletions in Lymnaea ules: What are the key players that govern and Patella . Roux’s Arch Dev Biol 192:75–85 molluscan biomineralization processes? How Asami T, Gittenberger E, Falkner G (2008) Whole-body does this compare to other animals that form enantiomorphy and maternal inheritance of chiral reversal in the pond snail Lymnaea stagnalis. J Hered 99:552–557 mineralized skeletons? Ax P (1999) Multicellular animals. Springer, Berlin Bandel K (1975) Embryonalgehäuse karibischer Meso- Acknowledgments AW expresses his sincere thanks to the und Neogastropoden (Mollusca). Abh Math Naturw numerous colleagues that have offered their time to discuss Kgl Akad Wiss Mainz 1:1–133 various issues on molluscan and metazoan morphology, Bandel K, Boletzky SV (1979) A comparative study of the development, and evolution over the past many years, often structure, development and morphological relationships of at most peculiar times and in most inspiring locations, in par- chambered cephalopod shells. The Veliger 21:313–354 ticular Gerhard Haszprunar (Munich), Bernie Degnan Baratte S, Bonnaud L (2009) Evidence of early nervous (Brisbane), Jens Hoeg (Copenhagen), Claus Nielsen differentiation and early catecholaminergic sensory 7 Mollusca 147

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Jörn von Döhren

Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger.

J. von Döhren Rheinische-Friedrich-Wilhlems Univerität Bonn, Institut für Evolutionsbiologie und Ökologie , An der Immenburg 1 , Bonn 53121 , Germany e-mail: [email protected]

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 155 DOI 10.1007/978-3-7091-1871-9_8, © Springer-Verlag Wien 2015 156 J. von Döhren

INTRODUCTION (Fig. 8.1C, D), Hoplonemertea (Fig. 8.1E, F), and Bdellonemertea, the latter compris- Anatomy and Systematics ing only one genus of commensal representa- tives ( Malacobdella) (Coe 1943; Gibson 1972 ). Nemertea is a clade of unsegmented, worm- Paleonemertea and Heteronemertea have been shaped Spiralia comprising about 1.300 classifi ed as Anopla due to their proboscis being described species (Fig. 8.1A–F ; Kajihara et al. uniformly organized and lacking a stylet armature. 2008 ). The vast majority inhabits marine benthic In Bdellonemertea a stylet armature of the probos- habitats, but several species are limnic, terres- cis is also absent, but this has been interpreted as trial, or marine pelagic. Most species have been secondary reduction due to the commensalic life- described as predators although a number of par- style of this group. Hence, Bdellonemertea and asitic, commensalic, and probably even scaven- Hoplonemertea have been classifi ed as Enopla, gers are known (Gibson 1972 ). Prey is captured characterized by a primarily armed proboscis, the by means of an eversible proboscis that may be brain being positioned behind the mouth opening armed with one to numerous calcareous stylets in and a more intimate connection of the probos- some clades. The proboscis apparatus comprises cis insertion and the foregut. In Hoplonemertea, the proboscis and the rhynchocoel. It represents two clades have been identifi ed due to their pro- the apomorphic character that has led to the alter- boscis armature: Monostilifera and Polystilifera. native name Rhynchocoela. The rhynchocoel is In Monostilifera the proboscis is armed with a a dorsally located, fl uid-fi lled secondary body single, comparably large stylet in its middle sec- cavity surrounded by muscle layers housing tion. In this clade the mouth opens to the ventral the proboscis. It opens to the tip of the head face of the rhynchodeum making its distal part a via a tube-shaped rhynchodeum (Fig. 8.2A, B ). bifunctional rhynchostomodeum. Polystilifera are Additional characters that unequivocally qualify characterized by a proboscis that is armed in its Nemertea as monophyletic are the ring-shaped middle portion with a cushion equipped with mul- brain surrounding the proboscis insertion instead tiple, relatively small stylets. The connection of of the esophagus, a pair of laterally located lon- the mouth opening with the rhynchodeum varies gitudinal medullary cords, and an endothelialized between species. Both rhynchodeum and mouth blood-vascular system. Apart from that nemer- open independently of each other but in nearby tean anatomy is marked by characters that are positions in many pelagic forms (Pelagica). In arguably plesiomorphic for Spiralia (Turbeville most benthic polystiliferan species (Reptantia), a 2002 ). These include a largely compact arrange- gradual fusion of both openings by sharing a com- ment of the tissue, a medullary cord type organi- mon atrial chamber at the tip of the head is pres- zation of the nervous system; a body wall muscle ent. Recent molecular analyses and reassessment tube comprising minimally two, an outer circular of morphological data, however, give a different and an inner longitudinal, muscle layers; and one picture putting the traditional classifi cation in to several paired lateral protonephridia that are jeopardy (von Döhren et al. 2010 ; Bartolomaeus not arranged in a segmental fashion. Characters and von Döhren 2010; Andrade et al. 2012 , 2014 ; that place Nemertea closer to Trochozoa are Kvist et al. 2014 ). Of the traditional higher-ranking a regionalized through-gut with mouth, fore- taxa, only Hoplonemertea and Heteronemertea are gut, midgut, and anus and the presence of glial recovered (Fig. 8.3 ). Paleonemertea has been rec- type cells in the nervous system (Turbeville and ognized as nonmonophyletic with Hubrechtiidae Ruppert 1985 ; Turbeville 1991 ). being more closely related to Heteronemertea than Nemertean ingroup systematics is pres- to the remaining paleonemertean clades (Fig. 8.3 ). ently in the consolidation phase with many sub- The presence of a specialized larva, the pilidium, clades still being unstable. Traditionally, four in both Hubrechtella dubia (Hubrechtiidae) and higher- ranking taxa have been distinguished: most heteronemertean species leads to them being Paleonemertea (Fig. 8.1A, B ), Heteronemertea combined in the clade Pilidiophora (Fig. 8.3 ). 8 Nemertea 157

A B

C D

EF

Fig. 8.1 Diversity of nemertean species, living speci- adult. ( F ) Riseriellus occultus (Heteronemertea, mens. ( A ) Procephalothrix oestymnicus (Cephalothricidae, Pilidiophora), adult. Note: in lightly colored or unpig- Paleonemertea), adult. (B ) Carinina ochracea mented species the brain ring ( br) is visible through the (Carininidae, Paleonemertea), adult. ( C ) Amphiporus body wall. ae anterior end, br brain ring, mid midgut lactifl oreus (Monostilifera, Hoplonemertea), juvenile. (D ) region, pb proboscis apparatus in rhynchocoel, pe poste- Emplectonema gracile (Monostilifera, Hoplonemertea), rior end (© Dr. J. von Döhren, All Rights Reserved) adult. ( E ) Lineus ruber (Heteronemertea, Pilidiophora),

The remaining paleonemertean species have weak the body wall muscles. In fact, there is no unequiv- support as monophylum based on molecular data ocally apomorphic morphological character that (Andrade et al. 2012 , 2014; Kvist et al. 2014 ). supports paleonemertean monophyly (Fig. 8.3 ). Anatomically, Paleonemertea represent the most Within the armed clades (Enopla), traditional inhomogeneous taxon within Nemertea with some taxonomy was not confi rmed either. On the one species having an additional inner circular muscle hand, Bdellonemertea proved to be an apparently layer, while others vary with respect to the position secondarily reduced member of Monostilifera of the brain and lateral medullary cords relative to according to molecular phylogenetic analyses, 158 J. von Döhren

A Anopla (Palaeonemertea & Heteronemertea)

rhd br es pb mid lmc rhc ret mdn

B Enopla (= Hoplonemertea)

rsd br es sty pb mid lmc rhc ret

ectodermal mesodermal entodermal

Fig. 8.2 Schematic representation (side view) of the organs, and gland cells have been omitted for clarity. br nemertean body plan (A ) Anoplan organization (Paleo- brain, lmc lateral medullary nerve cords, mdn middorsal and Heteronemertea). (B ) Enoplan organization nerve, mid midgut, es esophagus, pb proboscis, ret (Hoplonemertea). Color coding indicates the germ layer retractor muscle of proboscis, rhc rhynchocoel, rhd that the structure originates from. Note: body wall mus- rhynchodeum, rsd rhynchostomodeum, sty stylet appa- culature, nephridia, blood-vascular system, sensory ratus (© Dr. J. von Döhren, All Rights Reserved) rendering Hoplonemertea and Enopla synony- Polystilifera, they are situated far in front of the mous, while on the other hand with the same data brain in Monostilifera (Gibson 1972). Lateral the monophyly of Polystilifera remains a mat- sensory organs have a morphology that is remi- ter of debate (Thollesson and Norenburg 2003 ; niscent of cerebral sense organs in paleonemer- Sundberg and Strand 2007 ; Andrade et al. 2012 , teans but are located more posteriorly on the 2014 ; Kvist et al. 2014 ). Currently, there is strong sides of the animal in the vicinity of the nephro- support for a clade Neonemertea comprising pores. Lateral organs are typical of tubulanid Hoplonemertea and Pilidiophora (Figs. 8.1C–F paleonemerteans but have also been recorded and 8.3 ; Thollesson and Norenburg 2003 ; Andrade from some heteronemertean species (Gibson et al. 2012 , 2014; Kvist et al. 2014). All members 1972 ). Frontal organs that are connected to the of this clade share the presence of a median dorsal almost ubiquitously present cephalic glands and blood vessel between alimentary canal and rhyn- represented by a single epidermal pit in most chocoel (Gibson 1972 ). neonemertean species or by three epidermal pits A number of organs, characteristic of nemer- arranged in a triangular pattern in lineid teans, cannot be placed robustly in an evolution- Heteronemertea might be apomorphic for ary scenario due to their disparate distribution Neonemertea, although a row of median epider- within Nemertea. Cerebral organs connected to mal pits located in the head region have also the brain are a feature of many species, although been described in the paleonemertean genus they are quite different in shape and position. Carinoma (Gibson 1972 ). Pigmented eyes in While being located behind the brain in tubula- adults are characteristic of neonemertean species nid Paleonemertea, Pilidiophora, and reptant as well but show very high variation in size, 8 Nemertea 159

Fig. 8.3 Consensus phylogeny of Nemertea (modifi ed vascular system lined with endothelium; (2 ) Neonemertea: after Andrade et al. 2012, 2014). Numbers indicate apomor- middorsal blood vessel; (3 ) Pilidiophora: pilidium larva; (4 ) phic characters for respective clades. ( 1 ) Nemertea: dorsal, Hoplonemertea: proboscis equipped with stylet apparatus. eversible proboscis housed in fl uid-fi lled rhynchocoel, ring- Note: for the paleonemertean taxa, no apomorphic charac- shaped brain located around the proboscis opening, blood- ter exists (© Dr. J. von Döhren, All Rights Reserved) complexity, and number even between arguably nuptial dances reminiscent of those of epitokous closely related species. Moreover, the eyes in polychaetes in Nipponnemertes pulcher (Berg adults have also been described in a few isolated 1972). In most species, females spawn their paleonemertean species (Gibson 1972 ). eggs freely into the water. Egg masses depos- ited in mucus sheaths are reported from species living in marine (e.g., Tetrastemma candidum , Reproductive Biology Antarctonemertes phyllospadicola ), intertidal (e.g., Lineus ruber , Lineus viridis ), freshwater The majority of nemertean species are gono- and terrestrial habitats (e.g., Prostoma jenningsi , choristic. Only comparably few (mostly non- Apatronemertes albimaculosa ), as well as from marine) hermaphrodites have been described species that are living as parasites on or in other (Friedrich 1979 ). Free-living nemerteans have the animals (e.g., Carcinonemertes species on sev- tendency to aggregate during their reproductive eral crustaceans) (Thiel and Junoy 2006 ). In the season, and there have been anecdotic reports of monostiliferous hoplonemertean Amphiporus 160 J. von Döhren incubator , the female remains in the secreted Egg sizes in nemerteans range from 50 μm in mucus sheath with the developing offspring Carinina (Procarinina ) remanei (Nawitzki 1931 ) apparently providing them with nourishment by to 2.5 mm in Dinonemertes investigatoris (Coe means of complete histolysis of its intestinal tract 1926), with most being between 100 and 300 μm (Joubin 1914 ). in diameter (Friedrich 1979 ). With the relatively In general, spermatozoa are released freely into scarce data at hand, there seems to be no strict the water as well. In those species that aggregate correlation of egg size to the various phylogenetic during reproduction and in mucus spawning species entities, but there is a tendency of paleonemer- males typically come into close contact with either tean eggs being on the smaller side of the spec- the eggs or the female to ensure successful fertiliza- trum, while larger egg sizes are encountered in tion (Bartolomaeus 1984 ; Thiel and Junoy 2006 ). hoplonemertean species (e.g., Pantinonemertes In many species viscid mucous secretions around (Geonemertes ) agricola : 350–450 μm (Coe 1904 ); the worms putatively aid in scaling down the space Nipponnemertes pulcher : 280–340 μm (Berg into which eggs and sperm are shed. This behavior 1972 )). While there is usually only a very thin and has occasionally been termed “pseudocopulation” delicate chorion surrounding the eggs of anoplan (e.g., Carcinonemertes epialti, Lineus ruber, Lineus species, the eggs of hoplonemertean species are viridis ) (Gontcharoff 1951 ; Bartolomaeus 1984 ; invested with a chorion (also termed “vitelline Roe 1984 ; Stricker 1986 ). In Lineus viridis , one to envelope”) that is thicker and usually set off from several males enter a mucus sheath that is secreted the egg membrane by a fl uid-fi lled space of dif- by the female. Into this gelatinous mass, additional ferent dimensions (Stricker et al. 2001 ). In many mucus layers and the eggs are shed (Gontcharoff free-spawning species, a glutinous mucus layer 1951; Bartolomaeus 1984 ). During pseudocopula- surrounds the egg chorion to attach the eggs to the tion in some species (e.g., Carcinonemertes epi- substrate. In some species this mucus is reported to alti ), sperm may enter the female gonads through dissolve in the water soon after the eggs have been the gonopores, and internal fertilization occurs. shed. In this case the mucus possibly enhances Internal fertilization with direct sperm transfer (i.e., attraction and movement of the sperm to the egg. true copulation) has been assumed to occur in some Spermatozoa in nemertean species generally pelagic polystiliferans. Several structures have been comprise a sperm head consisting of an apical interpreted as accessory sperm transfer structures acrosomal vesicle, a condensed nucleus, a mito- such as muscular penes in Phallonemertes murrayi , chondrial mass, and diplosomal centrioles. From sucker- like attachment organs in Plotonemertes the distal centriole, a single fl agellum with a regu- adhaerens, and specialized glandular epithelia lar 9 × 2 + 2 axoneme emanates. Only in the pelagic around the male gonopores in Balaenanemertes polystiliferan Nectonemertes mirabilis afl agellate chuni and some other pelagic polystiliferans (Thiel sperm cells together with separate fl agella have and Junoy 2006). In species of the monostiliferous been observed in the testes (Stricker and Folsom hoplonemertean genus Carcinonemertes, the effer- 1997 ). Morphology of the sperm head is very vari- ent ducts of the testes open into a common duct, able with all components varying in both length termed Takakura’s duct, that widens into a seminal and width (Stricker and Folsom 1997 ; von Döhren vesicle which opens into the intestine near the anal and Bartolomaeus 2006 ; von Döhren et al. 2010 ). opening (Gibson 1972). In some Carcinonemertes Acrosomal vesicles and mitochondria may be dis- species, the anal opening is surrounded by a fl at- located from the terminal poles of the sperm cell in tened or concave muscular area that is used to trans- some species. In most paleonemertean and hoplone- fer sperm from the anus to the female gonopores mertean species, the single mitochondrion represents (Roe 1984). Some accounts of viviparous species the product of the fusion of numerous mitochondria from all phylogenetic lineages demonstrate that during spermiogenesis. Elongated headed sperm vivipary is not uncommon in this phylum. In these has been hypothesized to be correlated with either species internal fertilization has to be expected internal fertilization (e.g., Antarctonemertes phyl- (Thiel and Junoy 2006 ). lospadicola, Carcinonemertes epialti, Cephalothrix 8 Nemertea 161 rufi frons ) or the investment of the egg with a tough, 2013 and references therein). The egg, arrested in resistant vitelline membrane in free-spawning the metaphase of the fi rst meiotic division, has then species (e.g., Cerebratulus lacteus ) (Stricker and an eccentrically located nucleus and is ready for fer- Folsom 1997). In some hoplonemertean species, tilization. Fusion of the sperm with the egg induces a much of the elongation of the sperm head is due to sudden cortical calcium fl ash followed by repetitive a conspicuously elongated acrosomal vesicle (von calcium oscillations in the egg due to the action of a Döhren et al. 2010 ). Interestingly, in these species soluble sperm factor delivered from the sperm to the eggs are invested with both a vitelline envelope and egg. The fi rst two calcium waves start at the point a fairly thick glutinous mucus layer that is resist- where the sperm has fused with the egg, while the ing degradation well beyond the gastrulation of following waves are elicited from a pacemaker the embryo inside (e.g., Paranemertes peregrina , region in the vegetal cortex of the egg opposite of the Emplectonema gracile). site of prospective polar body extrusion. Along with In externally fertilizing species, eggs are com- the calcium waves, there is a global disassembly of monly shed arrested in the prophase of the fi rst mei- the ER microdomains. The calcium oscillations con- otic division, i.e., the germinal vesicle is clearly tinue through fi rst polar body formation and cease visible. In oviparous species with internal fertiliza- before the second polar body is extruded (Stricker tion, this is also the case only in Carcinonemertes et al. 2013 and references therein). Polar bodies are epialti where the eggs are reported to be shed in situated at the opposite pole of the cytoplasmic pro- early cleavage stages (Stricker et al. 2001 ). Freely trusion if one is present. This hints at the animal- spawned eggs typically show a somewhat com- vegetal axis of the egg being already established in pressed morphology due to them having been tightly the gonad during oogenesis (Fig. 8.4A ; Henry and packed in the ovaries. In some eggs, especially when Martindale 1997 ). Polar body formation is followed they are artifi cially extracted from the female, there by decondensation of the male pronucleus and sub- is a cytoplasmic protrusion representing the spot sequent karyogamy (Fig. 8.4B ). where the egg was contacting the ovarian lining dur- ing oogenesis (Iwata 1960 ; Stricker et al. 2001 ). In contact with seawater, they round up and usually EARLY DEVELOPMENT undergo meiotic maturation indicated by germinal vesicle breakdown (GVBD) (Stricker et al. 2001 ). In Diversity of Cleavage in Nemertea some species the cytoplasmic protrusion separates from the egg but remains in its vicinity; in other spe- Although exhibiting quite diverse larval types, cies it disappears soon after fertilization. Abolishment embryonic development of nemerteans is com- of prophase I arrest during GVBD in the heterone- parably uniform. In general, cleavage is of the mertean pilidiophoran Cerebratulus lacteus is medi- holoblastic, equal (homoquadrant), spiral type ated by intracellular signaling of nitric oxide (NO), (Fig. 8.4C–F ). The fi rst cleavage division is merid- cyclic guanosine monophosphate (cGMP), and an ional, passing through the anterior- posterior plane atypical protein kinase C (aPKC). Adenosine mono- as indicated by the location of the polar bodies. phosphate-activated protein kinase (AMPK) blocks It results in a pair of equally sized blastomeres. GVBD, but it can be resumed by the action of cyclic They are initially rounded, with little contact to adenosine monophosphate (cAMP) and protein each other. The embryo soon becomes compact kinase A (PKA) signaling. By these alternative path- prior to second cleavage division (Friedrich 1979 ; ways, an inactive form of the maturation promoting Henry and Martindale 1997 , 1998a ; Maslakova factor (pre-MPF) is activated. The active maturation et al. 2004a ; Maslakova 2010a ). In the heterone- promoting factor (MPF) triggers GVBD which is mertean species Lineus ruber, the fi rst cleavage accompanied by a drastic reorganization of the division has been reported to be unequal result- endoplasmic reticulum (ER) of the egg into numer- ing in a pair of slightly or signifi cantly differently ous ER microdomains about 5 μm in diameter, dis- sized blastomeres (Fig. 8.4D ; Nusbaum and tributed evenly within the cytoplasm (Stricker et al. Oxner 1913 ). This phenomenon, however, is not 162 J. von Döhren

Fig. 8.4 Diversity of the cleavage in Nemertea. Bold arrows and cross-furrows. The dorsoventral axis runs through the larger indicate the most common sequences. (A ) Unfertilized egg. D and the smaller B quadrant. (H ) Eight-cell stage after third The animal-vegetal axis is running through the germinal vesicle cleavage division with equally sized animal (1q ) and vegetal (dashed line), and the cytoplasmic stalk on the opposite vegetal (1Q ) blastomeres, showing an initially radial-like arrangement pole of the oocyte. (B ) Zygote. The animal pole is marked by of blastomeres (see also Fig. 8.5B ). (J ) Eight-cell stage after the polar bodies; the vegetally located cytoplasmic protrusion third cleavage division with animal blastomeres (“micromeres,” has been cast off. (C ) Two-cell stage, equal fi rst cleavage divi- 1q) being larger than vegetal blastomeres (“macromeres,” 1Q ) sion. (D ) Two-cell stage, unequal fi rst cleavage division result- most common in nemertean species studied to date. (K ) Eight- ing in a smaller AB and a larger CD blastomere. (E ) Four-cell cell stage after third cleavage division with animal blastomeres stage after second cleavage division with equal sized blasto- (“micromeres,” 1q ) being of the same size as vegetal blasto- meres and cross-furrows (see also Fig. 8.5A ). One dorsoventral meres (“macromeres,” 1Q ). ( L) Eight-cell stage after third axis is present running through the vegetal cross-furrow blasto- cleavage division with animal blastomeres (“micromeres,” 1q ) meres. (F ) Four-cell stage after second cleavage division with being smaller than vegetal blastomeres (“macromeres,” 1Q ). equal sized blastomeres lacking a cross-furrow. Two alternative Citations in brackets indicate confl icting reports that have been dorsoventral axes are present. (G ) Four-cell stage after second given for the respective species. For further explanations, see cleavage division with unequal-sized blastomeres (C and D ) text (© Dr. J. von Döhren, All Rights Reserved) 8 Nemertea 163 seen in all embryos of a given clutch and has sized blastomere pairs. It is situated between one therefore to be attributed to a certain variability of the larger and the opposing smaller blastomere of cleavage in this species. It is unclear to what in the majority of examined cases (Fig. 8.4G ; extend the differing blastomere size infl uences Nusbaum and Oxner 1913; Schmidt 1964). It is further development. The plane of the second not completely clear, however, how the cross- cleavage division is meridional and perpendicu- furrow is related to the further course of devel- lar to the fi rst, dividing the blastomeres into four opment. The closely related heteronemertean daughter cells of equal size (Fig. 8.4E, F). In the Cerebratulus lacteus does not show any sign of case of the unequally cleaving embryos of Lineus cross-furrows at this stage nor do any of the other ruber , the second cleavage division results in a examined heteronemertean species (Fig. 8.4F ; four-cell stage in which one pair of blastomeres, Friedrich 1979 ; Henry and Martindale 1997 , the progeny of the smaller two-cell stage blasto- 1998a ; Maslakova 2010a ). Due to its variability mere, is smaller than the progeny of the larger in occurrence, it can be assumed that in Lineus two-cell stage blastomere (Fig. 8.4G). This is in ruber the cross-furrow merely represents an contrast to what is observed in, e.g., unequally intraspecifi c variation of development that might cleaving annelids or mollusks (see Chapters 7 not have any infl uence on the determination of and 9 ) in which typically one of the four-cell the future embryonic quadrants. stage blastomeres, the precursor of the dorsal (D) The third cleavage division is synchronous and quadrant, is larger than the other three (Henry equatorial. It results in an eight-celled embryo and Martindale 1998b ). This also speaks for the (Fig. 8.4H–L ). In most species the third cleavage phenomenon of unequal cleavage in Nemertea division is described as clearly dexiotrophic (dex- not being homologous to the unequal cleavage of tral), but there are some accounts of animal blas- Annelida or Mollusca. The four-cell stage blas- tomere quartets being positioned initially exactly tomeres are initially rounded with little connect- opposite to their respective vegetal counterparts ing surface to each other but soon move toward (Figs. 8.4H and 8.5B ), resembling a radial cleavage each other just like in the two-cell stage. The pattern as found in, e.g., cnidarians (Vol. 1, Chapter occurrence of cross-furrows at the four-cell stage 6 ), ectoprocts (Chapter 11 ), phoronids (Chapter has so far been reported from only a few species. 10 ), brachiopods (Chapter 12 ), or invertebrate deu- The paleonemertean species Carinoma armandi terostomes (Vol. 6). In Amphiporus lactifl oreus and tremaphoros and Procephalothrix oestrymnicus Lineus viridis (as Lineus obscurus), the aligned show distinct cross-furrows (Figs. 8.4E and 8.5A ; animal and vegetal blastomeres shift position after Maslakova et al. 2004a ). In the former species, segregation to come to lie as if generated by a the cross- furrows are a result of a slightly leo- regular dexiotropic cleavage division (Fig. 8.4K ; tropic (sinistral) second cleavage division with Barrois 1877 ). This peculiar behavior might be one of the vegetal cross-furrow cells representing due to the high yolk content of the large eggs in the precursor of the future dorsal (D) quadrant. both of the mentioned species. In Cephalothrix However, it is not clear whether the specifi cation rufi frons there is certain variability regarding the of the D quadrant in this species is mediated by angle at which the eight-cell blastomeres are posi- the segregation of cytoplasmic components or by tioned, but subsequent cleavage divisions restore inductive cellular interactions (Maslakova et al. the spiral pattern by being regularly alternating 2004a ). In neonemertean species, cross-furrows leotropic (sinistral) and dexiotropic (Smith 1935 ). are reported from Lineus ruber (Heteronemertea) Regarding the relative sizes of the blastomeres at and Emplectonema gracile (Hoplonemertea), the eight-cell stage, there is some noteworthy vari- although in the latter species a more recent study ation among species investigated. In most species does not confi rm the existence of a cross-furrow the animal eight-cell stage blastomeres (micro- (Fig. 8.4E, G; Delsman 1915; Iwata 1960 ). In meres) are slightly or even considerably larger Lineus ruber the cross-furrow has been recorded than cells of the vegetal blastomere quartet (mac- only in four-cell stage embryos with unequal- romeres) (Fig. 8.4J ; Friedrich 1979 ; Henry and 164 J. von Döhren

Martindale 1997 ; Maslakova et al. 2004a ), while in this species is again reached by a transitory stage Tetrastemma vermiculus , Drepanophorus specta- with the macromeres and the vegetal-most micro- bilis , and Lineus ruber , the relative size differences mere quartet dividing fi rst, followed by the divi- accord to the general nomenclature, i.e., the animal sion of the two animal-most micromere quartets micromeres are smaller than the vegetal macro- (Nusbaum and Oxner 1913 ). Interestingly, there meres (Fig. 8.4L ; Lebedinsky 1897 ; Nusbaum seems to be a tendency in this species to arrange and Oxner 1913 ; Schmidt 1964 ). Both animal and the micromere quartet daughter cells in a planar vegetal blastomere quartets have been reported rather than a spiral pattern, a phenomenon that has to be of roughly equal size in Tetrastemma vari- also been described for the equatorial-most blas- color (Hoplonemertea), Cerebratulus lacteus , and tomeres in Amphiporus lactifl oreus (Barrois 1877 ; Lineus viridis (Figs. 8.4K and 8.5B ; Barrois 1877 Nusbaum and Oxner 1913 ). Relative asynchronies for Lineus viridis as Lineus obscurus ; Hoffman regarding the timing of division of blastomere 1877 ; Wilson 1900 ). In Cephalothrix rufi frons quartets have been described in a number of other there is no externally visible size difference of species, e.g., Carinoma tremaphoros , Cerebratulus animal and vegetal blastomeres in the eight-cell lacteus , Cerebratulus marginatus , Emplectonema stage, but sections reveal that the macromeres are gracile , Malacobdella grossa, and Tubulanus slightly larger, their additional volume projecting nothus (Wilson 1900 ; Zeleny 1904 ; Delsman 1915 ; interiorly into the blastocoel (Smith 1935 ). The Hammarsten 1918 ; Dawydoff 1928 ; Maslakova fourth cleavage division leading to the 16-cell et al. 2004a ). However, while in Lineus ruber the stage embryo is generally reported to be synchro- cleavage divisions of the vegetal pole precede those nous. The transition to the 16-cell stage in Lineus of the animal pole, the reverse is true in the other ruber passes through a series of stages in which exemplifi ed species. Nevertheless, cleavage gener- fi rst the two larger of the animal blastomeres divide ally generates four quartets of animal blastomeres in a leotropic manner, followed by the smaller pair. along with their respective progeny as well as one Finally, the macromere quartet (2A-D) divides to quartet of comparably small vegetal blastomeres. accomplish the 16-cell stage. The 32-cell stage in The paleonemertean Tubulanus nothus represents

Fig. 8.5 Embryonic stages of Nemertea. (A ) Four-cell furrow. (B ) Eight-cell stage of Lineus viridis (Pilidiophora), stage of Procephalothrix oestrymnicus (Paleonemertea) from vegetal; scanning electron micrograph. Note the from vegetal; live specimen; differential interference con- almost radial arrangement of equally sized animal and veg- trast (DIC ) light micrograph. Note the prominent cross- etal blastomeres (© Dr. J. von Döhren, All Rights Reserved) 8 Nemertea 165 an exception in that the regular spiral cleavage germinal vesicle situated on the opposite, animal pattern is largely given up after the fi fth cleavage pole of the egg (Fig. 8.4A). During meiotic matu- division, resulting in a chaotic cleavage pattern in ration the polar bodies form near the animal pole. which neither fourth micromere nor macromere After fertilization the fi rst two cleavage divisions quartets are discernible (Dawydoff 1928 ). pass through the animal-vegetal axis which thus becomes the anterior-posterior axis of the larva. Experimental alteration of the fi rst cleavage plane Determination of the Future by compressing fertilized eggs during fi rst cleavage Body Axes division results in its decoupling from the larval anterior-posterior and dorsoventral axis, indicating Cleavage of nemerteans does not only show the that the cleavage planes in normal development are characteristic pattern, but it also complies with the not the cause of the future larval axes but follow a general characteristics exhibited in Spiralia, such scaffold of the embryo that is precociously set up in as stereotypy and determination. Contributions the egg (Henry and Martindale 1995 , 1996a ). It has and capabilities of the embryo to form the future been shown that morphogenetic factors are evenly body parts have been most thoroughly studied in distributed within the egg prior to fertilization and the heteronemertean pilidiophoran Cerebratulus that a progressive restriction due to segregation of lacteus (Table 8.1 ; Henry and Martindale 1997 , these morphogenetic factors along the preformed 1998a ). Compared to the spiral cleavage pattern animal- vegetal axis is executed after fertilization. of annelids and mollusks, however, there are some The determinants seem to be fully segregated at the remarkable differences in terms of regulation and third cleavage division although vegetal determi- contribution of blastomeres to future organ systems nants (as exemplifi ed by formation of a larval gut) in this species (Henry and Martindale 1997 and ref- seem to be already confi ned to the vegetal pole of erences therein). The larval anterior-posterior axis the embryo after the second cleavage division, i.e., is already set up within the ovary and is outlined by at the four-cell stage (Zeleny 1904 ; Yatsu 1909 ; the stalk-like process on the vegetal pole and the Hörstadius 1937 ; Freeman 1978 ). The acceleration

Table 8.1 Clonal contributions of blastomeres during early cleavage (up to the 64-cell stage) in Nemertea Taxon Paleonemertea Heteronemertea Hoplonemertea Blastomere Contribution First quartet Apical tuft (1q1 ), ectoderm, 28 Apical tuft, larval Apical tuft, ectoderm “trochoblasts” (1q1 , 1q 2 ), eye (1c1 ) ectoderm, ciliated band, cephalic disks (1a, b), larval nervous system (1c, d) Second quartet 12 “trochoblasts,” esophagus, Larval ectoderm, ciliated Ectoderm, mesoderm primary somatoblast (2d), band, esophagus, larval (2a111 –d111 ) mesoderm a nervous system (2a, c, d) Third quartet ? Esophagus, larval Ectoderm muscles (3a, b), larval ectoderm (3c, d), larval nervous system (3c, d) Fourth quartet Mesoderm (3D)a Gut (4a–c), adult Gut mesoderm (4d) Macromeres Gut Gut Gut Species Carinoma tremaphoros Cerebratulus lacteus Malacobdella grossa References Maslakova et al. (2004a ); Henry and Martindale Hammarsten (1918 ) a Tubulanus nothus : Dawydoff ( 1998a ) (1928 ) a Marks data by Dawydoff (1928 ) for Tubulanus nothus 166 J. von Döhren or retardation of this segregation of determinants molecular mechanism underlying this delayed by stimulation or inhibition of the formation of determination is still unknown. mitotic asters during cleavage hints at a role the cytoskeleton, especially the microtubules, plays in the segregation process (Freeman 1978 ; Goldstein Cell Lineage and Freeman 1997 ). Contributions of blastomeres to the larval/adult body plan have most thoroughly been studied in Embryonic Regulative Capacities Cerebratulus lacteus for which the complete cell of Nemertea lineage is known up to the pilidium state (Table 8.1 ; Henry and Martindale 1998a ). Partial Additional remarkable modifi cations of the stereo- cell lineages are available for the paleonemertean typic spiral cleavage are seen in the capacity of the Carinoma tremaphoros and the hoplonemerteans embryo to regulate as well as in the contribution of Nemertopsis bivittata and Malacobdella grossa the blastomeres to the larval body. There is, how- (Hammarsten 1918 ; Martindale and Henry 1992 , ever, considerable difference regarding regulative 1995 ; Henry and Martindale 1994 ; Maslakova capabilities in Pilidiophora as opposed to other et al. 2004a ). In all species examined, all blasto- embryos. Embryos of Cerebratulus lacteus that meres of the fi rst quartet contribute equally to have been halved at the two-cell stage are capable forming the apical pit with a tuft of long cilia as of forming completely normal but miniature larvae, well as the majority of the larval epidermis. Due while embryos dissected at the four-cell stage do to the large size of the fi rst quartet micromeres, not regulate to complete larvae. Cleavage after iso- the ectodermal domains are positioned in a dor- lation of blastomeres is resumed in the stereotypic solateral and ventrolateral orientation, respec- spiral fashion, instead of being reinitiated from a tively, instead of being clearly dorsal, ventral, point that is analogous to earlier stages of cleavage and lateral as in other spiralian animals (see (e.g., zygote in the case of isolation at the two-cell Chapters 7 and 9 ; Henry and Martindale 1997 , stage) (Wilson 1900 , 1903; Zeleny 1904 ; Yatsu 1998a , 1999 ). In four-cell stages of Cerebratulus 1910; Hörstadius 1937 ). Embryos of the hoplone- lacteus and Nemertopsis bivittata that do not pos- mertean Nemertopsis bivittata in which blasto- sess cross-furrows, quadrant identities cannot be meres have been deleted at either the two-cell or predicted, while in Carinoma tremaphoros the A the four-cell stage develop into characteristically and C quadrants are formed by animal cross-fur- defi cient larvae, indicating that regulation does not row blastomeres, and the B and D quadrants by take place to such an extent as in Cerebratulus lac- vegetal cross-furrow blastomeres (Henry and teus (Martindale and Henry 1995 ). Compared to Martindale 1997 , 1998a ; Maslakova et al. 2004a ). other spiralian taxa, the inability to regulate as seen In Cerebratulus lacteus, the D quadrant along in Nemertopsis bivittata seems to be the ancestral with the dorsoventral axis is induced by the fi rst state and to be attributed to an early determination quartet micromeres after the third cleavage divi- of the clonal contributions of blastomeres to the sion. Deletion of all fi rst quartet micromeres juvenile tissues. results in radialized larvae, while deletion of a The regulative capacity of Pilidiophora repre- single or two adjacent fi rst quartet micromeres sents a modifi cation of the stereotypic determina- leads to the D quadrant being determined in a tion in spiral cleavage and can be attributed to the position where the fi rst quartet macromere had mode of development in which cells remain contact with most of the remaining fi rst quartet undifferentiated until forming the juvenile rudi- micromeres during the eight-cell stage (Henry ments much later in development. As a conse- 2002). In Cerebratulus lacteus the fi rst quartet quence of later determination of cell fates, the micromeres contribute to the apical epidermis cells already during cleavage retain a certain abil- down to the circumoral ring of elongated cilia ity to compensate for loss of blastomeres. The including the outer side of the lateral larval lap- 8 Nemertea 167 pets, while in Carinoma tremaphoros the fi rst 1998a ). As a characteristic of spiralian develop- quartet micromeres form the apical epidermis ment, mesoderm is derived from two sources. down to the apical row of trochoblast cells The so-called ectomesoderm is derived from (Table 8.1 ; Henry and Martindale 1998a ; micromeres of either the second or the third Maslakova et al. 2004a , b ). The second micro- quartet, while the so-called endomesoderm mere quartet cells contribute equally to most of originates from the fourth quartet micromere of the remainder of the epidermis, represented in the D quadrant, the so-called 4d mesendoblast Cerebratulus lacteus by parts of the ciliated band, (Table 8.1 ; Boyer et al. 1996 ; Boyer and Henry the inner side of the lappets, and parts of the 1998 ; Henry and Martindale 1999 ). In other esophagus (Table 8.1 ; Henry and Martindale nemertean species, there have been different 1998a ). In Carinoma tremaphoros , contributions accounts on the origin of the mesoderm. While of the second micromere quartet are unequal. In some researchers derive the mesoderm exclu- this species each of the second micromere quartet sively from ectomesodermal sources such as the cells forms three of the posterior trochoblast cells second micromere quartet (e.g., Hammarsten each and parts of the esophagus. The entire epi- 1918 for Malacobdella grossa), others claim dermis posterior to the trochoblast ring is formed both ectomesodermal and endomesodermal by the progeny of the dorsal second quartet sources to be present (e.g., second micro- micromere (2d), the so-called primary somato- mere quartet and 3D in Tubulanus nothus ; see blast (Table 8.1 ; Maslakova et al. 2004a , b ). The Table 8.1 and Dawydoff 1928 ). The mesoderm role of this blastomere is in accord to what is from both a mesendoblast (4d) and a multipolar reported from other spiralian taxa. In the hop- delamination from the archenteron after gastru- lonemertean Nemertopsis bivittata , there is no lation but without ectodermal contributions has somatoblast; all fi rst and second quartet micro- also been described (e.g., Nusbaum and Oxner meres contribute equally to the ectodermal 1913 for Lineus ruber). More recent studies on domains (Martindale and Henry 1995 ). The band the cell lineage of the pilidium of Cerebratulus of long cilia in the pilidium has to be regarded as lacteus have identifi ed the third quartet micro- not being homologous to the prototroch of the meres of the A and B quadrant as source of the typical trochophore larva. In the latter, the pro- ectomesoderm forming the array of larval mus- totroch is composed by a limited number of cells cles as well as some undifferentiated mesenchy- which originate from the fi rst and second quartet mal cells scattered in the blastocoel (Table 8.1 ; micromeres, while there are a large number of Henry and Martindale 1996b , 1998a ). It is cells in the ciliated band of the pilidium that are known from isolation experiments that blasto- additionally contributed by the C and D quad- meres isolated at the four-cell stage are capable rants of the third quartet micromeres (Maslakova of regulating for mesenchymal tissues in further 2010b ). Moreover, the ciliated marginal band development. This indicates that inductive inter- cells lack the expression of a trochoblast-specifi c actions from the A and B quadrants inhibit the β-tubulin found in other trochophores (van den formation of mesodermal cell types in the prog- Biggelaar et al. 1997 ). Due to the position of eny of their dorsally located counterparts (i.e., Pilidiophora within Nemertea and the absence of 3c and 3d; Martindale and Henry 1995 ). prototroch-type long cilia in either hoplonemer- Endomesoderm is formed by the fourth quartet tean or paleonemertean species, the long cilia micromere of the D quadrant (Table 8.1 ). In the that adorn the marginal band of the pilidium thus pilidium, endomesoderm is represented by a pair have to be regarded as newly evolved (Maslakova of loosely organized mesodermal bandlets situ- 2010b ). ated underneath the epidermis at the junction of Contributions of blastomeres after the the esophagus and stomach and some scattered, fi fth cleavage division, i.e., the 32-cell stage, undifferentiated mesenchymal cells (Henry and are known only from the pilidium larva of Martindale 1996b , 1998a ). The differentiation Cerebratulus lacteus (Henry and Martindale of adult mesoderm could not be followed but is 168 J. von Döhren assumed to originate from a population of undif- tive potential located in two adjacent quadrants ferentiated, dormant mesenchyme cells (Henry necessary to induce eye formation (Martindale and Martindale 1997 ). In paleonemertean and and Henry 1995 ; Henry and Martindale 1997 ). hoplonemertean larvae, assumed cell-cell inter- Although there is considerable variation with actions as operating during the development respect to some blastomere lineages in nemerte- of the pilidium to specify the dorsoventral axis ans, their mode of cleavage clearly operates within clearly do not inhibit that all quadrants contribute the framework of a spiral cleavage as seen in many equally to the muscle layers displayed in the lar- other spiralians. In contrast to most other spiralian vae (Iwata 1957 ; Martindale and Henry 1995 ). To species, relative blastomere size, inductive interac- what extent the musculature is derived from both tions, and, to a certain degree, regulation seem to ectomesodermal and endomesodermal sources play a prominent role in nemertean development in nonpilidial types of development has been a (Henry and Martindale 1999 ; Henry 2002 ). matter of debate and remains to be clarifi ed. The ectodermally derived nervous system of the pilid- ium of Cerebratulus lacteus is formed by progeny Gastrulation of the blastomeres 1c, 1d, 2a, 2c, 2d, 3c, and 3d with all of them contributing to the marginal cili- Accelerated divisions of the progeny of the ani- ary neuropil (Table 8.1 ). The oral and the suboral mal quartets and subsequent fl attening of the cells neuropils are formed without contribution of the lead to the widening of the blastocoel into which fi rst quartet micromeres (Henry and Martindale the more columnar vegetal progeny projects to 1998a ). Due to the nervous system generating different degrees. This process results in a coelo- blastomeres being situated mostly in dorsal posi- blastula in nearly all species studied, whereas the tions in the embryo, the authors suggested induc- shape of the blastula and the dimensions of the tive interactions of other blastomeres specifying blastocoel may vary. The blastula may be domed, neuronal fates of blastomeres in the pilidium as but in the majority of species, it is fl attened, has been shown in D quadrant specifi cation. resulting in an either rather spacious or highly There are no data available on the cell lineage compressed blastocoel (Friedrich 1979 ). While of the nervous system in paleonemerteans and the blastula is radially symmetric in most species, hoplonemerteans. The only sensory structures it attains a somewhat rectangular appearance that provenance from blastomeres is known of (having been termed blastosquare) due to some are the eyes of the monostiliferan hoplonemertean of the progeny of animal blastomere quartets jut- Nemertopsis bivittata and the carinomid paleone- ting out to the sides in Cephalothrix rufi frons , mertean Carinoma tremaphoros (Martindale and Malacobdella grossa , and Micrura alaskensis Henry 1995 ; Maslakova et al. 2004b ). Although (Hammarsten 1918 ; Smith 1935 ; Maslakova apparently different with respect to both number 2010a ). At this time of development, some spe- and ultrastructure (one pair of rhabdomeric eyes in cies show signs of beginning ciliation, while in the monostiliferan hoplonemertean Paranemertes other species ciliation only starts when gastru- peregrina versus a single ciliary eye in the carino- lation sets in. Some species with intracapsular mid paleonemerteans Carinoma tremaphoros and development develop cilia even later after gas- Carinoma mutabilis ), there seems to be a com- trulation (e.g., Pantinonemertes ( Geonemertes ) mon pattern in deriving at least one eye from the C australiensis , Antarctonemertes phyllospadic- quadrant (1c1 in Carinoma tremaphoros , right eye ola ) (Hickman 1963 ; Maslakova and von Döhren in Nemertopsis bivittata from so-called RD, i.e., C 2009 ). The apical pole in some species with quadrant; see Martindale and Henry 1995 ; Henry pelagic stages already shows fi rst rudiments of and Martindale 1997 , 1999 ; Maslakova et al. organs, namely, the apical pit characterized by 2004a ). The second eye in Nemertopsis bivittata is a group of columnar cells that are housed in a derived from the oppositely located (i.e. A) quad- shallow depression. In species with intracap- rant. In the latter species, there seem to be induc- sular development, an apical organ rudiment is 8 Nemertea 169 usually absent (Friedrich 1979 ; Senz and Tröstl so-called indirect development is characterized by 1999 ). A pair of large, sometimes slightly invagi- a catastrophic metamorphosis in which the larval nated cells situated on both sides of the apical pit epidermis is shed as a whole and usually eaten by has been reported in Procephalothrix simulus , the juvenile that has developed inside (Cantell Emplectonema gracile , Prosorhochmus vivipa- 1967; Maslakova 2010a). Larval types in rus , and Malacobdella grossa , already marking Pilidiophora are quite diverse, including several the bilateral symmetry of the blastula (Friedrich morphotypes of pilidia, the Desor larva of Lineus 1979 ). The large cells have been interpreted as viridis , the Schmidt larva of Lineus ruber , the the fi rst rudiments of the nervous system (Iwata Iwata larva of Micrura akkeshiensis, and other 1960 ). On the vegetal pole of the embryo, one unspecifi ed pelagic, lecithotrophic larvae of some ( Cerebratulus lacteus: Coe 1899 ; Wilson 1903 ; Micrura species (Norenburg and Stricker 2002 ; Lineus ruber : Nusbaum and Oxner 1913 , but Schwartz and Norenburg 2005; Schwartz 2009 ; see Schmidt 1964 ; Tubulanus nothus : Dawydoff Maslakova 2010b ). Larvae of hoplonemertean and 1928) or two pairs ( Procephalothrix fi lifor- paleonemertean species have traditionally been mis : Iwata 1960 ; Tetrastemma vermiculus and classifi ed as so-called planuliform larvae due to Drepanophorus spectabilis: Lebedinsky 1897 ) their superfi cial resemblance to the planula larvae of distinct blastomeres, having been interpreted of cnidarians (Fig. 8.6A–C ). Several hypotheses as primary mesoblast cells, are reported. In other have been put forward to derive the pilidium from species, however, the vegetal-most blastomeres planuliform nemertean or even other, non-nemer- are indistinguishable in size and form (Friedrich tean larval types, but the evolution of this aberrant 1979 ). larval form is presently still enigmatic (Iwata Gastrulation proceeds by invagination of the 1972 ; Jägersten 1972 ; Hiebert et al. 2010 ). blastomeres of the vegetal pole, although epi- Recent fi ndings, however, hint at a more com- bolic processes caused by the proliferation of the plex picture. The planuliform larval type includes ectodermal components have been indicated to the decidula larva that possesses a transitory larval be involved. During gastrulation the shape of the epidermis whose cells are replaced by cells of the embryo changes to become more domed. Some defi nite juvenile epidermis and the hidden trocho- species, especially hoplonemerteans with very phore that is characterized by a distorted preoral yolky eggs, gastrulate by polar ingression of the belt of large cells that have been homologized with vegetal cells (Malacobdella grossa, Prostoma grae- the trochoblasts of annelids and mollusks cense, Argonemertes ( Geonemertes) australien- (Maslakova et al. 2004b; Maslakova 2010b ). The sis, Gononemertes australiensis) (Hammarsten decidula is typically present in Hoplonemertea but 1918; Reinhardt 1941; Hickman 1963 ; Egan and has also been suggested to occur in the tubulanid Anderson 1979 ). paleonemertean Tubulanus punctatus (Iwata 1960 ; Maslakova 2010b and references therein). The hidden trochophore has been found in LATE DEVELOPMENT Carinoma tremaphoros based on cell lineage stud- ies (Maslakova et al. 2004a , b ). The larval type of Diversity of Larval Types in Nemertea the remaining paleonemertean species has never been specifi ed. Species with intracapsular devel- Although most nemerteans develop via planktonic opment typically show characteristics of the larval stages, their development has traditionally been type present in their respective phylogenetic lin- characterized as direct and indirect development, eage (Maslakova and Malakhov 1999 ; Maslakova according to the transformation of the larva to and von Döhren 2009 ; von Döhren 2011 ). Larvae form the juvenile body (Fig. 8.6). While in nemer- of hoplo- and paleonemerteans develop from the tean, so-called direct development morphological gastrula by differential growth of one side of the transformations are gradual, resulting in a smooth body which becomes the dorsal side of the animal. transition from the pelagic stage to the juvenile, As a consequence, the blastoporal region moves to 170 J. von Döhren

Fig. 8.6 Pelagic larvae of Nemertea. (A ) Five-day-old larva pair of dorsal eyes. The stylet apparatus has already formed of Carinina ochracea (Paleonemertea), fi xed and osmi- inside the larva. (D ) Six-week-old, advanced pilidium of cated specimen¸ differential interference contrast (DIC ) light Riseriellus occultus (Pilidiophora), live specimen, DIC. Note micrograph. Note the single median ventral eye. (B ) Three- the advanced juvenile with eyes inside the pilidium. The ima- day-old larva (at 18 °C) of Procephalothrix oestrymnicus ginal disks are fused; only the dorsal part of the juvenile has (Paleonemertea), live specimen, DIC. Note the pair of dor- yet to form. at apical tuft, ct caudal tuft, de defi nite (adult) eye, sal eyes. (C ) Four-day old larva of Emplectonema gracile jr juvenile rudiment, la lateral lappet, le larval eye, mo mouth (Hoplonemertea), live specimen, bright fi eld. Note the fi rst opening, sty stylet (© Dr. J. von Döhren, All Rights Reserved) a more frontal position marking the ventral side of sequent researchers (e.g., Bürger 1894 ; Dawydoff the larva, thus changing the angle between the api- 1940 ; Cantell 1969 ; Chernyshev et al. 2013 ). More cal and the former vegetal pole of the embryo. than half a dozen morphotypes have been described The pilidium was originally described by Müller of which the original pilidium gyrans represent the (1847 ) as a previously unknown, putatively larval archetype but not necessarily the ancestral state animal from the North Sea. It was consequently (Figs. 8.6D and 8.7 ). Apart from different shapes given a binomen: pilidium gyrans. Although being of the lappets, the dome, and the bands of elongated aware of the larval nature of the organism, the tradi- cilia, the pilidial types have been distinguished by tion of binomial nomenclature was adopted by sub- the orientation that the juvenile rudiment assumes 8 Nemertea 171

both of which are functional. In the majority of hoplonemertean species, the blastopore has been reported to close. A secondary, ectodermal mouth mid at opening along with an esophageal tube is subse- quently formed in various ways according to dif- hlb es ferent authors (Friedrich 1979 ; Maslakova and flb von Döhren 2009 ). In some hoplonemertean spe- cies, especially those with intracapsular devel- opment, no functional mouth opening is formed until much later in development (Salensky 1914 ; mcb Hammarsten 1918 ; Hickman 1963; Iwata 1960 ; td crd cpd Egan and Anderson 1979 ). The larvae are ini- tially ovoid in shape and gradually elongate with age. While the surface of the larva is covered with cilia of equal length, the apical pole of the larva lap 100 µm is equipped with an apical tuft of longer cilia housed in an apical depression, the apical pit. In Fig. 8.7 Schematic representation of a young pilidium many species the posterior end of more advanced gyrans (lateral view). The positions of the prospective stage larvae is endowed with another tuft of elon- paired epidermal invaginations forming the juvenile are shown in shaded areas (black , exumbrellar invagination; gated cilia termed caudal tuft or posterior cirrus light grey , subumbrellar invagination). at apical tuft, cpd (Fig. 8.6C ; Iwata 1960 ; Stricker and Reed 1981 ; cephalic disk rudiment, crd cerebral organ disk rudiment, Maslakova and von Döhren 2009 ). One to several fl b forelobe of pilidium, hlb hindlobe of pilidium, lap lat- pairs of lateral cirri located anterior of and at the eral lappet of pilidium, mcb marginal band of elongated cilia, mid midgut, es esophagus, td trunk disk rudiment level of the mouth opening have been described (© Dr. J. von Döhren, All Rights Reserved) for members of the cephalothricids (Iwata 1960 ). In most larvae a pair of epidermal invaginations with respect to the larval apical-vegetal axis. In the on either side of the apical pit is formed which archetypical pilidium (e.g., pilidium gyrans), the has variously been interpreted as rudiments juvenile anterior-posterior axis assumes a roughly of the nervous system or the cerebral organs rectangular orientation to the larval apical-vegetal (Lebedinsky 1897 ; Salensky 1909 ; Hammarsten axis as indicated by the larval apical tuft and veg- 1918 ; Smith 1935 ; Iwata 1960 ; Maslakova and etal mouth opening. There is a hypothetical series von Döhren 2009 ; but see Hiebert et al. 2010 ; to derive the archetypical pilidial forms (pilidium Maslakova 2010b ). gyrans, pilidium pyramidale) from types in which Some larvae of paleonemerteans and most the larval and juvenile main axes are roughly par- hoplonemertean larvae possess eyes (Fig. allel, such as the pilidium recurvatum (Jägersten 8.6A–C ). They are usually simple pigment 1972 ). However, more recent fi ndings indicate that cup ocelli composed of only a few photore- the vast diversity of pilidial morphotypes might be ceptor cells surrounded by one to a few shad- a consequence of the uncoupling of morphogenetic ing pigment cells (von Döhren 2008 ). While in processes between the larval stage and the forma- Carinoma species and Carinina ochracea there tion of the juvenile (Hiebert and Maslakova 2015 ). is a single almost median ventral eye anterior to the mouth opening, cephalothricids and most hoplonemertean species possess a pair of eyes Comparative Larval Development situated dorsally, in front of the mouth opening (Fig. 8.6A–C; Hammarsten 1918; Iwata 1960 ; Paleo- and Hoplonemertean Larvae Maslakova et al. 2004a , b ; Maslakova and von In larvae of paleonemertean species, the blas- Döhren 2009 ; Hiebert et al. 2010; Maslakova toporal region differentiates into an ectodermal 2010b ). In the larva of Quasitetrastemma esophagus and an endodermal, sac-like midgut, ( Tetrastemma ) stimpsoni and Quasitetrastemma 172 J. von Döhren

( Tetrastemma ) nigrifrons , three pairs of dorsally through the esophagus to the gut (Maslakova located eyes have been described (Chernyshev 2010a). The larva is composed of two epithelial 2008 ). In paleonemertean larvae, the eyes are layers consisting of a single row of multiciliated, epidermal and possess ciliary receptor cells, large fl at cells. In the region of the apical tuft, the while hoplonemertean species possess subepi- marginal ciliary band, and the longitudinal ridges, dermal eyes equipped with rhabdomeric recep- the cells attain a more columnar shape. Inside the tors. Since the adults of those paleonemertean larva, an extensive, comparably loose extracellu- species studied lack eyes, the eyes present in the lar matrix with interspersed neuronal and meso- larvae have to be interpreted as transitory larval dermal cell clusters fi lls the body. organs (von Döhren 2008 ). In hoplonemerte- Both the larval musculature and nervous sys- ans, the subepidermal eyes persist to adulthood, tem are formed early in development, enabling increasing in size and number in most species. the larva to take up food particles and to loco- Only in Quasitetrastemma species the one pair mote. For the most part, nerves and muscles are of eyes has been hypothesized to be fused to the not taken over into the juvenile organization neighboring pair or completely reduced during (Maslakova 2010a ), but some neural components development (Chernyshev 2008 ). are incorporated into the postmetamorphic ner- vous system (Hindinger et al. 2013). The fi rst Pilidiophoran Larvae: The Pilidium structures of the nervous system that differenti- The archetypical pilidium develops from the ate in the pilidium are a prominent ring-shaped swimming gastrula by growth of the lateral rims neuropil underlying the marginal ciliary band of the invaginated vegetal fi eld to form the lateral (marginal band neuropil), a ring-shaped neuro- lappets. A ring of elongated cilia, the marginal pil encircling the opening of the esophagus into ciliary band, is present around the invaginated the gut (oral ring neuropil), as well as a pair of ciliary fi eld, marking border between the outer neurite bundles connecting the oral ring neuro- (exumbrellar, also: epispheric) and the inner pil on both sides of the larva with the marginal (subumbrellar, also: hypospheric) surfaces of the band neuropil at the posterior base of the lateral larva (Fig. 8.7 ; Friedrich 1979 ; Maslakova lappets. A pair of linear plexuses (circumesopha- 2010a ). In some forms of pilidia, the anterior and geal plexus) extending from the anterior to the posterior parts of the ciliated ring may form addi- posterior base of the lateral lappets alongside the tional anterior and posterior lobes in the larva. opening of the esophagus connecting to the mar- Denticle-like structures and chromatophores may ginal band neuropil has been described in some adorn the ciliated ring. Depending on the species, species (Lacalli and West 1985 ; Hay-Schmidt they may increase in number with age of the larva 1990 ; Henry and Martindale 1998a ; Maslakova (Cantell 1969; Norenburg and Stricker 2002 ; 2010a ; Zaitseva and Flyachinskaya 2010 ; Lacalli 2005 ). The exumbrellar part, housing the Hindinger et al. 2013 ). The epidermis is under- apical pit with a tuft of elongated cilia, expands lain by a loosely arranged epidermal plexus that to form the typical dome shape of the pilidium is in contact with the marginal band neuropil and (Figs. 8.6D and 8.7). The esophagus is formed exhibits a prominent pair of monociliated, seroto- from ectodermal cell material being dragged in nergic, putatively sensory neurons, laterally next during blastopore invagination (Friedrich 1979 ). to the apical pit (Maslakova 2010a ; Hindinger The larval gut is sac-like and located interior of et al. 2013). The apical pit itself is devoid of the posterior lobe. The gut is open to the esopha- neuronal elements (Cantell et al. 1982 ; Lacalli gus via a narrow opening with a valve that devel- and West 1985 ; Chernyshev et al. 2013 ). From ops from the blastopore. Along the posterior wall some of the mesodermal cells, the larval mus- of the esophagus, there is a pair of prominent culature forms underneath the epidermis, while longitudinal ridges that exhibit elongated cilia. others remain in an undifferentiated, almost They have been reported to serve as main struc- amoeboid state (Henry and Martindale 1998a ; tures to mediate transport of food particles Maslakova 2010a). The most prominent structure 8 Nemertea 173 of the larval musculature is a muscle ring situated quartet micromeres, hence 1a and 1b (Fig. 8.7 ; underneath the marginal ciliary band. The muscle Henry and Martindale 1998a ; Maslakova 2010a ). ring branches at the bases of the lateral lappets After some time, the trunk disks are formed, fol- to send a portion along the rim of the lappets, lowed by the cerebral organ disks. Both pairs while the other encircles the esophageal open- of imaginal disks are unanimously reported to ing. A ring-shaped sphincter muscle underlies invaginate from the subumbrellar larval epider- the opening between the esophagus and the gut. mis (Fig. 8.7 ; Friedrich 1979 ; Maslakova 2010a ). Underneath the epidermis there is a prominent Shortly after, an unpaired anterior-most putative array of muscles that radiate from the dome into proboscis rudiment is observed sitting dorsal of all parts of the larva. By branching they form an the cephalic disks underneath the larval epider- oblique muscle network being most prominent in mis. The exact origin of the putative proboscis the lateral lappets (Maslakova 2010a). A muscle rudiment is unknown. It has been described as extending from the apical pit downward (apical epidermal invagination similar in structure to the muscle or central muscle), branching around the paired imaginal disks, as a delamination of the esophagus, and serving to contract the larval apex larval epidermis or as a cluster of undifferenti- has been described in most of the larval forms ated mesenchymal cells forming underneath the (Bürger 1897 –1907; Lacalli 2005 ; Zaitseva and larval epidermis (Bürger 1894; Schmidt 1937 ; Flyachinskaya 2010 ). Maslakova 2010a). At the time the cephalic disks The juvenile develops inside the pilidium fuse with each other, incorporating the putative from a set of epidermal invaginations of the proboscis rudiment and thus forming the head larval envelope, termed imaginal disks. A mini- rudiment, the anterior end of each trunk disk fuses mum of three paired, bilaterally located disks are with the posterior rim of the cerebral organ disk of described: the most frontally located cephalic the respective side. During fusion of the proximal disks, the middle cerebral organ disks, and the cell layer of the imaginal disk, the amnion layers trunk disks situated ventral of the gut posterior are separated from the proximal layer to fuse to of the lateral lappets (Fig. 8.7 ). After invaginat- each other, forming a continuous amniotic layer ing from the larval epidermis, the imaginal disks distal to the respective rudiments. In most spe- are pinched off becoming fl attened, fl uid-fi lled cies an additional, unpaired, posterior- most dor- cavities. While the cells of the proximal wall of sal rudiment forms (Friedrich 1979 ; Maslakova the imaginal disk are initially more columnar and 2010a ). In contrast to the paired imaginal disks, differentiate into defi nite, juvenile epidermis, the the dorsal rudiment arises as a delamination from distal layer retains its fl at, thin structure reminis- the larval epidermis or a mesodermal cell layer cent of the larval epidermis. The distal, thin cell instead of an invagination from the larval epider- layer is referred to as amnion (Friedrich 1979 ; mis. Although initially single layered, the dorsal Maslakova 2010a). In addition to the three pairs rudiment soon becomes double layered, the outer of epidermal invaginations, two rudiments have layer forming an amnion as is seen in the ima- been observed, of which the ectodermal origin ginal disks that develop as invaginations of the has not been unambigouously demonstrated: the larval epidermis (Bürger 1894 ; Salensky 1912 ; anterior-most proboscis rudiment and the poste- Schmidt 1937 ; Maslakova 2010a). During further rior-most dorsal rudiment, both of which might development the dorsal rudiment fuses with the not be generally present in all species (Friedrich dorsoposterior rims of the bilaterally still sepa- 1979 ; Maslakova 2010a ). The imaginal disks do rate trunk rudiments which themselves fuse pos- not, however, form at the same time, but there is a teriorly soon afterwards. As well as during the stereotypic sequence of appearance. The fi rst pair fusion of the cephalic disks, a continuous amnion of disks that forms is the cephalic disks. Unlike layer is formed. the remaining imaginal disks, the cephalic disks The next stage is accomplished by the fusion are invaginated from the exumbrellar epidermis of the lateral posterior rims of the head rudiment as derivatives of the A and B quadrants of the fi rst to the lateral anterior rims of the trunk rudiment 174 J. von Döhren represented by the cerebral organ disks. Thus, of developing at an angle to the larval body, the a continuous torus-shaped rudiment is formed juvenile inside is parallel to the larva but with around the larval esophagus (Maslakova 2010a ). its head directed toward the side of the blasto- The anterior dorsal rim of the trunk rudiment sub- pore. Furthermore, in the Iwata larva, the number sequently fuses with the posterior dorsal margin of imaginal disks is restricted to fi ve, a pair of of the head rudiment, completing the formation frontal head disks, a pair of posteroventral trunk of the juvenile inside the pilidium. The juvenile disks, and an unpaired posterior dorsal disk, all of is almost completely separated from the larval which form from invaginations of the larval epi- tissues by the amniotic cavity which is lined by dermis. An anterior-posterior sequence of their the continuous amnion layer. The only contact of formation is likely (Iwata 1958 ). pilidium and juvenile is by the mouth opening that Similar nonfeeding larval types have been is shared by larva and juvenile. After the develop- described in Micrura rubramaculosa , Micrura ment of the juvenile inside the pilidial envelope is verrilli, and at least two other yet undescribed completed, the juvenile escapes from its envelope Micrura species; although in the larva of the by vigorous movements that rupture the pilidial former species there is an additional equato- tissues (Cantell 1967 ; Maslakova 2010a ). This rial band of elongated cilia present and the lar- process has been termed catastrophic metamor- val animal- vegetal axis coincides with the adult phosis and is usually a matter of a few minutes. anterior-posterior axis (Schwartz and Norenburg The larval envelope that is still connected to the 2005; Schwartz 2009). Another nonfeeding lar- mouth of the escaping juvenile is usually eaten val form possessing an equatorial as well as a by the juvenile (Cantell 1967 ; Maslakova 2010a ). posterior band of elongated cilia that have been termed proto- and telotroch analogously to the Aberrant Pilidiophoran Larvae ciliated bands of the trochophore larva has been Within Pilidiophora there seems to be an evolu- described for an undescribed lineid heterone- tionary tendency toward lecithotrophic stages mertean (Maslakova and von Dassow 2012 ; (Maslakova and Hiebert 2014 ). The larva of Maslakova and Hiebert 2014 ). Unfortunately, Micrura akkeshiensis, named Iwata larva after its there is virtually nothing known on the develop- discoverer, is a pelagic stage but does not take up ment in any of the abovementioned larval types. food during its development (Iwata 1958 ). Thus, Although being the fi rst developmental stage the larva does not show either lappets or bands of described in nemerteans, the Desor larva of elongated cilia used for food capture in the pilid- Lineus viridis might represent one of the most ium. It develops from a comparably fl at gastrula derived larval types. Despite of showing entirely that elongates and attains a pyriform shape. The intracapsular development within an egg cap- animal pole is marked by an apical pit with an sule, it shows distinct characteristics typical of apical tuft of elongated cilia. The opposite vege- pilidiophoran development, including imaginal tal pole is marked by the site of the blastopore. In disks and the devouring of the larval epidermis this type of development, the blastoporal opening during a catastrophic metamorphosis (Friedrich is not shifted to either side of the gastrula during 1979 ; von Döhren 2011 ). However, since this further development but remains in its original larval type is lecithotrophic, no food is taken position. Around the blastopore four larger, up until metamorphosis; hence, accessory feed- mesodermal cells have been observed which pro- ing structures as seen in the pilidium are absent. liferate to form a mesodermal mass between the Following gastrulation a spherical larva is formed epidermis and the invaginated archenteron. that is uniformly ciliated, lacking any apical tuft Apart from an apical tuft and an evenly ciliated or band of elongated cilia. Reminiscent of the epidermis, no distinct larval organs have been pilidium larva, three pairs of imaginal disks are reported. Similar to the pilidium, the epidermis formed by invagination along with two rudi- covering the larva is a transitory larval envelope ments formed by delamination from the larval in which the juvenile develops. However, instead epidermis (Nusbaum and Oxner 1913 ; Schmidt 8 Nemertea 175

1964). They correspond in position to the ima- Riserius (Cantell 1969 ; Thollesson and ginal disks and rudiments seen in the pilidium. Norenburg 2003 ; Hiebert et al. 2013a , b ). In contrast to canonical pilidial development, An apical organ situated basal to the api- cephalic and trunk disks appear almost simulta- cal pit present in all nemertean larval types neously in the Desor larva, followed by the pro- has variously been interpreted as homologous boscis rudiment, the cerebral organ disks, and to the apical organ in several trochozoan and fi nally the dorsal rudiment (Schmidt 1964 ). The lophophorate species, especially in Annelida sequence of fusion only differs from that seen in and Mollusca but also in Brachiopoda (see the pilidium in that the trunk disks fi rst fuse with Chapters 7 , 9 , and 12 as well as Nielsen 2013 each other and with the dorsal rudiment before and references therein). In many spiralian lar- fusing with the cerebral organ disks (Friedrich vae, the apical organ is composed of the apical 1979 ). At that time, the larva elongates posteri- tuft and a usually low number of fl ask-shaped, orly, attaining a rhomboid shape. serotonergic neurons at its base. These fl ask- The larva of the closely related species Lineus shaped neurons generally degenerate prior to ruber is smaller but has essentially the same metamorphosis (Nielsen 2013 and references larval morphology (Schmidt 1964 ). It has been therein). In nemertean species a single fl ask- given a different name due to its reported diver- shaped serotonergic neuron has been reported gent mode of feeding. In contrast to the Desor only for hoplonemertean species although it larvae of Lineus viridis , the so-called Schmidt does not project directly into the apical pit but larva of Lineus ruber is adelphophagic, feed- reaches the epidermal surface in its vicinity ing on putatively defi ciently developing sibling (Fig. 8.8A ; Chernyshev and Magarlamov 2010 ). embryos inside the same egg capsule. Hence, Moreover, this fl ask-shaped neuron persists differences in timing of the development of the through metamorphosis as part of the brain ring proboscis and the alimentary tract with its associ- (von Döhren J 2015, unpublished). In paleone- ated musculature in later-stage larvae have been mertean and pilidiophoran species, no fl ask- reported (Schmidt 1964 ). shaped serotonergic neurons have hitherto been found. In the pilidium, there are two spherical, Nemertean Larval Types – Evolutionary putatively sensoric serotonergic neurons in the Considerations vicinity of the apical pit, while the single apical Including the other nemertean larval types, there serotonergic neuron of paleonemertean species is a series of transitions from the paleonemer- has an ovoid shape and does not make contact tean larvae to the most derived developmental with the epidermis (Fig. 8.8B ). While the ovoid types, namely, the Iwata and Desor larva (Iwata apical serotonergic neuron in Paleonemertea dis- 1972 ). This series, however, has to be taken with appears when the brain rudiment starts to form, some reservation. On the one hand, the basally the apical serotonergic neurons of Pilidiophora branching, non-heteronemertean pilidiophoran are shed at metamorphosis along with the lar- Hubrechtella dubia possesses a pilidium of the val envelope. Similar to the situation seen in the pilidium auriculatum type which is most similar larvae of polyclad Platyhelminthes, homology to the pilidium gyrans type (Cantell 1969 ), of the serotonergic apical neurons is unclear while on the other hand, most pilidial morphot- (Rawlinson 2010). Two evolutionary scenarios ypes cannot be assigned to a certain species or seem possible: either the apical serotonergic clade, thus rendering phylogenetic inferences neurons of Nemertea are homologous to those needed to set up an evolutionary series impos- in Trochozoa, but there have been various sub- sible. The aberrant pilidium recurvatum type stantial transformations in both Paleonemertea has been assigned to members of the heterone- and Pilidiophora, or apical serotonergic neu- mertean pilidiophoran Baseodiscidae but has rons have evolved multiple times independently also been identifi ed as the larval type of the in Nemertea. Current hypotheses regarding putatively most basal heteronemertean genus the phylogeny of Nemertea favor the second 176 J. von Döhren

Fig. 8.8 Stereo pair images of confocal microscopy 12 °C) stained with antibodies against serotonin stacks of neural staining of nemertean larvae. (A ) (5HT-lir) and acetylated α-Tubulin (ac α-Tub-lir). Note: Emplectonema gracile (Hoplonemertea) , one-day-old there is no connection of the apical serotonergic neuron larva stained with antibodies against serotonin ( asn) to the apical pit that the apical tuft (at ) emanates (5HT-lir). Note: the fl ask-shaped apical serotonergic from. The rudiment of the brain (br ) is visible as well as neuron ( asn ) projects in the vicinity of the apical pit a few more posteriorly located neurons (psn ) showing ( ap). An additional, more posterior serotonergic neuron 5HT-lir. ap apical pit, asn apical serotonergic neuron, ( psn ) is visible at this stage. (B ) Procephalothrix oes- at apical tuft, br brain rudiment, psn posterior seroto- trymnicus (Paleonemertea) , three-day-old larva (at nergic neuron (© Dr. J. von Döhren, All Rights Reserved) alternative scenario as being more parsimonious so that generalizations are at best preliminary at (Andrade et al. 2012 ). this point in time. Comprehensive information is restricted to classical accounts on about a dozen species, most of which are neonemerteans with Formation of the Adult Body Plan arguably aberrant, i.e., intracapsular, develop- ment (e.g., Lineus viridis / ruber : Barrois 1877 ; Data on the formation of the defi nite adult orga- Arnold 1898 ; Hubrecht 1885 , 1886 ; Nusbaum nization in Nemertea are rather heterogeneous and Oxner 1913; Schmidt 1964 ; Prosorhochmus 8 Nemertea 177 viviparus : Salensky 1882 –1883, 1909 , 1914 ; development of Pantinonemertes ( Geonemertes ) Pantinonemertes ( Geonemertes ) agricola : Coe agricola (Coe 1904). Although it has been argued 1904 ; Prostoma graecense : Reisinger 1926 ; to be homologous to the pilidial larval envelope, Reinhardt 1941 ; Argonemertes (Geonemertes ) the larval epidermis of Hoplonemertea has been australiensis: Hickman 1963 ; Prosorhochmus given the name “decidula” due to its differing adriaticus: Senz and Tröstl 1999 ). Organs that fate (Maslakova 2010b ). are undisputedly of ectodermal origin are the epi- A fi rst generation of epidermal cells in dermis with the associated cephalic glands, the paleonemertean species has been hinted at in nervous system and its associated sense organs, Tubulanus punctatus (Iwata 1960), while in the outer (when seen extruded) epithelium of the Carinoma tremaphoros , there is a population of proboscis, and the rhynchodeum. The esophagus transitory trochoblast cells forming a distorted and anus have been reported to be ectodermal preoral belt that has been termed “vestigial pro- derivatives by the majority of authors although totroch” (Maslakova et al. 2004b ). The defi nite, there has been disagreement regarding the ori- adult epidermis consists of multiciliated cells, gin of the esophagus according to older accounts several types of gland cells, basal granular cells, (Barrois 1877 ; Salensky 1886 , 1909 , 1912 , and basal neurites originating from subepider- 1914 ; Arnold 1898 ). Of mesodermal origin are mal neurons constituting a pseudostratifi ed epi- the muscular systems, comprising the various thelium in paleonemertean and heteronemertean body wall muscle layers, the muscle layers and species (Turbeville 1991 ). In Hoplonemertea, due the inner epithelium of the proboscis facing the to the presence of a basal cup cell layer, the pseu- rhynchocoel, the rhynchocoel epithelium and its dostratifi ed appearance of the epidermis is even underlying muscle layers, as well as the endo- more pronounced (Norenburg 1985 ; Turbeville thelialized blood-vascular system. The intesti- 1991 ). The heteronemertean dermis is formed nal tract exclusive of the esophagus and anus is late in development, i.e., after metamorphosis, derived from the endoderm (Fig. 8.2A, B ). On by some gland cells sinking underneath the level the origin and formation of the protonephridia of the ciliated epidermis cells. Thus, proximal of in Nemertea, there has been considerable dis- the epidermal basal extracellular matrix, a dis- agreement as comprehensive data are presently tinct layer of differing extent containing gland missing. cells intermingled with muscle cells and neurons appears to be present at light microscopic resolu- Epidermis tion (Norenburg 1985 ). In Neonemertea there are two generations of epidermal cells that appear sequentially and Nervous System of which only the second is kept as the defi nite The nervous system in paleo- and hoplonemer- adult epidermis. The fi rst generation of cells tean species seems to have two areas of origin. covers the entire body as a larval epidermis in While the fi rst nervous elements represented by all Pilidiophora. It has been shown to be present a peripheral plexus are most likely derived from in Hoplonemertea according to more recent cells that sink in from the apical and the poste- fi ndings (Maslakova 2010b ). While the larval rior ectoderm of late gastrulae of paleonemertean epidermis is kept as intact larval envelope in and hoplonemertean species, the brain and lateral which the juvenile develops until metamorpho- medullary cords have been reported to origi- sis in Pilidiophora, in Hoplonemertea cells of nate from paired large ectodermal blastomeres the larval epidermis are intercalated by defi nite bilateral to the apical pit (Salensky 1909 , 1914 ; epidermal cells and later disappear (Maslakova Hammarsten 1918 ; Smith 1935 ; Iwata 1960 ). The and Malakhov 1999 ; Maslakova and von Döhren brain primordia sink underneath the epidermis, 2009 ; Maslakova 2010b ). In Pantinonemertes fi rst forming a pair of narrow epidermal invagi- californiensis larval epidermal cells have been nations that soon lose contact to the epidermis shown to be shed (Hiebert et al. 2010), a mecha- and proliferate (Coe 1904 ; Salensky 1909 , 1914 ; nism that has also been suspected to occur in the Hammarsten 1918 ; Smith 1935 ; Iwata 1960 ). 178 J. von Döhren

The different positions of the brain and lateral sural tract. Nervous structures innervating the medullary cords in different clades have been mouth opening and the esophagus are observable reported to originate from the depth to which the already in larval stages of paleonemertean spe- nervous system rudiments sink in. Remaining cies. In Cephalothrix rufi frons the esophageal in a distal position to mesodermal tissues in nerves have been derived from neuron precursors Tubulanus punctatus , the nervous system rudi- in the vicinity of the mouth opening, while in ments sink in underneath the mesodermal tissues Carinina ochracea esophageal nerves are puta- in Procephalothrix species and Hoplonemertea tively outgrowths of the lateral medullary cords (Iwata 1960 ). From the invaginated cell clusters, (Smith 1935 ; von Döhren J 2015, unpublished). the ventral and dorsal brain lobes on either side While paired ectodermal invaginations have been are formed that are later connected horizontally interpreted as the main rudiment of the brain and by the ventral and dorsal commissural tracts lateral medullary cords in most species, there (Coe 1904; Salensky 1909 , 1914 ; Hammarsten have been differing accounts in a few species 1918; Smith 1935; Iwata 1960 ). Formation of the from all clades. brain lobes, lateral medullary cords, and ventral In the intracapsularly developing species commissural tract prior to the dorsal commis- Argonemertes (Geonemertes ) australiensis and sural tract has been reported for the monostilifer- Prostoma graecense , the fi rst nervous system ous hoplonemertean Prosorhochmus adriaticus rudiments are represented by a cluster of cells (Senz and Tröstl 1999 ). In Pilidiophora, the that is not formed by an invagination (Reinhardt brain and lateral medullary cords are formed 1941 ; Hickman 1963 ), while in Cephalothrix independent of the extensive larval nervous sys- rufi frons paired groups of cells from the ecto- tem (Salensky 1886 , 1912; Maslakova 2010a ; derm in the vicinity of the stomodeum have been Hindinger et al. 2013 ). The fi rst nervous sys- suspected to contribute to the ventral brain lobes tem rudiment is observable in pilidia as a paired (Smith 1935 ). In Tetrastemma vermiculus and epidermal invagination or delamination on the Drepanophorus spectabilis , the brain is reported proximal side of either of the cephalic disks prior to originate from a ventrolateral and a dorsolat- to their fusion (Salensky 1886 , 1912). With the eral pair of epidermal thickenings that disconnect fusion of the disks, the brain lobes on either side from the epidermis to each form a brain lobe. are secondarily connected by formation of both Connection of the isolated brain compartments the ventral and later the dorsal commissural tract is later accomplished by secondarily forming (Salensky 1912 ). neuroectodermal ridges (Lebedinsky 1897 ). The In the Desor larva, the brain is formed later same mode of formation has been stated for the in development when the cephalic disks have lateral medullary cords in the abovementioned already fused with each other and with the trunk species and Argonemertes ( Geonemertes ) aus- rudiment. The brain is formed by a uniform, traliensis as well as for the median dorsal nerve ring- shaped rudiment that later differentiates into of Drepanophorus spectabilis (Lebedinsky 1897 ; the four brain lobes and their connecting commis- Hickman 1963 ). Likewise, in Pilidiophora, inde- sural tracts. A temporal sequence of development pendent rudiments of the dorsal brain lobes from can only be seen by the preceding separation the cephalic disks and the ventral brain lobes of the ventral commissural tract from the sur- as well as the lateral medullary cords from the rounding ectodermal layer (Nusbaum and Oxner trunk rudiments have been reported in pilidia 1913). The lateral medullary cords are reported to by Bürger (1894 ). The relevance of these rather be outgrowths of the ventral brain lobes in most particular fi ndings, however, has been doubted species. Immunohistochemistry data reveal that by several other contemporary authors (Salensky the serotonergic neurons of the brain ring start to 1912 , 1914 ; Nusbaum and Oxner 1913 ). show immunoreactivity in the ventral lobes and Development of the proboscis nerves or the the ventral commissural tract fi rst, while pepti- nerves innervating the sense organs has never dergic (FMRF-like) immunoreactivity is fi rst been concisely described, but they have been observed in the dorsal brain lobes and commis- reported to form comparably late during 8 Nemertea 179 development in Prosorhochmus adriaticus (Senz labeling studies have revealed that the sequence and Tröstl 1999 ). of formation of muscle layers is uniform in all species studied. The inner longitudinal body wall Body Wall Musculature muscle layer is formed prior to the outer circu- The body wall muscle layers develop from mes- lar layer. Furthermore, in the Desor larva, the enchymal cells situated between the epidermis outer longitudinal muscles as well as the dermal and the gut rudiment. In both paleonemertean muscles develop later during post- larval develop- and hoplonemertean larvae, the sequence of ment. Moreover, these fi ndings argue against an formation of musculature is uniform, the inner ectodermal origin of the dermal and outer longi- longitudinal muscles being differentiated prior tudinal muscle layers, since muscle cells of both to the outer circular layer (Salensky 1914 ; Iwata the dermal and the longitudinal muscle layers 1960 ). Additional dorsoventral muscles and are observed prior to the formation of the dermis the musculature of the anterior head region are (von Döhren 2008 ). reported to develop much later in the hoplone- mertean species Prosorhochmus adriaticus (Senz Proboscis Apparatus and Tröstl 1999). There is, however, considerable Data on the development of the proboscis are only disagreement over the nature of the mesoderm. available for neonemertean species (Friedrich In Tetrastemma vermiculus and Drepanophorus 1979 ; Senz and Tröstl 1999 ; Maslakova 2010a , spectabilis, four longitudinal, mesodermal bands b ). In paleonemerteans the formation of the pro- originating from four mesendoblasts form all of boscis has never been witnessed during develop- the body wall and the splanchnic musculature ment. Therefore, the onset of the formation of the (Lebedinsky 1897). In pilidia, however, there proboscis has been assumed to occur quite late, are four somatic rudiments underneath the two close to or even after the shift to the benthic life- cephalic and the two trunk disks, but only one style. While in heteronemertean pilidiophorans splanchnic mesodermal rudiment surrounding the proboscis epithelium is formed from a cone- the larval gut formed from initially scattered shaped, proximal process of the fused cephalic mesenchymal cells (Salensky 1912 ). disks, it is generally reported to be formed as an Classical accounts on the formation of muscle epidermal invagination close to the apical organ layers in heteronemertean Pilidiophora differ in hoplonemerteans. both concerning the origin of muscle layers as According to classical accounts on pilidia, the well as regarding the sequence of their forma- cephalic disks fuse and the proboscis is either tion. Hubrecht (1885 , 1886 ) derived the muscle formed by an invagination of the proximal layer of layers in the Desor larva from mesoderm. The the head rudiment or by a separate proboscis disk fi rst muscle layer to form is the outer longitudinal that is incorporated into the head rudiment during layer, that is typical of Heteronemertea, followed fusion of the cephalic disks (Bürger 1894 ; Salensky by the inner circular and longitudinal layers. 1912 ; Schmidt 1937 ). A more recent account According to other authors the dermal muscles shows a different picture (Maslakova 2010a ). and the outer longitudinal layer are derived from While the cephalic disks fuse to form the head rudi- the proximal layer of the imaginal disks, while ment by proliferation of their proximal cell layer, the inner circular and longitudinal muscles are the putative proboscis rudiment is represented by a derived from the underlying mesenchymal cells separate cluster of cells underneath the larval epi- (Bürger 1894 ; Salensky 1912 ). A similar origin of dermis. The arguably mesodermal proboscis rudi- muscle layers has been reported in the Iwata larva ment makes its way ventrally to come to lie adjacent of Micrura akkeshiensis, although the sequence to the inner side of the proximal cell layer of the of formation of muscle layers is reversed: fi rst, head rudiment. A portion of the proximal layer of the inner longitudinal and circular layers are the head rudiment invaginates to form a cone- formed, while outer longitudinal and dermal shaped structure that protrudes into the larva so that muscles have not been seen until 39 days after the putative proboscis rudiment sits on it like a metamorphosis (Iwata 1958 ). Recent fl uorescent shallow cap, forming a compound proboscis bud. It 180 J. von Döhren has been reported that in the proboscis bud, the derived from an invagination of the secondary cone-shaped structure merely forms the ectodermal epidermis, and its formation corresponds to the components, i.e., the outer epithelium and the classical description on the development of the gland cells of the proboscis. Its muscular layers, the proboscis in the pilidium larva (Iwata 1958 ). inner epithelium, as well as the rhynchocoel epithe- In hoplonemerteans the proboscis is lium and the rhynchocoel muscles originate from formed from an epidermal invagination. In the cap-like putative proboscis rudiment by means Malacobdella grossa , Emplectonema gracile , of schizocoely. The rudiment of the proboscis Oerstedia dorsalis , Gononemertes australien- apparatus elongates on the dorsal side of the esoph- sis , Carcinonemertes epialti , and Paranemertes agus until it reaches the level of the larval gut. It peregrina , the proboscis rudiment detaches from consists of two components. An outer tube-shaped the epidermis, while in Drepanophorus spec- structure, the forming rhynchocoel wall, is clearly tabilis , Tetrastemma vermiculus , Prosorhochmus separated from an inner, invaginated structure rep- viviparus, and Pantinonemertes ( Geonemertes ) resenting the developing proboscis (Maslakova agricola, the connection to the epidermis per- 2010a). After elongation of the proboscis, the dor- sists (Lebedinsky 1897 ; Coe 1904 ; Salensky sal margin of the trunk rudiment extends anteriorly 1914 ; Hammarsten 1918 ; Iwata 1960 ; Egan and over the larval gut and the proboscis rudiment. Anderson 1979 ; Stricker and Reed 1981 ). In the The hypothetical homology of the nemertean former group of species (except Paranemertes rhynchocoel with secondary body cavities of peregrina ), the rhynchodeum is formed by an other coelomate spiralians is said to be supported independent invagination located more frontally, by the mode of its development (Maslakova in some species underneath the apical pit, while 2010b ). The hypothesis of a schizocoelous devel- in the latter group the rhynchodeum is differ- opment of the rhynchocoel, however, demands a entiated from the anterior part of the epidermal complete fusion of the ectodermal cone with the proboscis rudiment invagination. The tripartite mesodermal cap of the proboscis bud. The com- organization of the armed hoplonemertean pro- ponents of the proboscis rudiment, however, seem boscis becomes apparent by a strong stylet bulb to be separated in all stages investigated; schizo- with a narrow canal separating an anterior tube coelous processes within the mesodermal shaped from a posterior sacculate portion. In the component have not been reported (Maslakova area of the stylet bulb, the stylet armature is later 2010a). In the Desor larva, the proboscis epithe- formed (Bürger 1895 ; Coe 1904 ; Stricker and lium develops from an independent proboscis Reed 1981 ; Stricker and Cloney 1982 ; Stricker rudiment, while its musculature, the rhynchocoel 1985 ; Senz and Tröstl 1999). While the mesoder- and the rhynchocoel muscles, are derived from mal components of the proboscis apparatus are the mesoderm. It forms a continuous layer around derived from the surrounding mesodermal cells the proboscis to separate into a proximal layer in most accounts by delamination of a mass of forming the proboscis muscles and the inner pro- mesodermal cells, Lebedinsky (1897 ) identifi ed boscis epithelium and an outer layer developing a dorsal and a ventral mesodermal strip indepen- into the rhynchocoel epithelium and the associ- dent of the somatic mesoderm that form from ated muscle layers. In the part where no separa- paired mesoblast cells at the junction of the pro- tion of the layers occurs, the retractor muscle boscis invagination in Drepanophorus spectabi- develops (Arnold 1898 ; Nusbaum and Oxner lis and Tetrastemma vermiculus . The rudiments 1913 ). Whether this separation is by delamination become hollow, associate with the proboscis or schizocoely is not reported. Fluorescent label- invagination, grow around it, and fuse. The inner ing of F-actin reveals that the formation of the wall of the fused mesodermal rudiment forms the proboscis musculature precedes that of the rhyn- proboscis musculature and its inner epithelium, chocoel in the Desor larva of Lineus viridis (von the outer rhynchocoel wall, and the associated Döhren J 2015, unpublished). In the Iwata larva of muscle layers. The hollow space between the Micrura akkeshiensis , the proboscis rudiment is two-layered rudiment represents the rhynchocoel. 8 Nemertea 181

Alimentary Canal the esophagus develops early and retains its func- The intestinal tract in nemerteans is a one-way tionality, the esophagus in the latter is reported to through-gut. It comprises the midgut and its be separated from the midgut. A defi nite connec- derivatives as well as a histologically different tion is formed late, i.e., after metamorphosis, by a foregut. An extensive hindgut connecting the secondary esophagus from two groups of cells midgut with the anus is absent (Bürger 1895 ; located laterally near the mouth opening (Arnold Gibson 1972). The midgut and its derivatives 1898 ; Nusbaum and Oxner 1913 ; Schmidt 1964 ). develop from the embryonic endoderm. In species The development of the mouth opening and with gastrulation by invagination, the gut persists pharynx in hoplonemerteans is much more diverse in the larva as a hollow cavity, while in species and complicated. In general, the blastopore is said that gastrulate by polar ingression, the midgut to be closed early in development. In some species rudiment is regularly a solid mass of cells that (e.g., Emplectonema gracile , Oerstedia dorsalis ), establish the gastric cavity later during develop- the blastopore reopens and a functioning larval gut ment (Friedrich 1979). In paleonemerteans and is formed (Iwata 1960 ). In other species the larval hoplonemerteans, the future mouth is moved to gut is formed by a secondary invagination the future ventral side by accelerated growth of of the epidermis, forming the esophagus in a the dorsal side of the body, while in canonic pilid- more anterior position than the blastopore iophoran development, it remains in its original (Drepanophorus spectabilis , Tetrastemma vermic- position (Iwata 1957 , 1960 , 1985 ; Friedrich 1979 ; ulus , Malacobdella grossa , Paranemertes pereg- Maslakova 2010a , b). During and following gas- rina ) (Lebedinsky 1897 ; Hammarsten 1918 ; trulation, parts of the ectoderm have regularly Maslakova and von Döhren 2009 ). In Tetrastemma been reported to be dragged inside the gastrula vermiculus and Drepanophorus spectabilis , the with the blastopore to form the ectodermally blastopore remains open even after the secondary derived esophagus (Friedrich 1979 ). In paleone- esophagus has formed. It has been reported to close mertean species and Pilidiophora that develop via later so that the remaining pouch develops into the a pilidium, the blastopore remains open, marking intestinal cecum present in most hoplonemerteans the connection between the esophagus and the (Lebedinsky 1897 ). While the mouth and probos- blindly ending, sac-like midgut. All or most of the cis pore are separate in most polystiliferan hop- larval intestinal tract is taken over from the larval lonemerteans, in monostiliferan hoplonemerteans to the juvenile organization. According to the larval mouth subsequently closes, and the Salensky (1912 ), the esophagus of the pilidium is esophagus gains connection with the rhynchodeum divided into a distal and a proximal part by a con- forming the rhynchostomodeum that is typical of strictor muscle in later-stage larvae. During meta- this clade. In Prosorhochmus viviparus , Prostoma morphosis only the proximal part is taken over graecence , Argonemertes (Geonemertes ) aus- into the juvenile; the distal part is shed along with traliensis, and Gononemertes australiensis , neither the pilidial envelope. In the Iwata larva of Micrura a reopening nor a secondary esophagus forms, akkeshiensis , the blastopore remains open resulting in a midgut that is closed for most of the although no food is taken up. The defi nite mouth time of the development (Salensky 1909 , 1914 ; opening and esophagus develop from a rudiment Hickman 1963; Egan and Anderson 1979 ). A func- composed of multilayered cells located in the dor- tioning intestinal opening is accomplished by sal wall of the stomodeum. By elongating in a fusion of the midgut with the ventral part of the slight curve, the rudiment becomes the esophagus rhynchodeum. of the juvenile. The stomodeal part distal of the The fusion of the esophagus with the rhyncho- esophageal rudiment is shed along with the larval deum is accomplished in various ways. While in epidermis during metamorphosis (Iwata 1958 ). Emplectonema gracile , Oerstedia dorsalis , and During intracapsular development of Lineus Gononemertes australiensis the esophagus and the ruber and Lineus viridis, the esophagus develop- proboscis rudiments gain access to the indepen- ment differs markedly. While in the former species dently invaginated rhynchodeum, the esophagus 182 J. von Döhren fuses to the distal part of the proboscis rudiment that as simple protonephridia anterior of the mouth later becomes the rhynchodeum in Prosorhochmus opening (Bartolomaeus et al. 2014 ). They consist viviparus (Salensky 1909 , 1914 ; Iwata 1960 ; Egan of few (two to three) multiciliated terminal cells and Anderson 1979 ). In Tetrastemma vermiculus , constituting the site of ultrafi ltration and two to Drepanophorus spectabilis, and Pantinonemertes three multiciliated cells that form the nephrod- (Geonemertes ) agricola , the anterior-most part of uct which modifi es the ultrafi ltrate. Distally, the the proboscis rudiment forms a ventral invagina- nephroduct opens through the epidermis at the tion that fuses with the intestine developing into level of the mouth opening via a nephropore the adult esophagus (Lebedinsky 1897 ; Coe 1904 ). cell in both species (Bartolomaeus et al. 2014 ). In Paranemertes peregrina, the proboscis rudiment The protonephridia develop from a subepider- gains its connection prior to the closure of the lar- mal rudiment underneath the trochoblast cells val mouth opening (Maslakova and von Döhren and are fully formed prior to the degeneration of 2009). In the aberrant commensal Malacobdella the so- called vestigial prototroch (Bartolomaeus grossa , there is no true rhynchodeum. The probos- et al. 2014). In hoplonemertean species proto- cis rudiment connects to the esophagus and the lar- nephridia have been observed in larvae but in a val mouth is closed. A secondary mouth opening is more posterior position behind the mouth open- formed anterior of the larval mouth (Hammarsten ing. Nothing is known about the ultrastructure 1918 ). Apart from the disparate development in of the protonephridia in hoplonemertean larvae. Malacobdella grossa, there is no conceivable rea- Structure, position, and time of development son for the diversity shown in esophagus devel- of the protonephridia in paleonemerteans are opment within the otherwise relatively uniform reminiscent of the “head kidneys” of the trocho- clade of Hoplonemertea. It is therefore very likely phore larva, although it is not clear whether the that the diversity shown in the development of the nemertean protonephridia share the same fate as esophagus and rhynchostomodeum depends rather transitory organs that are restricted to the larval on the different researchers than on diverging lines organization. of development (Friedrich 1979 ). The protonephridia of hoplonemertean larvae In some hoplonemerteans, an anus is formed correspond in position to the respective organs in early in development by means of a caudal epi- the adults. Therefore, it is very probable that the dermal invagination or ingression (Lebedinsky excretory organs in hoplonemerteans represent 1897; Hammarsten 1918; Iwata 1960 ; Friedrich early developmental stages of the adult organs 1979; Maslakova 2010b). Apparently, in the being elaborated during development to attain majority of hoplonemerteans, as well as in the adult morphology. In Pilidiophora, branched Pilidiophora and paleonemertean species studied, protonephridia located slightly anterior of the the anus is formed much later as the formation of midgut have been observed as early as 2 weeks an anal opening has not been reported in these. after fertilization (von Dassow and Maslakova Initially, the intestinal tract is an undifferentiated 2013). In later stages the protonephridia become tube. The morphological differentiation of the sandwiched between the intestinal tract and the alimentary canal in hoplonemerteans comprising distally developing juvenile rudiment (Maslakova the ectodermal esophagus, stomach, and pyloric 2010a). In Lineus viridis the fi rst protonephridia tube as well as the mesodermal intestinal ceca were observed in the juvenile after metamorpho- and lateral diverticula forms comparatively late sis. At this time no trace of the blood-vascular (Friedrich 1979 ; Senz and Tröstl 1999 ). system can be observed. The protonephridia are branched structures with a single terminal cell Nephridia on the proximal end of each branch. Initially Excretory organs have been observed to develop monociliated, the terminal cell becomes soon early in paleonemertean larvae of Carinoma multiciliated. The nephroduct is intercellular, mutabilis and Procephalothrix oestrymnicus and the nephropore is formed by four specialized 8 Nemertea 183 epidermal cells (Bartolomaeus 1985 ). Since the gland cells, and neurons located behind the fi rst protonephridia formed correspond in struc- brain in Pilidiophora and reptant Polystilifera. ture and position to those seen in the adults, it can In the heteronemertean pilidium and the Desor be assumed that protonephridia in Pilidiophora larva, the cerebral organs develop from the cere- are defi nite persisting adult organs (Bartolomaeus bral organ disks, a paired rudiment invaginated et al. 2014). Although it has been shown that con- from the subumbrellar larval ectoderm. They are trary to classical accounts the protonephridia in situated bilaterally between the frontal cephalic the pilidium are not derived from bilateral invagi- disks and the posterior trunk disks and are the nations of the esophagus, their origin as invagi- last of the invaginated rudiments to form in the nations of the subumbrellar epidermis could not respective larvae. The cerebral organ disks fi rst be substantiated either (Hubrecht 1885 , 1886 ; fuse posteriorly with the trunk disks and subse- Arnold 1898 ; Salensky 1912; Maslakova 2010a ). quently with the already fused head rudiment. In the Iwata larva of Micrura akkeshiensis , the The invagination of the original disks is retained origin of the protonephridia has been identifi ed to later form the cerebral organ canal, while as a pair of cell groups located in the ventral wall gland cells and neurons form in the proximal por- of the stomodeum on the level of the juvenile tion of the rudiment. The neurons connect to the mouth opening (Iwata 1958 ). Their further devel- developing dorsal brain lobes already before the opment, however, has not been followed. While juvenile body has completely formed (Salensky the abovementioned data hint at an ectodermal 1912 ; Nusbaum and Oxner 1913; Maslakova origin of the protonephridia, a mesodermal origin 2010a ; Hindinger et al. 2013 ). Although nothing has been hypothesized for the protonephridia in is known about the formation of cerebral organs the Desor larva. The rudiment is represented by a in non-heteronemertean Pilidiophora such as cluster of cells which later forms narrow tubular Hubrechtella species, it can be assumed that the structures situated between the esophagus, the process is similar as described in the heterone- cerebral organs, and the ventral rhynchocoel wall mertean pilidium judging from the correspond- (Nusbaum and Oxner 1913 ). Further develop- ing larval type and the morphological similarity ment and opening of the nephridial rudiments to of the cerebral organs in the respective taxa. the exterior have not been observed. In the Iwata larva, the cerebral organs are not derived from paired epidermal larval invagina- Sensory Organs tions but from paired lateral invaginations of the Sensory organs comprise cerebral sense organs stomodeum. The connection to the stomodeum present in many paleonemertean, nearly all is soon obliterated, and the cerebral organs open pilidiophoran and most hoplonemertean spe- secondarily into the cephalic furrow formed in cies; frontal organs and eyes, both of which are the cephalic rudiment. A connection to the dorsal present in the majority of Neonemertea; and brain lobes is accomplished prior to metamor- lateral organs that are confi ned to some tubu- phosis (Iwata 1958 ). Cerebral organs in larvae lanid species (Gibson 1972). Cerebral organs of hoplonemertean species have been reported to differ in their complexity and position relative develop from a pair of narrow invaginations of to the brain in different taxa. Cerebral organs thickened epidermal areas that can be observed comprise a ciliated, blind ending canal, running early in development during the larval phase from the epidermis proximally to end in a mass (Lebedinsky 1897; Iwata 1960; Maslakova and of neuronal tissue that is connected via a nerve von Döhren 2009 ). In Paranemertes peregrina to the brain. While they are comparably small, and Emplectonema gracile , they are situated simple canals situated in front of the brain in bilaterally on both sides of the larva on the level Monostilifera or behind the brain in Tubulanus of the mouth opening, while in Tetrastemma and Carinina species, cerebral organs are large, vermiculus and Drepanophorus spectabilis, the compound organs comprising ciliated cells, epidermal invaginations have been reported to be 184 J. von Döhren located between the dorsal and the ventral brain 2004b ; Chernyshev 2008 ; Maslakova and von lobes (Lebedinsky 1897; Iwata 1960; Maslakova Döhren 2009 ). But while the eyes in the larva are and von Döhren 2009). Epidermal invaginations transitory in most paleonemertean species, they that have initially been interpreted as cerebral persist and become more in number in organ rudiments in some hoplonemertean spe- Hoplonemertea (Fig. 8.4C ). In Heteronemertea cies have later been interpreted as invaginations pigmented eyes are confi ned to the juvenile rudi- that were hypothesized to be homologous to ment; no larval type of Pilidiophora is reported the imaginal disks of Pilidiophora (Maslakova to have eyes (Fig. 8.6D ; Cantell 1969 ; Norenburg 2010b ). In the direct developing hoplonemer- and Stricker 2002; von Döhren and Bartolomaeus tean Prosorhochmus adriaticus , the internal ner- 2007 ). The eyes in neonemertean species are vous and glandular portion of the cerebral organ subepidermal, rhabdomeric eyes (Jespersen and forms fi rst, while its connection to the exterior Lützen 1988; von Döhren and Bartolomaeus via the cerebral organ canal and pore opening in 2007 ; von Döhren 2008). In Lineus viridis the the cephalic furrows is established later in devel- eyes develop from an unpigmented, subepider- opment (Senz and Tröstl 1999 ). In Malacobdella mal rudiment comprising a small number of grossa larval cerebral organ rudiments have been cells of two types. A bundle of rhabdomeric suspected although the adult does not possess receptor cells is surrounded by undifferentiated, cerebral organs. The cerebral organ rudiments unpigmented corneal progenitor cells. From the comprise paired epidermal cell clusters that are latter type of cells, the closed optical cavity is situated a little posterior of the mouth open- formed that houses the receptor cells. Pigmented ing. After being invaginated they give off some cells that form a pigment cup to one side are neuronal cells to contribute to the ventral brain formed as the eye begins to function. The eye lobes. The remainder of the rudiment disappears spot enlarges as the number of all cell types shortly after (Hammarsten 1918). Although cere- increases (von Döhren and Bartolomaeus 2007 ). bral organs exist in some paleonemertean spe- Contrary to the statement that the eyes in cies, neither their formation nor any rudiments Heteronemertea form by fragmentation of exist- have ever been observed (Iwata 1960). While ing eyes, additional eyes in Lineus viridis juve- this appears logical in species lacking cerebral niles are formed de novo in the same manner as organs, it is somewhat astonishing judging from described above (Gontcharoff 1960 ; von Döhren the fact that cerebral organ rudiments are formed and Bartolomaeus 2007 ). early in other nemertean species. Moreover, The frontal sensory organ is typical of recent phylogenetic analyses suggest cerebral Neonemertea being represented by a single organs to be an ancestral character in Nemertea protrusible epidermal pit in most species, while that was reduced repeatedly (Thollesson and comprising three triangularly arranged pro- Norenburg 2003; Andrade et al. 2012 ). It would trusible pits in lineid heteronemertean species therefore be conceivable to interpret the bilateral (Gibson 1972 ). The frontal organ is commonly apical invaginations that had been attributed as associated with the head glands that discharge rudiment of the brain in paleonemertean species their glandular products through canals open- as a joint rudiment giving rise to not only the ing between the epithelial cells of the frontal brain and lateral medullary cords but also to the organ. In some species the head glands open cerebral organs. In species that do not possess independently of the frontal organ by numer- cerebral organs as adults, e.g., Procephalothrix ous smaller ducts to the epidermis (Gibson simulus , they are reduced later in development. 1972 ). According to Lebedinsky ( 1897 ), the In Hoplonemertea and some paleonemertean frontal organ develops from the apical pit of species, the eyes are formed during the larval the larva in Drepanophorus spectabilis and phase (Fig. 8.6A–C; Iwata 1960 ; Stricker and Tetrastemma vermiculus, although this has been Reed 1981 ; Martindale and Henry 1995 ; doubted by other authors (Bürger 1897 –1907; Norenburg and Stricker 2002 ; Maslakova et al. Hammarsten 1918). In Malacobdella grossa the 8 Nemertea 185 apical pit gives rise only to the cephalic glands by resorbing encased yolk material (Reinhardt (Hammarsten 1918 ). 1941). Ultrastructural data on Prosorhochmus Various sensory structures are restricted to americanus , however, provide a different picture certain lineages, such as tactile cirri and stato- (Turbeville 1986 ). The solid bands of the blood cysts in Ototyphlonemertes species; lateral sen- vessel rudiments become hollow by a process sory organs in some Tubulanus , Callinera , and that is reminiscent of the schizocoelous mode of Micrella species; or so-called integumentary and formation observed in annelid coelomic cavities. subcutaneous organs of pelagic Polystilifera (Coe In contrast to annelid schizocoely, the cells of 1927 ; Gibson 1972 , 1982 ). There are no data the nemertean blood vessels acquire intercellular available about the development of these organs. junctions after the onset of cavitation, whereas the respective intercellular junctions in annelids Blood-Vascular System are formed at the onset of cavitation (Turbeville The development of the blood-vascular system 1986 ; Koch et al. 2014). Due to these differ- occurs comparably late in Nemertea after the ences, the question of homology of these laterally majority of musculature has already formed. located endothelialized, hollow compartments to While, according to classical accounts, the blood the secondary body cavities found in Trochozoa vessels in Pilidiophora precede metamorpho- can be at best preliminarily answered; more com- sis, no trace of blood vessels could be found in parative data on the development of the blood the postmetamorphic juvenile of Lineus viridis vessels in other nemertean taxa are needed to according to more recent data (Bürger 1897 – consolidate this hypothesis. 1907; Nusbaum and Oxner 1913 ; Bartolomaeus In summary, the development of Nemertea is 1985). There is no account on formation of the generally working on the regular spiralian scaf- blood vessels in paleonemertean species, while fold; although in organs that represent interfaces in Hoplonemertea data are only available for spe- of two germ layers (e.g., proboscis, foregut, cies with intracapsular development (Salensky nephridia), there is considerable disagreement 1914 ; Reinhardt 1941; Turbeville 1986). In clas- over their respective origin and formation. Until sical accounts, the blood vessels have either been more comparative data are collected, concise derived from gaps remaining or been reopened statements regarding the evolution of organ sys- in the blastocoel or from postulated embryonic tem development in Nemertea cannot be made. coelems (e.g., Bürger 1897 –1907; Salensky 1914 ; Nusbaum and Oxner 1913 ). The problem in these classical accounts is that the blastocoel GENE EXPRESSION was considered to be devoid of matrix or that the matrix within it was later liquefi ed. This led to Gene expression studies in development of the misinterpretation that nemerteans possess Nemertea are presently scarce and restricted to the extensive primary and/or secondary body cavi- pilidiophoran species Cerebratulus lacteus , Lineus ties. Instead, nemerteans should be considered as viridis , Micrura alaskensis , and Ramphogordius largely compact with the exception of the rhyn- (Lineus ) sanguineus . The data available refer to chocoel and the blood vessels that represent sec- the expression of β -catenin, a trochoblast-specifi c ondary body cavities (Gibson 1972 ; Turbeville tubulin-4 gene from the basal gastropod Patella and Ruppert 1985 ; Turbeville 1991 ). More recent vulgata, homeobox-containing genes (Hox genes, data on two hoplonemertean species suggest that as well as Otx, and Cdx), genes encoding phot- the blood vessels are formed in the parenchyma- opigments ( opsins), and genes of the so-called tous tissue underlying the body wall muscula- retinal determination gene network (RDGN) ture by forming solid bands of cells (Reinhardt ( Pax-6, Six1/2, Six3/6, Six4/5, Dach) (Loosli et al. 1941 ; Turbeville 1986). According to light 1996 ; van den Biggelaar et al. 1997 ; Klerkx 2001 ; microscopic data in Prostoma graecense , the Charpignon 2007 ; Henry et al. 2008 ; Döring 2012 ; bands of radially arranged cells become hollow Hiebert and Maslakova 2015 ). 186 J. von Döhren

Axis Specifi cation Genes Wnt pathway, is both necessary and suffi cient to promote endoderm fates in the vegetal cells A transcriptomic assessment of the pilid- during cleavage of Cerebratulus lacteus (Henry iophoran Micrura alaskensis revealed a single et al. 2008 ). β -catenin is expressed only in the Hox cluster comprising nine Hox genes while vegetal-most quartet at each cleavage division only six to seven Hox genes have been identi- and is passed on through the cleavage cycles to fi ed by genomic screening of the pilidiophoran fi nally end up in the blastomeres forming the Ramphogordius (Lineus) sanguineus (Kmita- endoderm. Experimental overexpression leads Cunisse et al. 1998; Hiebert and Maslakova to vegetalized embryos that fail to form typical, 2015 ). In Micrura alaskensis (Ramphogordius anteriorly positioned apical tufts, while block- (Lineus) sanguineus), the genes of the Hox ing of β-catenin leads to animalized larvae with gene cluster have been identifi ed as MaLab supernumerary apical tufts. Animalized larvae (LsHox1), MaPb, MaHox3 (LsHox3), MaDfd gastrulate only to a limited extent. It has been (LsHox4), MaScr, MaLox5 (LsHox6), MaAntp hypothesized that the animal fates of blastomeres (LsHox7), MaLox4, and MaPost2 (LsHox9). In represent the default condition which is shifted Ramphogordius (Lineus) sanguineus a seventh to vegetal fates under the infl uence of β -catenin putative Hox gene, LsHox8 that is possibly dis- (Henry et al. 2008 ). A trochoblast-specifi c tubu- located from the chromosomal Hox gene region lin gene (tub4 ) was assessed for its expression was found using an alternative set of primers in the cells that form the marginal band of elon- (Kmita-Cunisse et al. 1998). Data on the expres- gated cilia on the lappets around the pilidium of sion of Hox genes during development are only Cerebratulus lacteus (van den Biggelaar et al. available for Micrura alaskensis (Hiebert and 1997 ; Klerkx 2001 ). However, no expression Maslakova 2015 ). Hox gene expression largely signal was recorded in cells bearing elongated complies both temporally and spatially to the cilia, underpinning the hypothesis that the mar- canonical bilaterian fashion, albeit with note- ginal band of cells bearing elongated cilia is worthy exceptions: All pilidial tissues are com- not homologous to the prototroch of the trocho- pletely devoid of Hox genes expression during phore (van den Biggelaar et al. 1997 ; Maslakova all stages of development. Hox gene expression 2010a ). An Otx-class and a Cdx-class gene were in Micrura alaskensis does not start before the identifi ed in Ramphogordius (Lineus ) sanguin- trunk disks are formed. It then proceeds posteri- eus (Ls-Otx , Ls-Cdx ) (Kmita-Cunisse et al. 1998; orly. Thus the anterior part of the juvenile com- Charpignon 2007). A Cdx ortholog ( MaCdx ) has prising the cephalic disks and the cerebral organ also been found in Micrura alaskensis (Hiebert disks as well as the later developing head rudi- and Maslakova 2015 ). Ls-Otx was found to be ment and the proboscis develop without showing expressed during the development of postmeta- Hox gene expression. The absence of Hox gene morphic juvenile Lineus viridis (Fig. 8.9A ; expression in the pilidial tissues has been inter- Charpignon 2007 ). First, it is expressed in the preted as an indication of a functional decou- entire brain ring and the anterior-most part of the pling of the early, larval from the later juvenile lateral medullary cords, the cerebral organs and development phases (Hiebert and Maslakova their canals, and additionally, but weaker, in the 2015 ). While giving a possible explanation of gut. In older stages there is an Ls-Otx expres- the diversity of pilidial morphotypes the genetic sion in the frontal organ and in the frontal organ mechanism for the patterning of the pilidial nerves. Expression in the brain and cerebral envelope remains unclear. Preliminary results of organs becomes weaker, although in the cerebral inhibition studies hint at an involvement of the organ canals, Ls-Otx is still clearly expressed. At Wnt and the fi broblast growth factor pathways this stage of development, no expression signal in patterning the early larval shape (Hiebert is detectable in the lateral medullary cords or and Maslakova 2015). Recent fi ndings indicate the gut (Charpignon 2007 ). Ls-Cdx is strongly that β -catenin , a downstream component of the expressed in Lineus viridis juveniles consistently 8 Nemertea 187

Fig. 8.9 Schematic representation of gene expression of Lineus viridis. br brain ring, co cerebral organ, coc cerebral several genes from Ramphogordius (Lineus ) sanguineus in organ canal, de developing eye, lmc lateral medullary cord, postmetamorphic juveniles of Lineus viridis. (A ) Expression mg midgut, mo mouth opening, pb proboscis rudiment of Ls-Otx and Ls-Cdx . ( B ) Expression of Ls-opsin and LsPax- (© Dr. J. von Döhren, All Rights Reserved) 6 . Note: LsPax-6 is not expressed in the developing eyes of

throughout development in the posterior-most et al. 1996; Charpignon 2007; Döring 2012). One part of the body. The signal is located in inter- opsin gene has been isolated from Ramphogordius nal tissues, arguably in the developing intestine (Lineus ) sanguineus , while two opsin genes have (Fig. 8.9A; Charpignon 2007). In Micrura been found in Lineus viridis (Charpignon 2007 ; alaskensis MaCdx is fi rst expressed in the trunk Döring 2012 ). The former, Ls-opsin, represents disk being shifted posteriorly during the course an unusual G-protein- coupled-receptor (GPCR) of development to be fi nally located as a ring opsin that does not cluster with canonical rhabdo- shaped pattern around the base of the developing meric-type opsins. Its expression in Lineus viridis caudal cirrus. In this species however, expres- is restricted to the developing eyes, while expres- sion of MaCdx in the intestinal tissue is absent sion in adult eyes is absent (Fig. 8.9B ; Charpignon (Hiebert and Maskakova 2015 ). 2007 ). The opsins identifi ed in Lineus viridis (LiVi-ops1 , LiVi-ops2 ) cluster with peropsins and photoisomerases (LiVi-ops1 ) or in a basal position Retinal Determination Gene to these together with opsin5 ( LiVi-ops2) (Döring Network Genes 2012 ). Although a G-protein alpha subunit Q gene (LiVi-Gq ) was also found in the same species, The most extensive expression data have been gath- expression experiments for any of the three genes ered on the eye development in Lineus viridis (Loosli were unsuccessful (Döring 2012 ). Genes marking 188 J. von Döhren the canonical neurotransmitters employed by pho- ing brain and the cerebral organs but also in two toreceptor cells of Platynereis dumerilii , the vesicu- bilateral stripes running from the brain lobes lar acetylcholine receptor gene in larval eyes, and anteriorly. A correspondence with the nerves the vesicular glutamate receptor gene in adult eyes connecting the eyes with the brain seems likely have both been extracted from juveniles of Lineus (Charpignon 2007 ). viridis ( LiVi-vacht and LiVi-vglut , respectively). Expression data on these genes in Lineus viridis have not been reported (Döring 2012 ). Other Genes Expression of genes of the RDGN during development was studied in Lineus viridis and Genes that have been identifi ed in Ramphogordius includes the genes LsPax-6 and Lv-Six1/2 , (Lineus ) sanguineus but have not been sub- Lv-Six3/6 , Lv-Six4/5 , and Dach , although for the jected to expression studies include Ls-Bmp2/4 , latter an antibody reaction was studied instead of Ls-Engrailed , Ls-Msx , LsNK , LsPax-2/5/8 , in situ hybridization (Loosli et al. 1996 ; Ls-Snail, and Ls-Twist (Charpignon 2007 ). Gene Charpignon 2007 ). In Micrura alaskensis a expression of the canonical developmental path- Six3/6 ortholog ( MaSix3/6) was found to be ways is very limited both with respect to the gene expressed during early larval development products involved and to the diversity of develop- (Hiebert and Maslakova 2015 ). LsPax-6 is mental trajectories found in Nemertea. Currently, a expressed in Lineus viridis during postmetamor- sound assessment of gene expression networks in phic development in the brain and the cerebral Nemertea is impossible. organs, but not in the eye region (Fig. 8.9B ; Loosli et al. 1996 ; Charpignon 2007 ). Due to dif- ferential splicing and a thus changed open read- OPEN QUESTIONS ing frame, two isoforms have been hypothesized to exist. One of them is shorter, missing the PST • How does the high yolk content in Pelagica domain. The possible role of this shorter isoform oocytes infl uence embryonic cleavage? has been suspected to be differential regulation • How are the dorsal quadrant and the meso- of developmental processes. However, there is derm precursor cell specifi ed in Paleonemertea no proof of the existence of the theoretical iso- and Hoplonemertea? form (Charpignon 2007 ). Of the remaining genes • Is there an apical neuronal structure that is assessed, only Lv-Six1/2 and the monoclonal homologous to the apical organ of Trochozoa Drosophila melanogaster antibody against Dach in larvae of Nemertea? show an expression of these genes in the devel- • How did the larval and adult eyes in Nemertea oping eyes of Lineus viridis . The former gene is evolve? also expressed in the lateral medullary cords • What is the developmental origin of nemer- near the brain and in the frontal organs. Lv-Six3/6 tean protonephridia? also shows expression in the frontal organ, but • What is the developmental fate of the fi rst its strongest expression is observed in the brain formed protonephridia – transitory larval or lobes. Expression of MaSix3/6 is restricted to defi nite adult organs? early developmental stages of Micrura alaskensis . • Where does the adult mesoderm originate In the blastosquare stage several cells on one from – entirely endomesodermal or with ecto- pole of the embryo are labelled while in the feed- dermal components? ing pilidium stage some expression signals are • Is there a taxon-specifi c mode of muscle for- detectable near the apical pit along with few mation in Nemertea? additional signals situated in the anterior region • How do the different components of the pro- of the developing lateral lappets (Hiebert and boscis and rhynchocoel develop? Maslakova 2015). In Lineus viridis Lv-Six4/5 is • Is there a generalizable mode of foregut for- expressed in the posterior region of the develop- mation in Hoplonemertea? 8 Nemertea 189

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Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger.

C. Bleidorn (*) • C. Helm • A. Weigert Molecular Evolution and Systematics of Animals , Institute of Biology, University of Leipzig , Talstraße 33 , Leipzig D-04103 , Germany e-mail: [email protected] M. T. Aguado Departamento de Biología, Facultad de Ciencias , Universidad Autónoma de Madrid , Canto Blanco , Madrid 28049 , Spain

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 193 DOI 10.1007/978-3-7091-1871-9_9, © Springer-Verlag Wien 2015 194 C. Bleidorn et al.

INTRODUCTION sister group relationship (Edgecombe et al. 2011 ). Annelids are a taxon of protostomes compris- Annelids show a huge diversity of body plans, ing more than 17,000 worldwide-distributed and it is diffi cult to describe a consistent anatomy species, which can be found in marine, limnic, matching most of this variety (Fig. 9.2 ). Most and terrestrial habitats (Zhang 2011). Their annelids are coelomate organisms, possessing phylogeny was under discussion for a long multiple segments which occur repetitively along time, but recent phylogenomic analyses the anterior-posterior body axis (Purschke 2002 ). resulted in a solid backbone of this group If segmentation is present, the annelid body is (Struck et al. 2011 ; Weigert et al. 2014 ). divided into a prostomium, an either homono- According to these analyses, most of the anne- mously (i.e., identical segments) or heterono- lid diversity is part of Errantia or Sedentaria, mously (i.e., segments differ from each other) which both form reciprocally monophyletic segmented trunk and a pygidium (Fauchald and sister groups (Fig. 9.1 ) and are now known as Rouse 1997 ). In many annelid taxa, the prosto- Pleistoannelida (Struck 2011 ). The Sedentaria mium contains the brain; however, in Clitellata also include the Clitellata, Echiura, and the brain may be found in the following segments Pogonophora (Siboglinidae) as derived annelid (Bullock 1965 ). The head of the annelids may taxa. Outside Sedentaria and Errantia, several bear appendages, as palps or antennae, but these groups can be found in the basal part of the are lacking in a number of taxa. The mouth can annelid tree, namely, Sipuncula, Amphinomida, be found in the fi rst segment which is termed Chaetopteridae, Magelonidae, and Oweniidae. peristomium. Several members of the Errantia as The latter two taxa together represent the sister well as Amphinomida bear sclerotized mandibu- taxon of all other annelids. Given this hypoth- lar structures, which may be replaced by a mech- esis, it has to be assumed that the early diversi- anism resembling molting (Paxton 2005 ). The fi cation of extant annelids took place at least in segments of many annelids contain a pair of the Lower Cambrian (520 Ma ago) (Weigert nephridia (usually metanephridia), coelomic cav- et al. 2014 ). The phylogenetic position of ities, ganglia, and ventral and dorsal groups of Myzostomida, a group of commensals or para- chitinous chaetae which might be organized in sites of echinoderms (and, rarely, cnidarians), parapodia (Purschke 2002 ; Bartolomaeus et al. remains still uncertain. Whereas there is strong 2005). Segments are generated by a posterior support for an annelid ancestry, its exact posi- growth zone which is located in front of the tion awaits to be determined (Bleidorn et al. pygidium (Nielsen 2004 ). The pygidium contains 2014 ). Likewise, the phylogenetic position of the anus, which is usually either dorsally or ter- several interstitial taxa is still under debate minally located and is often equipped with pairs (Westheide 1987 ; Worsaae and Kristensen of cirri. 2005; Worsaae et al. 2005; Struck 2006). A Annelids show a wide variety in the position of Diurodrilidae outside Annelida, as organization of their nervous system (Bullock suggested by Worsaae and Rouse (2008 ), was 1965 ; Orrhage and Müller 2005 ; Müller 2006 ). rejected by molecular data (Golombek et al. Müller (2006 ) proposed a nervous system with 2013), and the position of the enigmatic paired circumesophageal connectives, four cere- Lobatocerebrum and Jennaria remains unre- bral commissures, fi ve connectives, and numer- solved (Rieger 1980, 1991 ). Likewise, the ous commissures in the ventral nerve cord as a position of Annelida within Protostomia is still hypothetical ground pattern. However, many uncertain. However, recent phylogenomic variations of this pattern exist, and many taxa analyses recover a clade uniting annelids have not been investigated at all. Accordingly, with Mollusca, Nemertea, Brachiopoda, and alternative hypotheses suggest that a strict Phoronida, but without strong support for any rope-ladder-like nervous system with segmental 9 Annelida 195

Fig. 9.1 Phylogeny of Annelida based on Weigert et al. (2014 ). Placement of well-investigated model annelids indi- cated with asterisks

ganglia interconnected by a pair of connectives tional longitudinal nerves might constitute the and commissures was not present in the last com- ground pattern of Annelida (Lehmacher et al. mon ancestor of annelids (Purschke et al. 2014 ). 2014 ). Instead, an orthogonal arrangement of the periph- Annelids show varying grades of brain com- eral nervous system and the presence of addi- plexity which may comprise a number of ganglia. 196 C. Bleidorn et al.

A BC

DEFG

HI J

KLM

NOP

Fig. 9.2 The diversity of marine Annelida. ( A ) Brada Phyllodocidae. (K ) Polynoidae indet. (L ) Sabellidae villosa , Flabelligeridae. (B ) Glycera capitata , Glyceridae. indet., Sabellidae. ( M ) Spirobranchus giganteus , ( C ) Lepidonotus squamatus, Polynoidae. (D ) Nereis pela- Serpulidae. ( N ) Lysidice sp., Eunicidae. ( O ) Serpula sp., gia, Nereididae. (E ) Chaetopterus sp., Chaetopteridae. Serpulidae. ( P ) Travisia sp., Travisia (All images pro- ( F) Syllinae indet., Syllidae. ( G ) Branchiomma arctica , vided by Alexander Semenov ( www.clione.ru ). © Sabellidae. ( H ) Lumbrineris sp., Lumbrineridae. (I ) Alexander Semenov, 2015. All Rights Reserved) Amblyosyllis sp., Syllidae. (J ) Phyllodoce sp., 9 Annelida 197

Mushroom bodies have been reported for several Many annelids possess some kind of light taxa of the Phyllodocida (Heuer et al. 2010 ). As receptive photoreceptors which show great for the nervous system, many different variations structural diversity (Purschke et al. 2006). can be found in the muscular system. The pres- Generally, rhabdomeric, ciliary, and phaoso- ence of an outer layer of circular muscle fi bers mous photoreceptor cell types are distin- and an inner layer of four longitudinal bands of guished, and they might represent either larval muscle fi bers is often regarded as a possible or adult eyes (Purschke et al. 2006; Arendt ground pattern (Tzetlin and Filippova 2005 ; et al. 2009). Larval eyes are simple organized, Lehmacher et al. 2014 ), but circular muscles are and the eye spots of the trochophore of missing in several annelid taxa and may thus not Platynereis dumerilii consist of a rhabdomeric constitute a basal annelid feature. Additionally, photoreceptor cell and a pigment cell which other muscular fi ber bundles referred to as provide a direct coupling of light-sensing cili- oblique, diagonal, bracing, or dorso-ventral fi bers ary locomotory control (Jekely et al. 2008). might be present and are often compensating Eyes of adult annelids might be present on the missing circular musculature (Purschke and head, palps, segments (usually laterally), or Müller 2006 ). even the pygidium (Purschke et al. 2006 ). However, many taxa, such as myzostomids, Annelids show a variety of reproductive sipunculids, or echiurids, are clearly deviating strategies, and sexual as well as asexual repro- from the pattern described above in various duction is well-documented for many taxa aspects, and all of them seem to have lost seg- (Wilson 1991 ; Bely 2006 ). For sexual reproduc- mentation convergently. Interestingly, all these tion, different types of free spawning, brooding, examples still show some traces hinting to a sec- and encapsulation of embryos in cocoons can be ondary loss of segmentation (Purschke et al. distinguished, and all types involve either plank- 2000; Hessling 2003; Kristof et al. 2008 ; Helm totrophic or lecithotrophic developmental stages et al. 2014 ). Other taxa such as clitellates and (Thorson 1950 ; Wilson 1991 ). Multiple modes many other sedentarians lost their parapodia. of development (poecilogeny) are reported for Siboglinidae (Pogonophora + Vestimentifera) some annelid species, with the spionid show many reductions as adaptation to their life- Streblospio benedicti as the best-investigated style in close association with bacterial endosym- example (Levin 1984 ; Zakas and Wares 2012 ). bionts (Schulze and Halanych 2003 ). Loss of key By far the most spectacular diversity of repro- characters in Annelida is well-documented and is ductive modes can be found across syllids, regarded as one of the problems to converge to a including swarming and external fertilization, well-accepted phylogeny of the whole group internal fertilization, viviparity, and parthogen- (Purschke et al. 2000 ; Bleidorn 2007 ; Miyamoto esis, as well as different forms of hermaphrodit- et al. 2013 ). ism (Franke 1999 ). Several annelid taxa show a Several systems of sensory organs are pronounced sexual dimorphism resulting in described for annelids, including a type of che- dwarf male forms as found, for example, in mosensory organ called “nuchal organ.” This some echiurids, siboglinids, or antonbruunids type of sensory organ can be found in the pos- (Spengel 1879 ; Hartman and Boss 1965 ; terior part of the prostomium and usually con- Worsaae and Rouse 2010). Not surprisingly, sists of ciliated supporting cells, sensory cells, annelids are also a prime example for the inves- and retractor muscles (Purschke 1997). tigation of heterochrony, and several putative Clitellates as well as several other annelids paedomorphic taxa have been hypothesized such as the basal branching Oweniidae and (Westheide 1987 ; Struck 2006; Bleidorn 2007 ; Magelonidae lack nuchal organs completely. Osborn et al. 2007 ). 198 C. Bleidorn et al.

Platynereis dumerilii as a Model for Evolutionary elegans. Important insights into the evolution Developmental Biology of segmentation, vision, and the nervous sys- Platynereis dumerilii is a marine annelid tem in Bilateria were provided by evolution- belonging to the errant family Nereididae, ary developmental studies on P. dumerilii , and which emerged as a thoroughly investigated since a number of labs now use this animal as model species. The life cycle of this indirect a model, important results with considerable developing gonochoric species with plankto- relevance for our understanding of animal trophic larvae is well established and control- evolution and development are likely to keep lable in the lab. Immature, atokous worms live emerging in the near future. in self-constructed tubes. Sexually mature, epitokous individuals, which appear mor- phologically different to atokous individuals, leave the tube and begin swimming to fi nd partners for spawning during the night. The day of swarming is controlled by an endog- enous lunar cycle which can be triggered artifi cially in the lab. Cultures were estab- lished in the lab in 1953 and are bred since then without interruption. Experimental tech- niques such as cell ablation, whole-mount in situ hybridization, RNA interference, and Morpholino knockdowns are routinely appli- cable. First transgenic lineages have been cre- ated, and a project sequencing the genome is underway for this species ( http://4dx.embl. de/platy/ ). Comparative genomic studies sug- gest that the genome of P. dumerilii retains a more ancestral organization compared to other protostomian model organisms such as Atokous juvenile of Platynereis dumerilii (After Drosophila melanogaster or Caenorhabditis Fischer and Dorresteijn (2004 ) )

EARLY DEVELOPMENT endoplasm, and hyaloplasm (Lillie 1906 , 1909 ), the latter two regions forming the cytoplasm. The Egg Structure and Fertilization ectoplasm contains large membrane- bound spher- ules, nuage (a germ line-specifi c organelle con- Annelids with planktotrophic development usu- taining several proteins), and intracellular ally have small, non-yolky eggs, whereas species membrane systems. The endoplasm contains yolk with lecithotrophic development bear larger and and lipid, interspersed with mitochondria, granular yolk-rich eggs (Irvine and Seaver 2006 ). bodies, and endoplasmatic reticulum. In contrast, Ultrastructural studies of annelid eggs are scarce the hyaloplasm (or teloplasm) is characterized by given the immense diversity of this group. One of the absence of granular bodies (Eckberg 1981 ; the best-investigated examples is the egg of parch- Jeffery 1985 ). Eggs of many annelid species show ment worms of the genus Chaetopterus . Different a clear polarity with an accumulation of develop- regions are distinguished based on staining mental factors in the cortex of future polar regions properties, divided into a cortical ectoplasm, (Dorresteijn 2005 ). The spatial distribution of 9 Annelida 199 maternal mRNA in the ectoplasm has been oblique to the egg axis due to an inclination of the described for Chaetopterus (Jeffery and Wilson mitotic spindle (see Chapter 7). This cleavage 1983 ; Jeffery 1985 ). After fertilization and before starts with two orthogonal cell divisions which initiation of the fi rst cleavage, a reorganization of generate four blastomeres, called A, B, C, and D the yolk-free hyaloplasm (teloplasm) has been (Costello and Henley 1976 ). Correlating with the observed in several annelids (Dorresteijn 1990 ; differing developmental modes in annelids, blas- Weisblat and Huang 2001 ). It has been shown for tomeres usually exhibit the same size (equal Platynereis dumerilii that after attachment of the cleavage) in species with indirect development sperm to the egg surface, cortical granules released and planktotrophic larvae, whereas direct devel- by exocytosis from the ectoplasm start forming an opers show pronounced differences in blastomere egg jelly on the outside, which removes supernu- size (unequal cleavage) (Anderson 1966 ; Arenas- merary sperm from its surface (Dorresteijn 1990 ). Mena 2007). However, exceptions to this trend Ultrastructural investigations of the ectoplasm of exist. For example, Platynereis dumerilii and eggs of P. dumerilii and the clitellate Theromyzon Platynereis massiliensis both show unequal spi- rude reveal an extensive framework of actin fi la- ral cleavage patterns, even though the former spe- ments that are involved in remodeling the egg sur- cies develops indirectly and the latter directly face after fertilization (Fernandez et al. 1987 ; (Schneider et al. 1992 ). In most cases unequal Kluge et al. 1995 ). cleavage is achieved due to positioning of the Following fertilization a reorganization of mitotic spindle. However, in some annelids this the endoplasm can be observed. The distribu- cleavage pattern is facilitated due to the presence tion of the two cytoplasmic domains can be cat- of membrane-bound polar lobes (Freeman and egorized into different types, which seem to be Lundelius 1992 ). Such a polar lobe is also restricted to certain annelid taxa (Shimizu reported for the myzostomid Myzostoma cirrif- 1999 ). Most investigated non-clitellate annelids erum (Eeckhaut and Jangoux 1993 ). The differ- (e.g., chaetopterids, nereidids, and onuphids) ent modes of spiral cleavage across annelids have show a stratifi cation of the endoplasm into two been thoroughly reviewed in Dorresteijn ( 2005 ). domains, and, in most cases, the clear hyalo- The future axis of the developing embryo is plasm (teloplasm) is localized at the animal already determined, with A and C corresponding pole (Wilson 1892; Huebner and Anderson respectively to the left and right side of the 1976 ; Jeffery and Wilson 1983). In contrast, embryo and blastomeres B to D defi ning the three domains can be distinguished in the clitel- antero-ventral to postero- dorsal axis (Nielsen late endoplasm, with teloplasm localized at 2004 ). Due to uneven cleavage, starting from the both the animal and vegetal poles of the egg third cell division, a shifting in the angle of the (Shimizu 1999 ; Weisblat and Huang 2001 ). mitotic spindles, which alternates during subse- These cytoplasmatic movements are coordi- quent divisions, becomes obvious in almost all nated by complex cytoskeletal mechanisms annelids. These shifts are either dextral (clock- which even seem to vary among taxa. Whereas wise) or sinistral (anti-clockwise) and lead to the in the leech Helobdella triserialis , microtu- name-giving spiral arrangement pattern of blas- bules are shown to play an important role, tomeres. By oblique divisions animal and vegetal movement of the teloplasm in the oligochaete daughter cells are generated, referred to as micro- Tubifex is orchestrated by an actin network mere quartets and macromere quartets. Some (Astrow et al. 1989 ; Shimizu 1995 ). annelids such as the opheliid Armandia brevis show equal cleavage also in the third cleavage, generating micro- and macromeres of equal size Cleavage (Hermans 1964 ). In the oweniid Owenia collaris and some leeches, micromeres are larger than Annelids develop by spiral cleavage which is macromeres in the eight-cell stage, a pattern characterized by cleavage furrows which are which is also known from several nemerteans 200 C. Bleidorn et al.

(Dohle 1999 ; Smart and Von Dassow 2009 ). clitellates show a cleavage pattern that nearly Several authors introduced the idea that a spe- obscures the original spiral mode of cleavage. In cifi c, phylogenetically conserved pattern of blas- these cases where yolk content and egg size are tomeres can be seen at this stage, termed the reduced, the embryo is nourished by the sur- “annelid cross.” This pattern is regarded as typi- rounding fl uid within the cocoon (Dohle 1999 ). cal for most annelids (including echiurans) but Several siboglinids show elongated eggs with a cannot be found in sipunculids, which show the high yolk content leading to an aberrant pattern so-called molluscan cross. However, a continuum of spiral cleavage (Southward 1999 ). In most of different variants between these patterns is annelids, the cleavage pattern shifts from spiral to demonstrated, and consequently, these concepts bilaterally symmetric after the formation of the have been neglected for phylogenetic purposes fourth quartet of micromeres (Meyer and Seaver (see also Chapter 6 ; Maslakova et al. 2004a ; 2010 ). In P. dumerilii cell fates of sister blasto- Nielsen 2004 ). The cell fate of individual blasto- meres along the animal-vegetal axis are specifi ed meres is conserved across annelids, and a specifi c by levels of beta-catenin. High levels specify nomenclature is used to trace the fate of vegetal sister cell fates, while lower levels spec- blastomeres throughout development, using ify animal sister cell fates. Interestingly, no beta- capital letters for macromeres and small letters catenin asymmetry is observed after the fi rst for micromeres (Conklin 1897 ; Costello and bilaterally symmetrical and transverse cell divi- Henley 1976 ; Nielsen 2004). A number is used as sions (Schneider and Bowerman 2007 ). prefi x to designate the quartet of which the Descendants of the fi rst micromere quartet macromeres or micromeres originated from. The (1a–1d) form larval head structures including the micromeres continue to divide, and daughter apical organ, larval eyes, and the head ectoderm cells inherit the name of their mother cell, with as well as the primary trochoblasts (Nielsen modifi cation, to trace its origin (animal vs. vege- 2004 ). The trochoblasts received their name as tal) (Fig. 9.3 ). An alternative cell nomenclatural they will give rise to the prototroch, and this system is in use for leeches (Dohle 1999 ). Many has been demonstrated for several annelids,

B - quadrant B - quadrant

A - quadrant A - quadrant 22 1b 21 2b 1 21 1b 2 12 2 3b 2b 1a 1b 2c 3a 22 11 C - quadrant 1a 1b 12 1 1c 2c 3B 11 2a 1 1a 22 3C 3A 1c 3c 12 11 1a 1c 4D 2 11 2a 21 1d 21 2 1d 1c 2d 3d 4d 12 1d 22 1d 2d 1

D - quadrant D - quadrant

Fig. 9.3 Diagram illustrating the spiral cleavage cell nomenclature in the 33-cell stage of an unequally cleaving embryo of Arenicola cristata (Child 1900 ). The four quadrants (A – D ) are indicated by colors 9 Annelida 201 e.g., capitellids, dinophilids, and nereidids 6 and 7 ). In Clitellata, a fi fth teloblast (M) is (Wilson 1892; Eisig 1898; Nelson 1904 ). derived from the 4d cell, specifying a mesodermal Trochophore larvae of Myzostomida lack or germband (Goto et al. 1999). Progenitors of 4d show a reduced prototroch (Rouse 1999 ), but as it form bilaterally symmetrical mesodermal anlagen has been shown for the nemertean Carinoma in Platynereis dumerilii (Ackermann et al. 2005 ; tremaphorus, this need not be refl ected in the for- Fischer and Arendt 2013 ). Prior to gastrulation, mation of trochoblasts (Maslakova et al. 2004b ). four secondary mesoblast cells bud from descen- Three sets of trochoblast cells are involved in dants of the 4d cell and show the morphology and prototroch formation, a pattern which is highly gene expression signature of primary germ cells conserved across Spiralia (Henry et al. 2007 ). (Rebscher et al. 2012). These primary germ cells Besides the primary and accessory trochoblasts, stay in mitotic arrest until individuals enter game- which are derived from the fi rst micromere quar- togenesis (Lidke et al. 2014). In C. teleta the 4d tet, this includes secondary trochoblasts formed cell generates few muscle cells, primordial germ by some descendants of the second micromere cells, and the anus (Meyer et al. 2010 ). It has been quartet (2a–2c). Some annelids deviate from this suggested that mesoteloblast‐like mesodermal pattern, e.g., the terebellid Amphitrite ornata , stem cells forming continuous mesodermal bands who lacks the accessory trochoblasts (Damen are part of the Pleistoannelida ground pattern and Dictus 1994 ). Other descendants of the sec- (Fischer and Arendt 2013 ). ond micromere quartet generally develop into the foregut (stomodaeum) as well as part of the ecto- derm (Nielsen 2004 ). The 2d cell is the somato- Gastrulation blast, developing into the major part of the body ectoderm posterior of the prototroch (Meyer and The process of gastrulation in annelids has been Seaver 2010 ). Using cell ablation studies, it has reviewed in detail by several authors (Okada been shown for Capitella teleta that the 2d cell is 1957 ; Anderson 1973 ; Weisblat and Huang 2001 ; responsible for organizing activity during early Irvine and Seaver 2006 ), and the following embryonic development, as well as bilateral sym- descriptions provide a generalized pattern found metry and dorso- ventral axis organization of the in clitellate and non-clitellate annelids. head, and formation of neural, foregut, and meso- Gastrulation of embryos with less yolk starts derm tissue (Amiel et al. 2013 ). In clitellates, with the invagination (embolic gastrulation) of four pairs of ectoteloblasts (called N, O, P, Q) are putative midgut cells, and epithelia derived from descendants of the 2d cell and give rise to four the micromere cap grow toward the ventral side. germbands including smaller cells (Dohle 1999 ; Mesoteloblasts can be found in a posterior posi- Goto et al. 1999 ). Cells derived from the third tion in the blastocoel, whereas ectoteloblasts are micromere quartet (3a–3d) form the foregut and located adjacent to them, below the larval ecto- ectomesoderm and might be the origin of proto- derm. The fate of the blastopore differs across nephridia (Nielsen 2004 ; Ackermann et al. 2005 ). annelid taxa, and protostomy, where the blasto- Interestingly, in C. teleta , mesodermal bands are pore becomes the mouth, is found in most anne- generated by 3c and 3d (Meyer et al. 2010 ). lid taxa. Notably, deuterostomy, where the Usually the mesoderm and endoderm are blastopore becomes the anus, has been demon- formed by cells of the fourth micromere quartet strated for eunicids (Åkesson 1967 ). The concept (4a–4d), as characteristic for spiral cleavage in of amphistomy, in which both the mouth and the Lophotrochozoa in general (Chapter 7; Gline anus are derived from the corresponding ends of et al. 2011 ). Of special interest is the fate of the 4d the blastopore, which was claimed to be present cell, which has been called “mesenteloblast” or in Polygordius (Arendt and Nübler-Jung 1997 ), “primary mesoblast” (Wilson 1898). This cell might not occur in any organism at all (Hejnol gives rise to the adult mesoderm in most spira- and Martindale 2009 ). Some differences apply lians including mollusks or entoprocts (Chapters for the gastrulation of yolk-rich annelid embryos. 202 C. Bleidorn et al.

Here, the process is rather described as epiboly, whole episphere in early developmental stages where the micromere cap grows over putative (e.g., in Chaetopterus ; see Fig. 9.4L ). The pro- midgut cells and teloblasts (Irvine and Seaver totroch may be formed by equatorially arranged 2006). In clitellates, the blastopore is found in the ciliary tufts (Myzostoma cirriferum ; see point where the germinal bands coalesce to form Fig. 9.4G ), or the whole larva may be covered by the germinal plate. cilia, and a defi ned prototroch is hardly distin- guishable, e.g., in early larvae of the eunicid Marphysa (Fig. 9.4E). An epispheral ciliated LATE DEVELOPMENT band is represented by the meniscotroch, which is only known for Phyllodocida (Bhaud and Larval Ciliary Bands Cazaux 1982 ; Rouse 1999 ). Forming a tuft of short cilia, the meniscotroch is located in a Larval morphological characters vary across dif- ventral position within the episphere. Posterior to ferent annelid families (Fig. 9.4 ). A trochophore the latter structure, some annelids possess a has distinct larval ciliary regions forming promi- ciliated band situated anterior to the prototroch – nent bands or tufts, and the presence of a pro- the akrotroch (Häcker 1896 ). Forming a com- totroch is regarded as defi ning (Bhaud and plete ring separated from the apical tuft and the Cazaux 1987 ; Rouse 1999). However, detailed prototroch, an akrotroch can be found in syl- investigations concerning the homology of the lids, orbiniids (e.g., in Scoloplos armiger , see respective ciliated regions within the different Fig. 9.5 ), onuphids, cirratulids, and several annelid families are lacking. At the anterior end Eunicida (Rouse 1999 ). of the episphere, the apical tuft marks the posi- Situated posteriorly to the prototroch, the tion of the larval apical organ, a feature well metatroch is represented by a ciliated ring that known for most invertebrate taxa with ciliated often beats opposed to the latter one and lies larvae (Marlow et al. 2014 ). Appearing early in in a pre-segmental (= peristomial) position development, the apical tuft forms a sensory (Strathmann 1993 ; Nielsen 2012 ). Being present region that is located in the direction of larval in most annelid families, a metatroch seems to movement but often disappears in early larval be absent in Echiura (Fig. 9.4J ) and Opheliidae. stages (see Chapter 7 for details on apical tuft For Capitellidae, Siboglinidae, and Syllidae, the morphology). Although an apical tuft is wide- presence of a metatroch is still discussed (Rouse spread within annelids, larvae without an apical 2000a ). Planktotrophic larvae of several polynoid tuft are known for most cirratulids, histriobdel- scale worms possess another bundle of long cilia, lids, lopadorhynchids, orbiniids, sabellids, and the oral brush, which seems also to be involved in tomopterids (Rouse 1999 ). feeding mechanisms (Phillips and Pernet 1996 ). The prominent prototroch is represented by an A prominent ventral ciliary band is represented equatorial ring consisting of usually compound by the neurotroch, which is known at least in some cilia formed by a group of specifi c trochoblasts annelid families including many sedentarian taxa, (Damen and Dictus 1994 ). Situated anterior to e.g., Orbiniidae (Fig. 9.5 ), Sabellidae (Fig. 9.6 ), the mouth opening, a prototroch is known for and Maldanidae. In both planktotrophic and leci- most annelids, mollusks, and entoprocts (Nielsen thotrophic developmental stages, the neurotroch 2012 ). Dividing the larval body in an anterior forms a distinct ventral ciliated area interconnect- episphere and a posterior hyposphere (see ing proto- and telotroch, which often appears later Chapter 7 ), the prototroch is present mainly in in larval development (Rouse 1999 ). The telo- planktotrophic annelid larvae and some leci- troch, defi ned as a posterior ring of cilia used for thotrophic stages but absent in direct developing locomotion (Strathmann 1993 ), also appears later taxa such as clitellates, aelosomatids, and histri- in development and is known for both planktotro- obdellids (Rouse 1999 ). In some annelid taxa, the phic and lecithotrophic developmental stages cilia of the prototroch may cover almost the (Strathmann 1993 ). The telotroch marks the 9 Annelida 203

A BC

EF D

G HI

J K L

Fig. 9.4 Diversity of annelid trochophore larvae. Anterior Myzostoma cirriferum (Myzostomida) after Eeckhaut and (apical) is up in all aspects. ( A ) Polygordius sp. Jangoux ( 1993 ). (H ) Platynereis dumerilii (Nereididae) (Polygordiidae) after Woltereck (1904 ). (B ) Magelona fi li- after Fischer and Dorresteijn (2004 ). (I ) Phascolosoma per- formis (Magelonidae) after Wilson (1982 ). (C ) Eurythoe lucens (Sipuncula) after Jaeckle and Rice (2002 ). (J ) complanata (Amphinomidae) after Kudenov (1974 ). (D ) Urechis caupo (Echiura) after Pilger (2002 ). (K ) Osedax sp. Owenia collaris (Oweniidae) after Smart and Von Dassow (Siboglinidae) after Rouse et al. (2009 ). (L ) Chaetopterus (2009 ). ( E ) Marphysa sanguinea (Eunicidae) after variopedatus (Chaetopteridae) after Henry (1986 ). Prevedelli et al. (2007 ). (F ) Phyllodoce maculata Abbreviations: at apical tuft, ch chaetae, lc lateral cilia, mt (Phyllodocidae) after Voronezhskaya et al. (2003 ). ( G ) metatroch, nt neurotroch, pt prototroch, tt telotroch 204 C. Bleidorn et al.

ak cilia eye pt pt

mt mt

ch1

nt A B tt

C D

ak cilia ak cilia eye mouth eye

pt pt nt mt mt

g1 g1 g2 ch2 g2 g3 nt

ch5

tt tt E F G H

Fig. 9.5 Development of Scoloplos armiger (intertidal sal view. (F ) Late 6-day embryo, lateral view. (G ) Early clade) after Anderson ( 1959 ). Anterior (apical) is up in all 7-day embryo, dorsal view. ( H ) early 7-day embryo, ven- aspects. ( A ) Unfertilized egg. (B ) Early 4-day embryo, tral view. Abbreviations: ak akrotroch; ch 1, ch 2, ch 5 dorsal view. ( C) Late 5-day embryo, dorsal view. (D ) Late chaetiger 1, 2, 5; g 1, g 2, g 3 gastrotrochs of chaetigers; 5-day embryo, ventral view. ( E) Late 6-day embryo, dor- mt metatroch; nt neurotroch; pt prototroch; tt telotroch

position of the posterior growth zone (Nielsen trochophore (Fig. 9.7 ). In this developmental 2012). Further ciliated bands that may occur in stage, the fi rst signs of segmentation are visible, several families are the gastro- and nototroch e.g., the formation of the fi rst parapodia and (= segmentally arranged ventral and dorsal ciliary chaetae. In accordance with individual develop- bands), which are sometimes referred to as ment, several subdivisions of the metatrochopho- paratrochs (Bhaud and Cazaux 1982 ). ral stage are possible (Fischer et al. 2010 ). In some terebellids and pectinariids, the metatrochophore builds a tube and is called aulo- Post-trochophore Development phore (Bhaud and Cazaux 1982 ). and Larval Forms In Oweniidae a special type of trochophore occurs, the so-called mitraria (Fig. 9.4D ). The Within annelid ontogeny, the metatrochophoral mitraria larva exhibits prominent proto- and stage usually follows the prototrochophore/ metatrochal bands, as well as an apical tuft. 9 Annelida 205

A B

CD

EF

Fig. 9.6 Development of the lecithotrophic developmen- nonfeeding stage. ( D ) Shortly before metamorphosis, the tal stages of Megalomma vesiculosum revealed by anti- late nectochaete has lost the main ciliary bands and starts tubulin staining. All images are in ventral view except of development of the adult tentacles (te ). ( E) After meta- ( D) which is in dorsal view. Anterior is up. Confocal morphosis the juveniles settle within a tube and start feed- maximum projections. ( A) The early, nonfeeding trocho- ing. The parapodia ( pa ), the tentacles (te ), and remnants phore exhibits a prominent prototroch ( pt). An apical tuft of the metatroch ( mt) are exhibited. (F ) Late juvenile is lacking. ( B) The later trochophore gains a well-devel- worms show an adult-like morphology. The tentacles (te ) oped prototroch ( pt ) and cilia (ci ) at the anterior pole. are well-developed, and the animals start to elongate by Furthermore, the ventral neurotroch ( nt ) and the posterior posterior segment addition. an anus, ci cilia, mo mouth telotroch ( tt) develop at this stage. ( C) The early opening, mt metatroch, ne nephridia, nt neurotroch, pa metatrochophore exhibits three pairs of parapodia ( pa ) parapodia, pt prototroch, tt telotroch. Scale bars = 100 μm and a metatroch (mt ). Neurotroch (nt ), prototroch (pt ), and (© Conrad Helm, 2015. All Rights Reserved) telotroch ( tt) are still present in this free-swimming but 206 C. Bleidorn et al.

AB

CD

E F

Fig. 9.7 Development of Platynereis dumerilii . (A ) of body segments. Combined after Fischer et al. ( 2010 ). Cleaving embryo, where the third cleavage forms micro- Abbreviations: ac anal cirrus, ae adult eyes, adc anterior meres and macromeres. ( B) Early trochophore, with pro- dorsal cirrus, ant antenna, apt apical tuft, avc anterior ven- totroch and apical tuft. ( C ) Late trochophore, with tral cirrus, ch chaetae, chs chaetal sac, j jaws, ld lipid simultaneous appearance of the fi rst three larval segments. droplet, le larval eye, mg , midgut, mm macromere, mt ( D ) Mid metatrochophore, with developing chaetae reach- metatroch, pat 1 , fi rst paratroch, pat2 second paratroch, pl ing over the body wall. (E ) Early nectochaete, with forma- palps, pp parapodia, pro proctodeum, pt prototroch, sf sto- tion of the metatroch and elongation of the trunk. ( F ) modeal fi eld, sto stomodeum, tt telotroch, 4CS 4th chae- Juvenile, with rapidly growing jaws and further addition tigerous segment, 5CS 5th chaetigerous segment 9 Annelida 207

Notably, all ciliary bands are monociliated, an eaten or discarded (Rouse 2006 ). Such a transi- unusual feature for annelid larvae. The hypo- tion of lifestyles seems to be less distinct or miss- sphere of the mitraria is strongly reduced, and the ing in annelids with lecithotrophic developing juvenile segmental body develops within the lar- stages, where ciliary bands are absorbed or the val body (Wilson 1932 ; Smart and Von Dassow chaetal morphology changes (Figs. 9.5 and 9.6 ). 2009). Another unusual larval type is repre- Although late development and subsequent sented by the rostraria in Amphinomidae and metamorphosis may differ between several taxa, Euphrosinidae (Mileikovsky 1960 , 1961 ). After in almost all annelid larvae, the larval episphere a trochophore stage with a proto- and metatroch becomes the adult prostomium, and the poste- and an apical tuft (Fig. 9.4C ), the episphere of rior hyposphere becomes the pygidium and the the metatrochophore elongates, and tentacles are posterior growth zone (Fig. 9.8). The remaining formed for feeding (Jägersten 1972 ). Remarkably hyposphere forms the peristomium, which lacks elongated metatrochophore stages can be found chaetae in adult annelids (Nielsen 2004 ). The within siboglonids (Southward 1999 ). The segmented body between the peristomium and metatrochophore of investigated vestimentiferan the pygidium develops by segment formation siboglinids is sessile and bears a prostomium, a from the posterior growth zone (Irvine and peristomium, and two chaetigers. A prototroch, Seaver 2006 ). a neurotroch, and an apical organ are present as well as juvenile/adult organs such as tentacles and pyriform glands (Bright et al. 2013 ). Larval Feeding Modes The end of the metatrochophore stage is usu- ally marked by the point when the parapodia are Several types of larval feeding behaviors and fully developed. If present, the next larval stage is developmental modes occur in different annelid represented by the nectochaete (Fig. 9.7E ), which families, mostly divided into either feeding and is characterized by the presence of functional para- free-swimming planktotrophic larvae with “indi- podia which are used mainly for swimming. In rect” development or nonfeeding and mostly less Poecilochaetus (Spionidae) this stage is some- motile embryonic and juvenile forms with times called nectosoma; in other spionids, it refers “direct” development. The latter rely on maternal to the chaetosphaera stage (Bhaud and Cazaux sources of nutrition in the form of yolk stored in 1982 ). In this stage or the previous one, most lar- the egg during oogenesis, feeding on yolk-rich vae start body elongation and segment formation nurse eggs, or translocation of nutrition directly through the posterior growth zone (Irvine and from the parent (Qian and Dahms 2006 ). An Seaver 2006 ). A unique swimming larval form overview of larval feeding in annelids is summa- called pelagosphaera is known for Sipuncula (Rice rized by Rouse ( 2006). As direct development 1976 ). The body of this larval type can be divided occurs without a metamorphosis, developing into three body regions: head, mid region includ- stages are usually referred to as “embryonic” or ing metatroch, and a large trunk. These larvae can “juveniles,” avoiding the term “larvae” (Nielsen be either lecithotrophic or planktotrophic forms, 2009 ; Winchell et al. 2010). However, debates whereas the latter can live up to 6 months in the remain about the defi nition of the term “larva,” plankton (Jaeckle and Rice 2002 ). and alternative terminologies exist (McEdward After the nectochaetal (or pelagosphaera) and Janies 1993 ; Pechenik 1999 ). stage, metamorphosis occurs. During metamor- Many authors regard a biphasic life cycle phosis, the free-swimming larvae change behav- including a planktotrophic trochophore larva as ioral characteristics and start an adult-like representing the plesiomorphic condition of lifestyle as juveniles. This metamorphic step can annelid development (Heimler 1988 ; Nielsen differ drastically between various species and is 2012 ). Nevertheless, this idea is doubted by some markedly pronounced in larvae of Polygordius , authors (Haszprunar et al. 1995 ; Rouse 2000a , b ). which rupture the larval body that is later either Based on cladistic analyses and ancestral state 208 C. Bleidorn et al.

Fig. 9.8 Development of annelids indicating the contri- pygidium develop from descendants of the (D) quadrant. bution from the four quadrants (A–D), after Nielsen Abbreviations: at apical tuft, pt prototroch, mt metatroch, (2004 ). Note that the whole segmented body and the tt telotroch, nt neurotroch, gt gastrotroch reconstruction, multiple events in the evolution planktotrophy seems to be the likely ancestral of feeding larvae from lecithotrophic ancestors state in annelids. seem more parsimonious to assume. According Most planktotrophic larvae exhibit a prominent to this hypothesis, the prototroch had a primary proto- and metatroch, used for locomotion function for locomotion and became indepen- and food uptake by “downstream feeding” dently associated with feeding in several lineages (Rouse 2000a ). This is the case for larvae of (Rouse 2000a ). By studying larval forms of Amphinomidae, Chrysopetalidae, Glyceridae, sabellids, Pernet (2003 ) was able to demonstrate Nephtyidae, Oweniidae, Pectinariidae, Polynoidae, the persistence of reduced ciliary structures for and Sabellaridae. However, other annelid families food uptake in nonfeeding larvae. Consequently, with planktotrophic larval development use differ- it is suggested that the direction of evolution goes ent modes of feeding behavior. Taxa with nonfeed- from planktotrophy to lecithotrophy in this case. ing larvae possessing maternally derived nutrition Further on, a functional relation between egg size can be found all over the annelid tree within vari- and gut development has been hypothesized. ous families, and lecithotrophy may have evolved Non-feeding stages usually bear larger eggs, and secondarily (Rouse 2000a ). A special case of leci- due to the increased cell size, gut development thotrophic development is represented by adel- might be hindered, resulting in nonfunctional phophagy, where larvae develop by uptake of guts in nonfeeding stages (Pernet 2003 ). As this nutrients from nurse eggs, which are only pro- scenario may be generalized for all annelids, duced as nutrition reserve (Fig. 9.9 ), as in some 9 Annelida 209

AB

CD

E F

Fig. 9.9 Development of ciliation in larvae of the adel- ( vp), and a telotroch (tt ) are present. First signs of segmen- phophagous spionid Boccardia cf. polybranchia revealed tation are visible in this stage. ( E ) In the late metatrocho- by anti-tubulin staining. All images are in ventral view; phore the ventral ciliated patches ( vp ) are reduced. anterior is up. Confocal maximum projections. ( A ) Early Instead, a distinct neurotroch (nt ) and several gastrotrochs larvae lack ciliary regions and are full of yolk. ( B ) The ( gt) form. ( F) In the nectochaetal stage, shortly before trochophoral stage attaches to a nurse egg ( ne) and starts leaving the egg capsule, three bands of gastrotrochal cili- to digest its nutrients. Ciliation is only exhibited within ary bands ( gt ) are developed, and notopodial cilia (no ) are the mouth opening ( mo) and in the region of the ventral exhibited. The remaining ciliated bands are still present. ciliated patches ( vp), which are used for attachment. ( C ) gt gastrotroch, mo mouth opening, mt metatroch, ne nurse The late trochophore develops a distinct prototroch ( pt ) egg, no nototroch, nt neurotroch, pt prototroch, tt telo- arranged of ciliated patches and a less-prominent troch, vp ventral ciliated patch. Scale bars = 50 μm (© metatroch ( mt ). (D ) In the metatrochophoral stage, the Conrad Helm, 2015. All Rights Reserved) prototroch ( pt), metatroch (mt ), the ventral ciliated patches 210 C. Bleidorn et al. spionid taxa (Gibson and Carver 2013 ). Moreover, surface of the embryo. During gastrulation, the poecilogeny, showing different types of develop- germinal bands from both sides of the embryo ment in one species, seems to be common in some coalesce into the germinal plate, the origin of spionid genera (Blake and Kudenov 1981 ; Levin segments. The stereotyped cell divisions of all 1984 ; Chia et al. 1996 ). blast cells contribute to the forming segments (Shimizu and Nakamoto 2001 ; Weisblat and Huang 2001 ; Irvine and Seaver 2006 ). Each Segmentation ectodermal band contributes neural and epider- mal progeny, but two thirds of the neurons are Segmented annelids generate their fi rst seg- derived from the N teloblast (Weisblat and ments, usually simultaneously, as larvae, and Huang 2001 ). Ectodermal segmentation can be later segments are sequentially added from a divided into two steps. In the fi rst step, distinct posterior growth zone (Irvine and Seaver 2006 ). cell clusters are generated autonomously by each Consequently, some authors differentiate bandlet. These separate clusters are aligned with between primary and secondary segments, an the cell cluster derived from the mesodermal idea that goes back to Iwanoff ( 1928) who postu- bandlet in a second step (Shimizu and Nakamoto lated a distinct ontogenetic origin for both sets of 2001 ). It has been shown experimentally for segments. Segment formation on the cellular Helobdella that hemilateral ablation of mesoder- level is well understood for clitellate embryos, mal precursor cells results in the loss of ectoder- which are all direct developers and often show mal segmental organization. In contrast, huge and therefore experimentally manipulable mesodermal boundaries are determined autono- eggs. Segmentation in leeches is strongly corre- mously, without positional cues from ectodermal lated with cell division patterns of teloblast cells tissue (Blair 1982 ). and their descending blast cells (Fig. 9.10). All The cellular basis of segment generation in teloblasts originate from descendants of the D non-clitellate annelids is less well studied. One quadrant, with a pair of mesoteloblasts (M) reason seems to be that they are more diffi cult to being derived from 4d and four pairs of ectoder- handle experimentally due to the smaller size of mal teloblasts (N, O, P, Q) from 2d descendants. their embryos and larvae (Irvine and Seaver Mechanisms of the specifi cation of the ectotelo- 2006 ). An obvious difference is the absence of blast lineage are different between the hirudin- large visible teloblasts in non-clitellate annelids ean Helobdella and the oligochaete Tubifex . In (Seaver et al. 2005 ). However, using semiauto- Helobdella the O/P teloblasts constitute an mated cell tracking, mesoteloblast-like stem cells equivalence group, as they are both pluripotent were revealed for Platynereis dumerilii which and may subsequently follow either the O or the also form mesodermal bands of daughter cells P fate (Weisblat and Blair 1984 ). On the con- (Fischer and Arendt 2013 ). For this species, two trary, in Tubifex , the fate of the P teloblast is distinct sets of stem cells could be described for determined by birth, whereas the O teloblast is the posterior growth zone, where many rounds of initially pluripotent and is restricted to the fate of division of small populations of teloblast-like the O lineage due to signaling from the P lineage stem cells generate new segments (Gazave et al. (Arai et al. 2001). Each teloblast undergoes 2013 ). In contrast, Seaver et al. ( 2005) could not repeated series of unequal division, producing fi nd evidence of a teloblastic growth zone in bandlets of the so-called blast cells. The N and Q Capitella teleta and the serpulid Hydroides lineages produce two different types of blast elegans using incorporation of BrdU, therewith cells, which appear in alternation. The four ecto- confi rming older studies (Wilson 1890 ; Shearer dermal bandlets of each side of the bilateral 1911). Instead, segments in larvae arise from a embryo join and form together with the meso- fi eld of mitotically active cells located in lateral dermal band, the germinal band, which lies at the body regions. However, the authors could not 9 Annelida 211

Fig. 9.10 Schematic representation of the stem cell-mediated, lineage- 140 dependent segmentation in leeches and other clitellate annelids. Anterior is to the top . Pairs of diagonal lines indicate discontinuities in the depicted structures. One bilateral pair of mesodermal stem cells (M teloblasts) and four bilateral pairs of ectoder- mal stem cells (N , O/P , O/P , and Q teloblasts) 90 constitute the posterior growth zone. Two types of blast cells are contributed by the N lineage, designated ns (red ) and nf 80 ( blue ), which arise in alternation. The numbers on the left side indicate the progressing time during segment formation given in hours of clonal age. Arrows indicate the delimitation of two ganglionic primordia 29 (From Rivera and Weisblat 26 2009 , with permission from the publisher) 10 Q 0 M P N

rule out if inconspicuous teloblast-like cells Muscular System might be present. For Chaetopterus , it has been suggested that at least the fi rst 15 segments are Annelids show a huge variety of muscular orga- formed by subdivision of existing anlagen and nization, with longitudinal musculature orga- not by a posterior growth zone (Irvine et al. nized in separate bands or massive plates and 1999 ). Similarly, formation of repetitive struc- circular musculature that can be fully developed, tures in myzostomids differs from an addition incomplete, or even completely missing (Tzetlin governed by a posterior growth zone. As such, and Filippova 2005 ). Therefore, it comes without during development the third pair of parapodial surprise that differences in the development of structures appears fi rst, followed by the fourth the muscular system have been found across the pair, second and fi fth pair (simultaneously), and investigated taxa. Phalloidin labeling coupled fi rst pair (Jägersten 1940 ). Future studies of espe- with confocal microscopy revealed an origin of cially non-clitellate annelids are necessary to fur- muscular development posterior of the apical ther assess the existing variability in the process organ in the phyllodocid Phyllodoce groenland- of segment formation throughout Annelida. ica, with distinct transversal muscles starting to 212 C. Bleidorn et al. grow posteriorly. Subsequently, several longitu- and the leech Erpobdella octoculata , anterior dinal muscle fi bers start to develop and grow in circular muscles are formed synchronously posterior direction. Simultaneously, outer circu- (Wanninger et al. 2005 ; Bergter et al. 2007 ; lar muscle fi bers begin to appear in a progression Kristof et al. 2011 ; Helm et al. in press ). from anterior to posterior. The longitudinal mus- Muscular development in the non-segmented cle fi bers reach the anal region approximately sipunculids as analyzed for Phascolion strom- 7 days after hatching, and additional circular bus , Phascolosoma agassizii , Thysanocardia muscle fi bers forming distinct rings develop from nigra, and Themiste pyroides shows that the fi rst anterior to posterior. Additional longitudinal anlagen of circular body wall musculature muscle fi bers develop in the dorsal region, form- appear simultaneously. Rudiments of four lon- ing a continuous layer. Musculature of the diges- gitudinal retractor muscles appear at the same tive system is hardly recognizable in early stages. time, with longitudinal muscle fi bers forming a Notably, the organization of the body wall mus- pattern of densely arranged fi bers around the culature starts before the formation of the fi rst retractor muscles (Kristof et al. 2011 ). segments (Helm et al. 2013 ). Similarities to this kind of musculature development have been found in several other annelids. As such, an ante- Neurogenesis rior origin in either lecithotrophic embryos or planktotrophic larvae is also reported for, e.g., Annelids show a huge variety of adult nervous capitellids, clitellates, nereidids, and sabellariids system organization, and until today the ancestral (Hill 2001 ; Bergter and Paululat 2007 ; Hunnekuhl ground pattern remains under discussion (Bullock et al. 2009 ; Brinkmann and Wanninger 2010a ; 1965; Orrhage and Müller 2005 ). The develop- Fischer et al. 2010 ). No circular musculature ment of the nervous system, however, has been could be detected in developing stages of the investigated in a surprisingly small number of maldanid Axiothella rubrocincta , even though taxa. Several transmission electron microscopy they are present in adults (Brinkmann and (TEM)-based studies on the larval nervous sys- Wanninger 2010b ). tem of phyllodocids and serpulids were published In contrast to the description for Phyllodoce , by Lacalli (1981 , 1984 , 1986 ). Some detailed musculature of the digestive system develops comparative studies were conducted concerning before the body wall musculature in planktotro- the anatomy of the larval apical organ. phic larvae of serpulids and sabellariids Immunocytochemical studies revealed an almost (McDougall et al. 2006 ; Brinkmann and universal occurrence of an apical organ with Wanninger 2010a ). Temporal shifts in the devel- fl ask-shaped cells in larvae of Annelida, opmental onset of several muscle groups, a phe- Mollusca, Entoprocta, and Platyhelminthes, nomenon described as heterochrony, are a exhibiting FMRFamide- and serotonin-like common theme when comparing myogenesis immunoreactivity (e.g., Hay-Schmidt 2000 ; between different and even closely related anne- Wanninger 2009 ). Usually, the apical organ in lid species and are even more pronounced in the annelid trochophores is simple, containing a few comparison of planktotrophic with lecithotro- fl ask-shaped cells which have slender necks, phic developing species (McDougall et al. 2006 ; dense cytoplasm, and a single projecting cilium Brinkmann and Wanninger 2010a ; Helm et al. (Lacalli 1981 ). Whereas these cells are missing 2013 ). As in Phyllodoce , many annelid species in echiurans and many other annelids, sipuncu- show a successive appearance of circular mus- lans show a more complex apical organ with up culature from anterior to posterior, which has to ten fl ask-shaped cells (Wanninger 2008 ). been also described, e.g., for the tubifi cid Marlow et al. (2014 ) analyzed the molecular fi n- Limnodrilus and the serpulid Filograna implexa gerprint of apical organ cells in Platynereis (Bergter et al. 2007 ; Wanninger 2009 ). In con- dumerilii . They found that orthologs of six3 and trast, in sipunculids, Platynereis massiliensis foxq2 are involved in the formation of the apical 9 Annelida 213 plate, whereas the apical tuft is formed in a cen- neurogenesis shows major shifts between com- tral six3 -free area of the apical plate. pared species and is regarded as cases of heter- Besides this, only few comprehensive studies ochrony (McDougall et al. 2006 ; Brinkmann and for developmental sequences of annelids com- Wanninger 2008 ; Helm et al. 2013 ). bining immunocytochemical staining coupled A different picture is found in direct develop- with confocal laser scanning microscopy exist ing lecithotrophic species as investigated for (Hessling 2002 ; Hessling and Westheide 2002 ; nereidids. In Nereis arenaceodentata, the ner- Voronezhskaya et al. 2003; McDougall et al. vous system has developed already much of the 2006 ; Brinkmann and Wanninger 2008; Kristof complexity of the adult at hatching. This includes et al. 2008 ; Fischer et al. 2010; Winchell et al. a large brain and the presence of circumesopha- 2010 ; Helm et al. 2013, in press ). Main targets geal connectives, nerve cords, and segmental for these studies were serotonin, a biogenic nerves. Within 1 week after hatching, cephalic amine involved in neuronal signaling, and the sensory structures and brain substructures are dif- neuropeptide FMRFamide. Labeling of tubulin ferentiated, and the nervous system architecture is additionally used to stain neurotubules. In resembles that of adults (Winchell et al. 2010 ). A summary, these studies show that neurogenesis similar developmental pattern of the nervous sys- in annelids is variable, following different devel- tem has been described for Platynereis massilien- opmental pathways. Planktotrophic larvae typi- sis (Helm et al. in press ). cally bear a serotonergic nerve ring underlying Analyses of the development of the nervous the prototroch and an apical organ that bears system of the non-segmented Echiura and serotonergic and FMRFamidergic cells. The Sipuncula gained major interest, as they pro- development of the larval nervous system usu- vided direct ontogenetic evidence for the indi- ally starts from two subsystems (Fig. 9.11 ). rectly inferred loss of segmentation in these taxa FMRFamidergic immunoreactivity increases as suggested by molecular phylogenies. Using from anterior toward posterior during nervous immunocytochemistry, a metameric organiza- system development. In Phyllodoce and some tion of the nervous system has been demon- other annelids, a single serotonergic neuron strated for two echiuran species: Urechis caupo , located at the posterior pole of the larva is pres- which has planktotrophic larvae, and Bonellia ent (Fig. 9.11A). From here, anteriorly project- viridis with directly developing lecithotrophic ing nerve fi bers start to grow, outlining the future stages (Hessling 2002 ; Hessling and Westheide ventral nerve cords (Voronezhskaya et al. 2003). 2002 ). The development of the nervous system Such a posterior origin of serotonin-like immu- in Bonellia viridis proceeds from anterior to noreactivity was also detected in another phyl- posterior. This is obvious in early larvae, which lodocid, in syllids, nereidids, and orbiniids show a full set of serotonergic perikarya in the (Orrhage and Müller 2005 ; McDougall et al. anterior region, while this pattern is incomplete 2006 ; Fischer et al. 2010; Helm et al. 2013 ). In in the posterior area. This pattern suggests the contrast, the investigated sabellariids, spirorbids, presence of a posterior growth zone (Hessling and sipunculans show no evidence for a posterior and Westheide 2002 ). Similarly, in larval stages serotonergic cell (Brinkmann and Wanninger of Urechis caupo, a serial repetitive distribution 2008 ; Kristof et al. 2008 ; Brinkmann and of serotonin-containing neurons and their cor- Wanninger 2009 ). Later, the adult nervous sys- responding pairs of peripheral nerves, both tem starts to develop along pathways established formed in an anterior-posterior gradient, imply by the earliest peripheral neurons of the larva. a segmental pattern. Moreover, larvae show a However, other authors propose a separate devel- paired origin of the ventral nerve cord (Hessling opment of the larval and adult nervous system 2002 ). Another case of “ontogeny recapitulating (Lacalli 1984 ) or fi nd that the larval nervous sys- phylogeny” has been demonstrated for sipuncu- tem is integrated in the adult one (Hay-Schmidt lans, where neurogenesis of Phascolosoma 1995). The relative timing of events during agassizii follows a segmental pattern. During 214 C. Bleidorn et al.

Fig. 9.11 Schematic representation of neuronal AB development in larval stages of Phyllodoce groenlandica exhibiting formation from two subsystems (Modifi ed from Helm et al. (2013 )) Diagrams are in ventral view; anterior is up. Major types of neuronal structures are color coded. ( A ) 0.5 days past hatching ( dph ). First serotonergic immunoreactivity appears at the posterior pole. (B ) 2 dph. Nervous system originates from posterior CD (serotonin) and anterior (FMRFamide). ( C ) 7 dph. Serotonergic and FMRFamidergic immuno- reactivities start to overlap to the greatest extent. ( D ) 11 dph. Numerous serotonergic cells are detectable within the epi- and the hyposphere. ( E ) 20 dph. Serotonergic and FMRFamidergic nerve cells are limited mainly to anterior regions. ( F ) 34 dph. The larval nervous system is fully developed. at apical tuft, pt prototroch EF

SerotoninFMRF Serotonin + FMRF 9 Annelida 215 development of this species, a pair of becomes ciliated, and the most proximal cells are FMRFamidergic and serotonergic axons gains separated during coelomogenesis. During coelo- four pairs of associated serotonergic perikarya mic cavity growth, the proximal part of the anlage and interconnecting commissures in an anterior- is passively opened, forming the metanephridial posterior progression. During later larval stages, funnel. A truncation of this process due to sup- the commissures disappear and the two seroto- pression of the separation of duct cells leads to a nergic axons fuse, forming a single ventral differentiation into a protonephridium, as, for nerve cord, and after cell migration, a nonmeta- example, observed in several taxa of the meric central nervous system can be found in Phyllodocida (Bartolomaeus 1999 ). The develop- adults (Kristof et al. 2008). Interestingly, neuro- ment of metanephridia in the non-segmented genesis of the sipunculan Phascolion strombus sipunculids as investigated for Golfi ngia minuta lacks any signs of a segmented origin, and sero- seems to be similar as in segmented annelids. An tonergic structures are missing completely in overview of annelid nephridial organs is given by their larvae, which may be the result of the Bartolomaeus and Quast (2005 ). abbreviated larval phase in this species Segmented annelids show a heteronomous (Wanninger et al. 2005 ). coelomogenesis, and the coelomic lining is of mesodermal origin. Prior to metamorphosis (if present), a pair of unsegmented coelomic cavities Nephridia and Coelomogenesis stretches out over the fi rst larval segments. This process has been studied in detail for the serpulid Most adult annelid taxa possess metanephridia as Spirorbis spirorbis, where two caudally located excretion organs; however, in the development mesodermal cell clusters proliferate cells, which they are usually preceded by protonephridia, merge to surround the gut ventrally close to the which can be found in larvae and sometimes also anus. Fluid starts to accumulate between spaces in developing juveniles (Bartolomaeus and Quast of desmosomes, and at the same time myofi brils 2005). These larval excretory organs were termed appear, and due to growth and separation pro- “head kidneys” by Hatschek (1886 ) and are cesses, this myoepithelium develops into the coe- located anteriorly in trochophore stages, closely lomic lining. After migration, the two coelomic behind the larval eyes (Bartolomaeus and Quast cavities meet dorsally, completely surrounding 2005 ). Later, homologous organs were also the gut. Postmetamorphic stages develop strictly described from lecithotrophic developmental segmental coelomic cavities during segment for- stages, as shown for the direct developing species mation. Coelomic cavities are highly reduced or Scoloplos armiger (intertidal clade), which has completely missing in leeches and several meio- no free-swimming larval stage (Bartolomaeus fauna annelids with a presumed progenetic origin 1998 ). Head kidneys are present in some echiu- (Koch et al. 2014 ). rans but are missing in sipunculid developmental stages. It is hypothesized that the ancestral state of larval annelid protonephridia was an organ GENE EXPRESSION composed of three cells: a terminal cell, a nephro- pore cell, and a duct cell (Bartolomaeus and Only few model annelids are well characterized Quast 2005 ). This simple construction has been concerning gene expression patterns. Prime can- modifi ed in adaptation to different developmental didate taxa are leeches of the genera Helobdella modes in several annelid lineages and especially and Hirudo , the sedentary annelid Capitella in planktotrophic larvae, where the nephropore teleta, and the errant annelid Platynereis dumeri- cell is often missing (Kato et al. 2011 , 2012 ). lii . Besides this, some studies exist for the Adult segmental nephridia differentiate from a chaetopterid Chaetopterus , some sipunculid single anlage, consisting of few cells which line a worms, and a few serpulids as well as for addi- small lumen fi lled with microvilli. This duct tional nereidid and clitellate species. These 216 C. Bleidorn et al.

studies mainly used candidate gene approaches, In Chaetopterus , engrailed is expressed during where orthologs were chosen based on studies in all larval stages in different structures or organs, arthropods and vertebrates. Main points of and no signs of a putative segment polarity pat- interest are the genomic basis of segmentation, tern of expression are obvious (Seaver et al. Hox gene expression, and nervous system 2001). Congruently, no conserved segment polar- development as well as gastrulation and gut ity pattern was found investigating the expression development. of this gene in developing C. teleta or Hydroides elegans individuals (Seaver and Kaneshige 2006 ). Finally, ablation of individual cells Segmentation expressing engrailed in the leech Helobdella did not hinder remaining segmental clones in their Metameric segmentation can be found in verte- normal development (Seaver and Shankland brates, arthropods, and annelids, and the distant 2001 ). Consequently, the establishment of seg- phylogenetic position of these taxa gave rise to ment polarity in the leech (and possibly many the question of how often this feature evolved in other annelid taxa) seems to be independent of animals (Seaver 2003). Traditionally well inves- cell interactions across the anterior- posterior axis tigated is the molecular background of segmenta- as known for arthropods (Seaver and Shankland tion in vertebrates and arthropods, which show 2001 ). profound differences (Tautz 2004 ). Given the fact A set of pair rule genes is expressed in that a close relationship between arthropods and Drosophila and many other arthropods to pattern annelids was suspected as formulated in the the embryo across the anterior-posterior axis, Articulata hypothesis (Scholtz 2002 ), a possible including eve , hairy , and runt (Damen 2007 ). All common ancestry of segmentation in these taxa or some of these genes were investigated in detail became a focus of many evolutionary develop- for C. teleta and Helobdella robusta. In contrast mental studies of annelids. Genes or gene fami- to Drosophila , where it is expressed in stripes in lies identifi ed to play a vital role in segment the growth zone, the Capitella ortholog of hairy formation in arthropods were used as candidates ( Cap - hes1) shows an expression limited to a in several studies (Wedeen and Weisblat 1991 ; small band of cells in each larval segment. In Prud’homme et al. 2003 ; Seaver and Kaneshige juveniles its expression is limited to a small 2006 ; Saudemont et al. 2008 ; Dray et al. 2010 ; mesodermal domain of the posterior growth zone Steinmetz et al. 2011 ). (Thamm and Seaver 2008 ). In vertebrates and Best investigated is the molecular background some arthropods, the expression of hairy is con- of segmentation in Platynereis dumerilii and trolled by the Notch pathway (Stollewerk et al. Capitella teleta. For Drosophila it has been dem- 2003 ). In Capitella , Notch and hairy do not show onstrated that para-segmental borders are gener- a broadly overlapping expression, with a Notch ated by an interaction between the segment localization in already formed segments, anterior polarity genes wingless and engrailed (Tautz to the hairy signal (Thamm and Seaver 2008 ). In 2004 ). The gene wingless (or Wnt1 ) is part of the Helobdella , hairy is expressed in teloblasts and Wnt gene family, and engrailed is a primary blast cells. The expression peak corre- homeodomain-bearing transcription factor. As in lates with the production of blast cells by the arthropods, a role in segment formation is sug- teloblasts. However, no striped pattern suggest- gested for this pair of genes in P. dumerilii ing a pair rule function was found (Rivera et al. (Prud’homme et al. 2003 ), where an expression 2005 ). Similarly, Notch is also expressed in telo- of continuous ectodermal stripes is observed for blasts and blast cells, and functional studies these genes at the border of the segments during revealed that the disruption of the Notch / hairy their formation. However, investigation of other signaling results in a disruption of segmentation annelid taxa questions the conserved nature for (Song et al. 2004 ; Rivera and Weisblat 2009 ). For engrailed as a “segmentation gene” in annelids. Platynereis dumerilii , 15 hairy paralogs could be 9 Annelida 217 identifi ed, which are expressed in mesodermal for these species (Iwasa et al. 2000 ; Werbrock tissue, forming segments, and during neurogene- et al. 2001 ). sis, where it may be involved in the patterning of NKL genes are a family of homeodomain the nervous system (Gazave et al. 2014 ). transcription regulators that are involved in the However, these authors also found no overlap patterning of mesodermal derivatives in with the expression of Notch . Drosophila (Holland 2001; Jagla et al. 2001 ). The expression patterns of the arthropod pair The expression of seven genes of this cluster has rule genes eve and runt do not suggest a similar been investigated in developing Platynereis role in Capitella . Instead, the expression of runt dumerilii , and all are involved in the specifi cation can be found in the brain and ventral nerve cord, of mesodermal derivatives including muscular as well as the fore- and hindgut. Two eve paralogs precursors (Saudemont et al. 2008 ). Notably, fi ve were characterized for Capitella , both showing a of the investigated genes (NK4 , Lbx , Msx , Tlx , complex expression pattern, which does not cor- and NK1 ) show an expression in complementary respond to segmental stripes as expected by stripes in the mesoderm and/or ectoderm of results from Drosophila (Seaver et al. 2012 ). developing segments. Moreover, genes of the Likewise, de Rosa et al. (2005 ) did fi nd such a Hedgehog signaling pathway show a similar pattern of eve expression in developing striped pattern of expression, and segment forma- Platynereis dumerilii . However, these authors tion in P. dumerilii is disrupted when treated with speculate about a role of this gene in the posterior molecules antagonistic to this signaling (Dray addition of segments. A detailed functional study et al. 2010 ). for eve has been conducted for Helobdella (Song Wnt genes regulate a wide range of develop- et al. 2002 ). Segments arise sequentially from mental processes, including axis elongation and fi ve pairs of teloblasts in leeches (see above), and segmentation (Cadigan and Nusse 1997 ). This eve is expressed in these teloblasts and their pri- gene family ancestrally includes 13 paralog mary blast cells in Helobdella . Later embryos groups, of which several metazoan lineages lost express eve in cells of the ventral nerve cord some of the genes (Janssen et al. 2010 ). In which stem from the N teloblast. Morpholino Platynereis dumerilii and Capitella teleta , all knockdowns suggest a role of eve in early cell paralog groups besides Wnt3 could be discov- division through early segmentation in ered. In the leech Helobdella robusta, only nine Helobdella. However, no pair rule pattern is paralog groups are present, with additionally found for this gene in the leech. Wnta , Wnt8, and Wnt9 missing. Most Wnt genes The zinc fi nger transcription factor hunchback in P. dumerilii are expressed in ectodermal seg- plays the role of a gap gene in Drosophila , which mental stripes and/or in the area around the defi nes expression domains of pair rule and Hox pygidium (Janssen et al. 2010 ). Expression anal- genes (see Vol. 5, Chapter 1). Moreover, hunch- yses in H. robusta and C. teleta led to comparable back is involved in mesoderm development and results (Cho et al. 2010 ). Due to similarities with neurogenesis. In Platynereis dumerilii , hunch- arthropods, a role of Wnt genes in segment for- back expression is detected in mesodermal cells mation in both annelids and arthropods is sug- belonging to the posterior growth zone of juve- gested by some authors (Janssen et al. 2010 ). nile worms. Additionally, expression in the pre- In summary, the candidate gene approach led cursors of the somatic segmented mesoderm, to the discovery of many similarities as well as formed during larval development, could also be differences between annelids and arthropods in confi rmed, a striking similarity with arthropods gene expression patterns during the formation of (Kerner et al. 2006). However, an expression of segments. The expression of some genes at seg- hunchback could not be detected in segmental mental boundaries in Platynereis dumerilii precursor cells of the posterior growth zone in shows a remarkable similarity to arthropods. Capitella and Helobdella , and a role in the pat- However, for other annelid taxa investigated for terning of the anterior-posterior axis was rejected these candidate genes (as, e.g., for engrailed or 218 C. Bleidorn et al.

hunchback), the picture becomes less clear, and 2008 ; Simakov et al. 2013). In Capitella , assem- future studies covering segment formation in bled whole genome shotgun data found Hox more annelid taxa are clearly wanted. Moreover, genes distributed on three scaffolds, with one fewer similarities are found compared with scaffold containing the Post1 genes clearly sepa- arthropods when investigating pair rule genes. In rated from the others. In contrast, the leech the discussion of a putative common ancestry of Helobdella shows an extensive fragmentation of segmentation in annelids and arthropods, differ- the Hox cluster (Simakov et al. 2013 ). For ent authors come to different conclusions using Capitella , 11 Hox genes (lab , pb , Hox3 , Dfd , Scr , basically the same set of evidence (de Rosa et al. lox5 , Antp , lox4 , lox2 , Post2 , post1 ) correspond- 2005 ; Thamm and Seaver 2008). However, ing to 11 different paralog groups were detected, homology of genes expressed during segment and the presence of these genes are regarded as formation must not imply a homology of a seg- ancestral for lophotrochozoans in general mented body plan itself. At present, available (Fröbius et al. 2008 ; Simakov et al. 2013 ). developmental, paleontological, and phyloge- Interestingly, Helobdella also shows a derived netic evidence supports a convergent evolution pattern here, with the duplication of two paralog of segmentation in arthropods and annelids groups (fi ve copies of Scr and two copies of (Couso 2009; Chipman 2010). Given this Post2) and the loss of orthologs of pb and Post1 hypothesis, co-option of the same set of genes (Simakov et al. 2013 ). For many other annelids, into the process of segment formation leading to information about the Hox gene complement are a convergent pattern of gene expression can available through PCR and cloning studies; how- explain the similarities found between annelids ever, genomic organization and absence of genes and arthropods (Chipman 2010 ; Ferrier 2012 ). cannot be derived from this approach (Dick and Buss 1994 ; Snow and Buss 1994 ; Irvine et al. 1997 ; Cho et al. 2003 , 2006 ; Kulakova et al. Hox and ParaHox Genes 2007 ; Bleidorn et al. 2009 ). The expression of Hox genes during develop- Hox genes comprise a family of transcription ment has been only investigated for a few annelid factors bearing a DNA-binding homeodomain taxa, Capitella teleta , Chaetopterus variopeda- (Gellon and McGinnis 1998). Hox genes are usu- tus , Alitta (Nereis ) virens , Platynereis dumerilii , ally found as linked chromosomal clusters and Hirudo medicinalis, and two Helobdella species show spatial and temporal collinearity (Garcia- (Irvine and Martindale 2000 ; Peterson et al. Fernandez 2004 ). This means that genes from 2000 ; Kulakova et al. 2007 ; Fröbius et al. 2008 ; the 5′-end of the cluster are usually expressed Gharbaran and Aisemberg 2013). The most more anteriorly than the ones from the 3′-end. In inclusive study deals with C. teleta , where for the similar fashion we also see a temporarily earlier fi rst time spatial and temporal collinearity for onset of genes from the 5′-end compared to those Hox genes could be demonstrated for a lophotro- from the 3′-end. In bilaterian animals Hox genes chozoan taxon (Fröbius et al. 2008 ). Capitella are mainly involved in the patterning of body Hox genes, except for Post1 , are all expressed in regions; however, several examples of co-option ectodermal domains during larval development, into other areas of expression are described with a spatial correlation of anterior expression (Wagner et al. 2003 ). All these characteristics borders and location of genes in the Hox cluster. made this set of genes a prime target for evolu- Anterior class Hox genes (lab , pb , Hox3 ) are the tionary developmental biologists to understand fi rst genes expressed, occurring before the major transitions in animal body plan evolution appearance of segments. The expression of Dfd (Akam 1998 ). and Scr can be detected after appearance of the For annelids, the genomic organization of the fi rst segments, followed by the expression of Hox cluster is only fully described for Capitella lox5 , Antp , and lox4 . The expression of lox2 and teleta and Helobdella robusta (Fröbius et al. Post2 appears last. Interestingly, all Hox genes in 9 Annelida 219

C. teleta show their highest expression level at a expression of Post2 was detected in the pygidium unique stage during the course of development, of nereidids (Kulakova et al. 2007 ). Expression refl ecting the order of activation for each gene. A of Post1 , the gene which seems to be separated unique Hox gene expression boundary can be from the rest of the Hox cluster, could not be detected for all nine thoracic segments, and the detected in any investigated stages of C. teleta , posterior-most located Hox genes (lox2 and besides some signals in chaetal sacs (Fig. 9.12 ; Post2) are only expressed in the abdomen Fröbius et al. 2008 ). This result is congruent with (Fig. 9.12). Whereas no expression of Hox genes analyses of expression of this gene in nereidids could be detected in the pygidium of C. teleta , (Kulakova et al. 2007 ).

Fig. 9.12 Hox gene expression profi le in larvae and juve- weaker expression. Abbreviations: A1–4 abdominal seg- niles of Capitella teleta after Fröbius et al. ( 2008 ). Solid ments 1–4, Gz growth zone, Pe peristomium, Pr prosto- bars indicate strong expression; dashed bars indicated mium, Py pygidium, T1–T9 thoracic segments 1–9 220 C. Bleidorn et al.

A staggered expression of fi ve Hox genes gen- Genes Involved in Neurogenesis erally in line with spatial and temporal collinear- ity can also be found in Chaetopterus , even The development of the central nervous system though the genomic organization of the Hox clus- has been deeply studied for Platynereis dumeri- ter remains unknown in this species (Irvine and lii. Neural progenitor cells are located close to Martindale 2000 ). No strict temporal collinearity the ventral midline and express axin , a negative was found in expression studies for nereidid regulator of the Wnt/β-catenin pathway, which worms and Helobdella (Kourakis et al. 1997 ; controls the transition between these proliferat- Kulakova et al. 2007 ). However, all studies sug- ing cells and differentiating neurons (Demilly gest an involvement of Hox genes in body pat- et al. 2013). Wnt-controlled proliferation of neu- terning along the anterior-posterior axis, a ral progenitors is also well-documented for verte- function that seems to be the ancestral role of brates and arthropods, especially Drosophila Hox genes in bilaterian animals (Kulakova et al. (Bielen and Houart 2014 ). Using a candidate 2007 ; Butts et al. 2008 ). Notably, annelids show gene approach, genes with a conserved expres- a predominant expression of Hox genes in neuro- sion in developing vertebrate and arthropod genic structures such as the ganglia and the ven- brains were chosen as major targets in studies on tral nerve cord. This is especially obvious in annelids. The developing head of annelid larvae Chaetopterus and leeches (Shankland et al. 1991 ; is demarcated by the expression of six3 and otx Aisemberg and Macagno 1994 ; Wong et al. 1995 ; homeobox genes (Fig. 9.13 ), a patterning system Kourakis et al. 1997; Irvine and Martindale 2000 ; that might be universal to bilaterian animals Gharbaran and Aisemberg 2013 ). (Steinmetz et al. 2010 ). MicroRNAs are short The ParaHox cluster is a paralog of the Hox noncoding RNAs that posttranscriptionally regu- gene cluster, containing three genes ( Gsx , Xlox , late gene expression (Ambros 2004 ). The expres- and Cdx ) (Brooke et al. 1998 ). As for Hox genes, sion of several microRNAs is highly tissue temporal collinearity has been likewise demon- specifi c and conserved across animals strated for ParaHox genes in many instances. (Christodoulou et al. 2010 ). In P. dumerilii and However, the ParaHox cluster seems to be lost in Capitella teleta , the microRNAs mir - 7 , mir - 137 , several investigated ecdysozoan taxa which show and mir - 153 show a localized expression in dis- a breakup of the cluster and missing genes tinct neurosecretory brain tissue, a pattern which (Ferrier and Minguillon 2003 ). All three ParaHox was also found in zebra fi sh (Tessmar-Raible genes seem to be present in annelids (Ferrier and et al. 2007 ; Christodoulou et al. 2010 ). The Holland 2001 ; Fröbius and Seaver 2006 ; Park expression of the complementary pair mir - 9 and et al. 2006 ). The genomic organization of mir - 9 */mir - 131 is restricted to two sets of differ- ParaHox genes has been studied in detail for entiated neurons in the developing annelid brain, Platynereis dumerilii (Hui et al. 2009). In this with the most apical cells located at the base of species, a head-to-head location of Gsx and Xlox the antennae (Christodoulou et al. 2010 ). The could be demonstrated, with Cdx located in a three transcription factors rx , otp , and nk2.1 are separate position on the same chromosome. all expressed in the developing forebrain of P. Expression analyses of these genes in Alitta dumerilii. All cells expressing these genes, as ( Nereis ) virens suggest a role in anterior- posterior well as mir - 7, are vasotocinergic extraocular pho- patterning of the digestive system and in the toreceptors. The expression pattern matches specifi cation of neuroectodermal cell domains those known from the same cell type in zebra fi sh (Kulakova et al. 2008 ). Especially Gsx seems to (Tessmar-Raible et al. 2007 ). Gene networks be involved in the development of the brain in all controlling the pattern along the anterior- posterior investigated annelids (P. dumerilii , A . (N .) virens , axis of the central nervous system are conserved C. teleta), a function which is regarded as ances- across bilaterians, and the involved genes are tral for these genes for bilaterians in general mainly Hox genes (Ferrier 2012 ). The patterning (Fröbius and Seaver 2006 ; Kulakova et al. 2008 ). of the dorso-ventral axis in P. dumerilii is 9 Annelida 221

Pro- stomium Head Larval eye

Peri- stomium Six3

otx

Meta- gbx stomium

Trunk hox1

hox4 Pygidium

lox5

Fig. 9.13 Expression of six3 , otx , gbx , and Hox genes in prostomium and the peristomium, from which the cere- neuroectodermal regions of Platynereis dumerilii larvae. bral ganglia and eyes develop. Dark gray region marks the The six3 and otx expressing regions cover the developing mouth (From Steinmetz et al. (2010 ) )

controlled by a gene network including nk2.2 , innervate the male penis (Gharbaran and nk6 , Pax2 / 5 / 8 , Pax6 , Pax3 / 7 , dlx , msx , gsx , sim , Aisemberg 2013 ). For the same species, expres- and dbx (Denes et al. 2007 ; Ferrier 2012 ). A sim- sion of the axon migration guiding protein netrin ilar patterning of the neuroepithelium is obvious is shown to be involved in forming intergangli- in vertebrates (Denes et al. 2007 ). onic neuronal tracts and in defi ning ventrodorsal Besides the detailed investigations summa- boundaries of peripheral innervation (Gan et al. rized for Platynereis dumerilii , some gene 1999 ). Another protein family investigated in H. expression studies dealing with the development medicinalis is the innexins, where several cloned of the nervous system in leeches and Capitella members show a restricted expression in neurons teleta are published. As shown for P. dumerilii , (Dykes and Macagno 2006 ). In C. teleta , Delta otx shows a largely head-specifi c expression in and Notch expression was detected during brain Helobdella (Bruce and Shankland 1998 ). The development in larvae, as well as in the forming expression of Lox10 , a putative nk2.1 ortholog, ganglia of the ventral nerve cord of juveniles, was detected in the developing brain of suggesting a role of Notch signaling in neurogen- Helobdella, congruent with the results for P. esis (Thamm and Seaver 2008 ). dumerilii (Nardelli-Haefl iger and Shankland In summary, development of the central ner- 1993 ). In Hirudo medicinalis, the central class vous system and expression patterns of genes Hox gene Lox1 controls the differentiation of the localized in the brain show strong similarities so-called “rostral penile evertor neurons” that between vertebrates and annelids. Based on 222 C. Bleidorn et al. these results (and further studies involving ily are exclusively expressed in the developing other taxa), the presence of a centralized ner- midgut, with a prominent expression of gataB1 at vous system in the last common ancestor of its boundaries. In contrast, FoxA expression can protostomes and deuterostomes seems plausi- be detected surrounding the blastopore during ble for several authors (Arendt et al. 2008 ; development as well as in the foregut and hindgut Holland et al. 2013 ). during organogenesis. Partly similar expression patterns are reported for Themiste and Chaetopterus and might refl ect the differences in Genes Acting in the Development the gut architecture of these species with differ- of the Digestive Tract ent feeding mechanisms (Boyle and Seaver 2010 ). Moreover, expression of the ParaHox The gut of annelids consists of a foregut (stomo- gene Cdx is also reported for anterior and poste- deum) and a hindgut (proctodeum), both origi- rior regions of the gut in C. teleta (Fröbius and nating from ectoderm, as well as of the midgut Seaver 2006 ). In nereidids, the ParaHox gene which is of endodermal origin. All three parts Gsx is expressed during the formation of the fore- can usually be subdivided into different func- gut and the midgut. The expression of the tional regions (Tzetlin and Purschke 2005 ). ParaHox gene Xlox has been detected in all inves- Several genes involved in bilaterian foregut and tigated annelids, including Helobdella , H. medic- hindgut patterning have been investigated for inalis , C. teleta, and nereidids (Wysocka-Diller Platynereis dumerilii (Arendt et al. 2001 ). As et al. 1995 ; de Rosa et al. 2005 ; Fröbius and such, the T-box transcription factor brachyury is Seaver 2006 ; Kulakova et al. 2008 ; Hui et al. expressed in the ventral part of the developing 2009 ). foregut as well as in the hindgut of late trocho- Bilaterian animals are divided into deutero- phore larvae, resembling the pattern known from stomes, ecdysozoans, and lophotrochozoans larvae of basal branching deuterostomes. The (Edgecombe et al. 2011 ). Whereas research on homeobox gene goosecoid is fi rst expressed in a several well-established model organisms in the small number of cells at the anterior blastopore former two groups (e.g., Drosophila , margin which develops into the foregut. Caenorhabditis , Danio , Mus) has provided Expression can be additionally detected in adja- detailed insights into molecular mechanisms of cent cells which will contribute to the develop- the development, lophotrochozoans have tradi- ment of the foregut nervous system. As the tionally been chronically understudied in this expression patterns of the investigated genes regard (Tessmar-Raible and Arendt 2003 ). The seem to be conserved in protostomes and deu- rise of Platynereis dumerilii (and other annelids terostomes, a single origin of the tripartite bilat- like Capitella and Helobdella ) as EvoDevo mod- erian through gut has been hypothesized (Arendt els has provided major insights into the evolution et al. 2001 ). This idea has been later challenged of the nervous system and segment formation in based on expression studies of the same set of annelids, a key lophotrochozoan phylum (see genes in acoels, which are lacking a through gut above). Interestingly, the genomic architecture of (Hejnol and Martindale 2008 ). Platynereis seems to be little derived from a Genes involved in the patterning of ectoder- hypothetical bilaterian ground pattern, enabling mal and endodermal parts of the gut have been many insights into comparative developmental studied for Capitella teleta , Chaetopterus vari- genomics (Raible et al. 2005 ; Ferrier 2012 ). opedatus , and the sipunculid Themiste lagenifor- Future studies focusing on additional annelid lin- mis . In C. teleta , the transcription factor FoxA eages, such as the basal branching oweniids or and genes of the GATA family are expressed the non-segmented sipunculans, will certainly across the entire developing gut (Boyle and improve our understanding of the evolution of Seaver 2008 ). Different genes of the GATA fam- bilateria in general. 9 Annelida 223

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Zootaxa 3147:1–237 Dev Biol 235:476–488 Phoronida 1 0 Scott Santagata

Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger.

S. Santagata Biology Department, Long Island University-Post , Greenvale , NY 11548 , USA e-mail: [email protected]

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 231 DOI 10.1007/978-3-7091-1871-9_10, © Springer-Verlag Wien 2015 232 S. Santagata

INTRODUCTION or three true coelomic compartments, this distinction does not clarify their phylogenetic Phoronids are epibenthic (or infaunal) tubiculous position relative to other lophophore-bearing marine invertebrates closely related to brachio- animals (ectoprocts and brachiopods) or to pods (and perhaps ectoprocts; see Nesnidal et al. annelids, nemerteans, and mollusks. (2013 )) that have oval, U-shaped, or spiraling Similar to ectoprocts, the adult phoronid gut rings of ciliated tentacles called the lophophore is U-shaped and the anus is positioned outside of used for feeding and respiration (Temereva and the tentacles. Unlike ectoprocts, each of the ten- Malakhov 2009a). Although phoronids can dom- tacles has a blind capillary with nucleated red inate the density and coverage of some benthic blood cells containing a form of hemoglobin marine habitats (Larson and Stachowicz 2009 ), (Garlick et al. 1979). These capillaries are con- very little is known about their ecological role in nected to lophophoral ring vessels that are fed such habitats. The majority of taxonomic studies and drained by the efferent and afferent blood of phoronids have been conducted by Emig vessel loop traversing the trunk region. The adult ( 1974 ). Although at least 23 species have been trunk is divided into a more posterior ampullary described by various authors indicative of wide region and a more anterior and tapered muscular morphological diversity in adult forms, the region that contains some diagonal muscle fi bers majority of phoronid morphotypes have been in the body wall (Chernyshev and Temereva synonymized under 11 cosmopolitan species and 2010 ). The main body wall musculature of the two genera, Phoronis (Wright 1856 ) and trunk epithelium consists of a layer of circular Phoronopsis (Gilchrist 1907 ). muscles underlying numerous feathery or bush- Adult phoronid bodies have generally been like longitudinal muscles (Herrmann 1997 ). considered to be tripartite, divided by transverse The number and distribution of the latter mus- septa into the epistome, tentacle crown, and trunk cles in the four mesenteric divisions of the trunk regions. Each of these body regions was origi- have been used as species-level morphological nally described as having a true coelomic cavity characters (Emig 1974 ); however, some “cosmo- lined by mesoderm and were interpreted as the politan” species have wide muscle formula protocoel (epistome), mesocoel (lophophore), ranges and may actually be cryptic species. In at and metacoel (trunk), linking this arrangement to least one species, Phoronis pallida , distinct mus- the tripartite coeloms found in echinoderms cular cinctures separate regions of the trunk (Masterman 1898 ). The epithelial lining of the (Santagata 2002 ), which may be a derived adap- epistome in two species of Phoronis was found tive feature linked to this species living as a instead to be myoepithelial cells lacking adherent commensal in the burrows of (at least) two junctions surrounding a gel-like extracellular thalassinid shrimps, Upogebia pugettensis matrix, thus not exhibiting the features of a true (Thompson 1972; Santagata 2004a ) and epithelial layer (Bartolomaeus 2001 ; Gruhl et al. Upogebia major (Kinoshita 2002 ). 2005). Further complicating these fi ndings are One main feature of the adult nervous system ultrastructural observations of the epistome lin- is a group of basiepidermal neuronal cell bodies ing of Phoronopsis harmeri, which does contain concentrated between the mouth and the anus adherent junctions (Temereva and Malakhov (Fernández et al. 1996 ). Although this structure 2011 ). This cavity can be derived from the proto- has often been called the dorsal ganglion (Silén coel formed in Phoronopsis harmeri larvae but 1954a ) or a dorsal neural plexus (Temereva and may collapse and break down during develop- Malakhov 2009b ), some aspects of its post- ment among various other species (Zimmer metamorphic development are not consistent 1978 ). Alternatively, this larval cavity may be with a completely dorsal origin (Santagata 2002 ). lost during metamorphosis and reform during Regardless of the differences between these post-metamorphic growth (Santagata 2002 ). structural interpretations, this anterior concentra- Regardless of whether adult phoronids have two tion of ciliated neuronal cells appears to be the 10 Phoronida 233 adult “brain” and is connected to a collar nerve genus-level relationships among phoronids ring with unciliated neuronal cells along its based on either morphological or molecular length at the base of the tentacles (Temereva characters are moderately congruent (Santagata and Malakhov 2009b ). Sporadically distributed and Cohen 2009 ) and support Phoronis ovalis as neuronal cells and fi bers are found throughout a divergent lineage, a Phoronopsis clade and a the surface of the trunk epithelium. A subset of subclade of Phoronis spp. It should be noted these cells and fi bers are serotonergic (Santagata that the molecular phylogenetic data discussed 2002 ), but the most centralized neuronal struc- here are comprised largely of ribosomal and ture is the giant nerve fi ber embedded in the ante- mitochondrial markers, and a more recent phy- rior portion of the trunk epithelium (Temereva logenetic analysis of phoronid species based on and Malakhov 2009b ). Among various phoronid these genes plus additional nuclear and mito- species, the giant nerve fi ber(s) takes on different chondrial genes resolves previously incongruent morphologies with respect to their number, posi- aspects of the morphology- and molecular- tion, and where the posterior limit of these fi bers based evolutionary inferences (Santagata, can be found (Emig 1974 ). 2014 ). The soft tissues of phoronids do not make for particularly good fossils, but their chitinous and sandy tubes have been linked to some trace fossil EARLY DEVELOPMENT types. A Phoronis ovalis-like boring pattern may have produced the ichnofossil Talpina found in Gonads develop from the peritoneal lining the Devonian period (Thomas 1911 ). However, it covering the capillaries of the efferent blood is likely that the crown phoronid lineage is much vessel on the surface of the stomach (Ikeda 1903 ; older and may be linked to vermiform fi lter- Rattenbury 1953 ; Zimmer 1991 ). Adults are feeding forms of tommotiids from the lower either gonochoristic or hermaphroditic, with Cambrian that had unfused organophosphatic some species having a small temporal bias toward sclerites such as Eccentrotheca (Skovsted et al. protandry (Zimmer 1991 ). Zimmer ( 1991 ) recog- 2008, 2011 ). Molecular estimates of the phoronid- nized three reproductive types among phoronid brachiopod root are even older, reaching as far species that are largely based on the sex of the back as the Ediacaran (Sperling et al. 2011 ). adult, mature egg diameter, and how embryos are Debate still exists as to whether phoronids brooded (if present). Group one is comprised should be considered as an ancestral sister taxon almost entirely of gonochoristic species (except to all brachiopods (Sperling et al. 2011 ; although Phoronis pallida ) that freely spawn 60 μm ova see Thomson et al. 2014 ) or instead as a sub- and includes members of both genera. Members taxon within the brachiopods as a whole (Cohen of group two are for the most part species that and Weydmann 2005; Santagata and Cohen produce larger mature eggs (≥ approximately 2009 ; Cohen 2013 ). Phoronids are clearly 100 μm) and brood embryos to an early larval or closely related to brachiopods and share evolu- competent larval stage on specialized nidamental tionary affi nities with spiralian protostomes glands at the base of the lophophore. Patterns such as nemerteans, annelids, and mollusks, but within group two are somewhat muddled due to their phylogenetic position relative to ectoprocts little or incomplete knowledge regarding the has been unresolved (Halanych et al. 1995 ; reproductive properties of species such as Dunn et al. 2008; Hejnol et al. 2009 ; Hausdorf Phoronopsis albomaculata and other disputed et al. 2010 ; Mallatt et al. 2012 ). Interestingly, a species morphotypes ( Phoronis capensis and recent phylogenomic study supports the mono- Phoronis bhadurii). Group three has only one phyly of all three lophophore-bearing phyla in member, Phoronis ovalis , whose sex type remains which phoronids are closely related to brachio- unconfi rmed, produces approximately 125 μm pods and considered an ancestral sister taxon to eggs, and broods embryos in the parental tube ectoprocts (Nesnidal et al. 2013 ). Species- and (Harmer 1917 ; Silén 1954b ). 234 S. Santagata

In species where it has been studied in any exact mechanism is not known) lyses through the detail, fertilization is internal. Primary oocytes epithelial tissue into the collar coelom (Zimmer are arrested at metaphase and released from the 1991 ). From there, the activated sperm swims ovary to fuse with activated V-shaped sperm down to the septum that divides the collar and (Zimmer 1964 , 1991 ; Reunov and Klepal 2004 ). trunk coelom and lyses through that wall. Sperm Male pronuclei have been found in the coelomic must then swim down the trunk to its base to oocytes of Phoronopsis harmeri (Rattenbury fertilize oocytes from the ovary. As complicated 1953 ) and Phoronis ijimai (currently = vancouve- as this latter fertilization scenario may be, rensis; Zimmer 1991 ; Hirose et al. 2014 ). There yet another variant of this method may exist, as is one largely undocumented report of external spermatophores have also been observed being fertilization in Phoronis muelleri (Herrmann ingested by nearby adults. 1986), but this observation requires more data. Once expelled via the metanephridial ducts How active internal sperm reach the ovary is one into the surrounding seawater, the fertilized of the more intriguing aspects of phoronid primary oocytes become activated and complete biology. Even in species that are simultaneous meiosis forming two to three polar bodies (the hermaphrodites, self-fertilization has not been fi rst polar body often divides). At this point, reported, as mature sperm released from the tes- cleavage begins, and the zygote divides equally tes are not yet activated (Zimmer 1964 ; Reunov (or approximately equally) and totally along the and Klepal 2004 ). Small masses of mature sperm animal-vegetal axis (Fig. 10.1B, C ). Second released from the testes make their way through cleavage repeats this process, but with a cleavage the trunk coelom until gathered up by the ciliary angle perpendicular to the fi rst cleavage, result- currents of the paired metanephridial funnels. ing in a four-cell embryo (Fig. 10.1D ). Although some authors have observed sperm Blastomeres isolated at the latter stage and two- directly released from the metanephridial ducts cell stages are capable of forming the whole into the surrounding seawater (Rattenbury 1953 ; larva; accordingly, early development is regula- Silén 1954b ), more often released sperm are tive (Zimmer 1964 ). enclosed by the paired lophophoral organs at the Whether subsequent cleavage stages of vari- base of the lophophore. Here, sperm are pack- ous phoronid species exhibit aspects of radial aged into spherical, bean- or club-shaped sper- cleavage or spiral cleavage has been debated matophores, some of which have elaborate since the fi rst embryological observations were sail-like structures (Zimmer 1967 , 1991 ). gathered. Radial (or biradial) cleavage patterns Completed spermatophores drift away and the have been found in several species (Masterman enclosed sperm remain inactive until reaching 1900; Ikeda 1901 ), but spiral-like cleavage pat- another adult conspecifi c individual. Perhaps the terns have also been observed (Foettinger 1882 ; most common pathway for sperm to reach the Brooks and Cowles 1905), and further, both pat- ovary of another individual is via the metane- terns have been observed in the same species phridial ducts (Brooks and Cowles 1905 ; (Herrmann 1986 ). A more typical, spiral cleav- Rattenbury 1953 ; Zimmer 1967). Once contact- age pattern was described for Phoronopsis viridis ing the duct, sperm in the head of the spermato- (junior synonym of P. harmeri) by Rattenbury phore enter the duct and continue down the (1954 ), but this was interpreted by Zimmer funnel, to the base of the trunk, to the ovary. (1964 ) as an artifact introduced by compaction of Another less common route has been observed in the blastomeres by a tightly fi tting vitelline enve- Phoronopsis viridis (currently considered to be a lope. It should be noted that all of the embryos junior synonym of Phoronopsis harmeri ; see having spiral-like cleavage patterns (or both spi- Emig 1974 ; Santagata and Cohen 2009 ), in which ral and radial) came from species that produce the spherical portion of the spermatophore con- smaller (60 μm) eggs, but other species that pro- tacts the tip of a tentacle where sperm using duce larger (100 μm) eggs such as Phoronis iji- enzymes from their acrosomes (presumably, the mai (= vancouverensis ) more consistently have 10 Phoronida 235

A B C D

E F G H

I J K

L M N

Fig. 10.1 Early cleavage stages of Phoronis pallida counterclockwise twist (denoted by the arrows) to the (A–H) and Phoronis vancouverensis (junior synonym of 16-cell stage. Three polar bodies are found in the center of Phoronis ijimai; I–N). ( A ) Adult P. pallida collected from the image. ( G ) Animal pole view of a 16-cell embryo False Bay, WA, USA; note the characteristically bent dividing with a clockwise twist (denoted by the arrows ) to sandy tube (S ) and delicate U-shaped lophophore ( L ) (© the 32-cell stage. ( H) Two 32-cell embryos. ( I ) Ventral Scott Santagata 2015. All Rights Reserved). ( B ) Zygotes view of an adult Phoronis vancouverensis collected from (approx. 60 μm) with polar bodies ( PB) and a two-cell Westcott Bay, WA, USA. This species occurs in muddy stage. ( C) Confocal z-projection of a two-cell stage clumps with other conspecifi c individuals and broods stained for nucleic acids (blue ) and fi brous actin (yellow ). various early developmental stages (early cleavage to The embryo is undergoing division to the four-cell stage. four-tentacle actinotrochs) in paired lophophoral masses. ( D ) A four-cell (4C ) and an eight-cell (8C ) embryo. (E ) A A fertilized primary oocyte (approx. 100 μm; arrow ) is radially arranged eight-cell stage ( 8C ), an eight-cell stage present in one of the metanephridial funnels at midlevel in undergoing a spiral-like division to the 16-cell stage ( 8– the body. ( J) Two-cell stage. (K ) Four-cell embryo. (L ) 16C), a 16-cell stage with an open and radial blastomere Eight-cell embryo with radial arrangement of blasto- arrangement ( 16C ), and a 32-cell stage showing the apical meres. ( M) 16-cell stage with an open radial arrangement quartet of cells (32C ). All embryonic images are from dif- of blastomeres. ( N) 32-cell embryo. Scale bars in A and I ferent embryos placed into one composite fi gure. ( F ) equal 2 mm, scale bars in B and J – N equal 50 μm, scale Animal pole view of an eight-cell embryo dividing with a bar in C equals 10 μm, and scale bars in D – H equal 25 μm 236 S. Santagata radially cleaving embryos (Zimmer 1964 ; Wu adult that will begin cleaving once exposed to et al. 1980 ; Malakhov and Temereva 2000 ; seawater, there can be developmental differences Freeman and Martindale 2002 ), although irregu- between embryological cultures started from lar early cleavage stages are also reported. oocytes gathered this way as compared with nat- Cleavage variation also begins at different early urally spawned oocytes. What changes may cleavage stages among species; some cleave vari- occur to the fertilized oocytes while housed in the ably at the eight-cell stage (Brooks and Cowles nephridial funnels or what signals are received 1905 ; Herrmann 1986 ) and others (more com- when the oocytes are extruded through the monly) at the 16-cell stage (Temereva and nephridial ducts are not known, but naturally Malakhov 2007 ). More recently, early cleavage spawned embryos do develop more synchro- was investigated in Phoronis muelleri using 4D nously. For these reasons, it is best to gather microscopy techniques (Pennerstorfer and fertilized primary oocytes that are packed into the Scholtz 2012 ), which showed that oblique rather nephridial funnels rather than those isolated from than perpendicular cell divisions occurred at third the posterior part of the trunk and ovary and subsequent cleavages of most of the embryos (Santagata, pers. obs.). Early cleavage stages in the study. Interestingly, cell divisions at third shown in Fig. 10.1 were imaged with these con- cleavage and beyond also exhibited alternating siderations in mind. Four-cell stages of Phoronis dextral-sinistral twists, a feature typical of spira- pallida typically have a gap between the pairs of lian embryos (Hejnol 2010 ). Through these data sister cells and do not make cell contacts or cross- coupled with their thorough review of cleavage furrows as found in Phoronis muelleri (approx. variation in phoronid embryos, Pennerstorfer and 25 % of embryos; Pennerstorfer and Scholtz Scholtz (2012 ) make a strong case for the spira- 2012). Third cleavage of Phoronis pallida lian affi nities of some aspects of phoronid devel- embryos produces some eight-cell stages that opment. That said, further detailed accounts of have a radial arrangement of blastomeres and phoronid embryogenesis are required to discern others that have animal cells slightly offset (with the nature, variability, and signifi cance of early a clockwise twist) from those in the vegetal half cleavage patterns, especially in those species that of the embryo (Fig. 10.1 ). Fourth cleavage is also produce larger eggs and tend to have more con- variable among embryos; some 16-cell stages sistently radially cleaving embryos. For these have the characteristic open radial arrangement reasons, in the next section, the cleavage patterns of blastomeres (Fig. 10.1E ), while other embryos of Phoronis pallida and Phoronis vancouverensis have four tiers of cells offset from one another, (a junior synonym of P. ijimai) are compared arranged by oblique divisions with a using confocal and time-lapse microscopy. counterclockwise twist (Fig. 10.1E, F ). Embryos Numerous exogenous factors can affect the at fi fth cleavage continue this pattern of alternat- arrangement of blastomeres and the way early ing the directional twist of cell divisions embryos cleave. Obviously, temperature is a cru- (Fig. 10.1G ) and 32-cell stages have centrally cial variable and care must be taken to ensure that positioned quartets of cells at the animal and veg- embryos are not heated beyond their optimal etal poles (Fig. 10.1H ). ranges (see data in Staver and Strathmann 2002 ). The larger embryos of Phoronis vancouveren- Another less acknowledged variable is salinity – sis typically have a radial arrangement of blasto- it is known to affect fertilization success (Allen meres (Fig. 10.1I–N). Although irregular early and Pechenik 2010 ), but its nonlethal effects on cleavage stages have been observed (Zimmer early development (which may cause embryonic 1964 ; Malakhov and Temereva 2000), none of spaces to swell or shrink) are not well under- these patterns exhibit spiral-like divisions. At the stood. One issue particular to phoronids is the four-cell stage, transient cell contacts between typical way in which embryological cultures are non-sister blastomeres do occur, creating a cross- started. Even though numerous fertilized oocytes furrow similar to what is described for Phoronis may be gathered from the trunk coelom of an muelleri (Pennerstorfer and Scholtz 2012 ). 10 Phoronida 237

Eight- cell and 16-cell stages have the radial cell space and aggregate at the anterior portion of arrangements discussed previously, and 32-cell the late gastrula stage (see Fig. 10.2B ). These stages also have centrally located quartets of cells cells will form a small cavity underneath the at the animal and vegetal poles with a cross- apical tuft region of the late gastrula. This cavity furrow similar to what is observed in P. pallida has been interpreted as the fi rst coelomic cav- (Fig. 10.1 ). All of these radial-like and spiral-like ity (“protocoel”), but as discussed previously, cleavage patterns in phoronids support their spi- whether these cells form a true epithelium in ralian affi nities, and perhaps also indicate inde- all or in just select species is debated (Zimmer pendent switches to radial cleavage patterns in 1978 ; Bartolomaeus 2001 ; Temereva and particular species lineages, or may point toward Malakhov 2006 ). At this asymmetric late gas- radial-like cleavage being plesiomorphic for both trula stage, the gut is a blind sac. Cell labeling deuterostomes and protostomes. Although pho- and other embryological experiments show that ronid spiral-like cleavage patterns are clearly dif- much of the larval mesoderm is derived from ferent from the mosaic cell lineages of most ectodermal cells, especially where the ecto- mollusks (see Chapter 7 ; Hejnol et al. 2007 ) and derm is in contact with the endoderm, and the annelids (Chapter 9 ; Meyer et al. 2010 ), genetic remaining mesoderm is derived from the endo- mechanisms that underlie patterns of chirality in derm (Freeman and Martindale 2002 ; Santagata some gastropods may shed light on variable pho- 2004b; Temereva and Malakhov 2007). The fate ronid cleavage patterns in that the sinistral mapping studies of Freeman also found that the embryos of Lymnaea stagnalis are initially radi- development of phoronids was most similar to ally arranged before undergoing a relatively late that of rhynchonelliform brachiopods (Freeman sinistral twist (see Chapter 7 ; Shibazaki et al. 1991 , 2003) in that similar morphogenetic 2004 ). Furthermore, dextral and sinistral embryos movements reposition the anterior ectoderm. in this species are genetically determined at a Similar animal-vegetal inductive signals con- single locus (dextral is dominant), and so at least tribute to the formation of the anterior-posterior for this species of mollusk, allelic differences can axis in phoronids, rhynchonellid brachiopods, account for positional shifts in the spindle appa- and discinid brachiopods, but one difference ratus (Chapter 7 ; Kuroda et al. 2009 ). found in phoronids is that the animal half of Subsequent cell divisions yield either a thin- the embryo is not specifi ed until late in gastru- or thick-walled round blastula depending on the lation (Freeman 1991 , 2003). During the earli- size of the egg with a spacious or more compact est phase of larval development, the ectoderm blastocoelic space, respectively (Zimmer 1980 ; anterior to the mouth grows quickly, forming Herrmann 1986 ; Santagata 2004b ; Temereva the preoral hood, which has an upper exumbrel- and Malakhov 2007 ). Gastrulation begins lar epithelium with a developing apical organ with the fl attening of the vegetal region of the (Fig. 10.2D ) and a lower subumbrellar epithelial blastula forming a vegetal plate that is then layer (Fig. 10.2D ). Mesenchymal cells now line internalized. In species with smaller eggs, the the subumbrellar portion of the hood and the blastocoelic cavity remains spacious during the ectodermal portion of the trunk. These cells dif- early stages of gastrulation, but not in species ferentiate into muscle cells that form the initial with larger eggs (Zimmer 1991 ; Malakhov and body wall muscle fi bers of the larva (Santagata Temereva 2000 ; Santagata 2004b ). The blasto- 2004b ; Temereva and Tsitrin 2013 ). Where the pore is initially rounded but becomes slit-like as blind-ended gut meets the posterior ectoderm, the embryo elongates. The slit-like blastopore the anus and intestine form from an ectoder- is progressively closed from the posterior end, mal invagination (Freeman and Martindale leaving only a small anterior remnant to form 2002). As the early larval stage grows, the hood the mouth region. During this process, mesen- enlarges signifi cantly and drapes over the trunk chymal cells from the epithelial regions near region (Fig. 10.2E ). At this stage, a diagonal the developing oral region enter the blastocoelic band of epidermis on the future collar increases 238 S. Santagata

A B C D E

F G H

I J K L

Fig. 10.2 Gastrulation and larval development of various has grown rapidly and drapes over the collar. Clear demar- species of phoronids. ( A ) Light micrograph of a blastula cations exist between the gut ( G) and intestine ( I ), which stage of Phoronis pallida (adults collected from False are separated by a sphincter valve. ( F) Scanning electron Bay, WA, USA) that has a spacious blastocoel ( BL ). (B ) micrograph of a tentacle ridge ( TR) stage of Phoronis van- Light micrograph of a late gastrula stage of Phoronis couverensis (this morphotype is a junior synonym under architecta (adults collected from Alligator Bay, FL, Phoronis ijimai). The brooding adults were collected from USA). This morphotype is currently a junior synonym Westcott Bay, WA, USA. Early stages are held in the under Phoronis psammophila but is kept separate here for brood mass by mucous cords attached to the developing reasons discussed in Santagata and Cohen ( 2009 ). The apical organ (AO ) of the hood (H ). ( G ) Scanning electron preoral hood is precociously developed and the “proto- micrograph of an early tentacle bud (TB ) stage of Phoronis coel” (PC, whose lining’s structural composition as a true vancouverensis in which the simple ciliation on the sur- epithelial tissue has been questioned) is evident. Similar face of the larva becomes denser and the trunk lengthens. to many other larval forms, the developing apical organ ( H ) As the fi rst pair of tentacles becomes better defi ned, has centrally located cells with long cilia (apical tuft, AT ). their ciliation transforms into lateral ( LC ) and frontal cili- The blind-ended gut ( G) is also present at this stage. (C ) ary bands. The posterior part of the trunk also develops a Late gastrula stage shaded for embryological germ tissues telotrochal ( TH) band of cilia (© Scott Santagata 2015. based on cell-marking experiments in Phoronis vancouve- All Rights Reserved). ( I) Light micrograph of a two-ten- rensis (Freeman 1991 ). The animal pole of the egg is tacle stage of Phoronis vancouverensis with mouth ( M ) marked by the position of two polar bodies ( black ), the and anus ( A ). ( J) Light micrograph of a Phoronis pallida anterior ectoderm is shaded blue, the anterior mesoderm larva with eight tentacles. The trunk sac (TS ) grows rela- is shaded red, and the gut endoderm is shaded green . (D ) tively fast at this stage in preparation for metamorphosis. Light micrograph of an early tentacle ridge stage of ( K ) Light micrograph of a competent ten-tentacle stage of Phoronis pallida in which amoeboid mesenchymal cells Phoronis pallida that has a single, red blood corpuscle (MC ) on the subumbrellar portion of the hood and in the mass ( BC) and a muscular trunk sac (TS ) (© Scott trunk blastocoelic cavity will form the fi rst muscle fi bers Santagata 2015. All Rights Reserved). ( L ) Light micro- of the body wall (see Santagata ( 2004b )). The developing graph of a competent 18-tentacle stage of Phoronopsis apical organ (AO ) is more defi ned at this stage. The anus harmeri collected from the plankton near Friday Harbor and intestine are initially formed from an ectodermal Laboratories, WA, USA. Scale bars in A – E equal 25 μm, invagination. ( E ) Light micrograph of a late tentacle ridge scale bars in F – I equal 50 μm, scale bars in J and K equal (TR ) stage of Phoronis architecta in which the hood (H ) 100 μm, and scale bar in L equals 200 μm 10 Phoronida 239 in thickness and cell number, forming the cili- LATE DEVELOPMENT ated tentacle ridge (Fig. 10.2E, F ). Paired lateral projections develop at the ventral midpoint of Competent actinotroch anatomy varies mainly in the tentacle ridge, forming the fi rst two tentacle maximum size, number of blood corpuscle buds (Fig. 10.2G ). As these tentacle buds grow masses, maximum number of larval tentacles, into functional locomotory and feeding struc- and how the juvenile tentacles are differentiated tures, their ciliated epithelia differentiate into in the larva (see Fig. 10.3 ; Santagata and Zimmer lateral and frontal bands of cilia (Fig. 10.2H ). 2002 ). Only one species, Phoronis ovalis , pro- Lateral cilia generate downward currents that duces a non-feeding slug-like larva that lacks bring food particles to the frontal surface of tentacles. Once released from the parental tube, the tentacles, where specialized sensory cells Phoronis ovalis larvae swim in the plankton for (laterofrontal cells) likely detect them, causing about 4 days and then transition to a demersal localized ciliary arrests and/or reversals. These phase, swimming near the bottom for about changes in ciliary beat couple with a lifting of 3 days before undergoing metamorphosis (Silén the hood creating additional suction, thus lift- 1954b ; Zimmer 1991 ). Feeding actinotroch lar- ing the particle to the mouth to be ingested vae have a complex complement of striated mus- (Strathmann and Bone 1997). By the time the culature for lifting and closing the preoral hood; fi rst pair of tentacles is formed, the middle collar extending, fl icking, or lowering the tentacles; and region of the actinotroch larva is well defi ned, adjusting the angle of the teletrochal band of cilia dividing the body into three regions (hood, col- while swimming (Zimmer 1964 ; Santagata and lar, and trunk). Pairs of new tentacles will be Zimmer 2002 ; Santagata 2002 , 2004b ; Temereva added laterally and dorsally as the larva grows and Tsitrin 2013 ). Although many aspects of lar- and the trunk lengthens (Fig. 10.2J). How many val musculature are shared among several phoro- larval tentacles are formed varies among species nid species, the morphology of particular larval and larval sizes (Santagata and Zimmer 2002 ). muscles does differ, especially when considering Late in larval development, a midventral thick- the smaller and more compact actinotroch types ening in the trunk epidermis invaginates to form ( P. pallida , P. ijimai, and P. hippocrepia ) as an internal sac (Fig. 10.2J, K ). Typically called opposed to the larger and more elongate types the metasomal sac, it will be referred to here as ( Phoronopsis harmeri and Phoronis muelleri ; see the trunk sac to avoid unnecessary comparisons Santagata and Zimmer 2002 ; Temereva and to the anatomy of ambulacral deuterostomes. Tsitrin 2013). However, some reported differ- The trunk sac continues to elongate, displacing ences in larval musculature are the result of inter- the stomach and intestine dorsally. The trunk pretational differences among various authors. sac differentiates into what will be the trunk Both conserved and derived features also epithelium of the juvenile, replete with numer- comprise the larval nervous system among vari- ous circular and longitudinal muscles (Zimmer ous phoronid species. The greatest concentration 1964 ; Santagata 2002 ; Temereva and Tsitrin of neuronal cells in the larval nervous system is 2013 ). Other general features of the competent found in the apical organ, a U-shaped structure actinotroch larva are the compact mass(es) of containing at least four different neuronal cell red blood cells that form in the collar region types, all of which send axonal fi bers into a cen- (Fig. 10.2K ), the precociously developed juve- tral neuropil (Hay-Schmidt 1989 , 1990 ; Lacalli nile blood vessels, and variation in the differen- 1990 ; Santagata 2002 ; Temereva and Wanninger tiation of the juvenile tentacles (Santagata and 2012 ; Sonnleitner et al. 2013 ; Temereva and Zimmer 2002 ). Tsitrin 2014). Cellular domains within the 240 S. Santagata

A B C

D E F

G

HIJ

Fig. 10.3 Aspects of competent larval characteristics of the apical sense organ (AO ) and central neuropil (NP ) and metamorphosis among actinotroch morphotypes. ( A ) in a late-stage actinotroch larva labeled for serotonin Scanning electron micrograph ( SEM) of a competent ( green ). The three median hood nerves ( HN ) project from stage of an unidentifi ed actinotroch larva, “Actinotroch the apical organ to the margin of the hood. (F , G ) C,” collected from the plankton near the Port of Los Serotonin-like sensory cells in the frontal organs of two Angeles, CA, USA (Santagata and Zimmer 2002 ), show- different actinotroch morphotypes. ( H) Early stage of ing the hood ( H ), collar (C ), and trunk (T ) regions as well metamorphosis when the juvenile trunk sac (TS ) is everted as the heavily ciliated telotroch ( TH ). (B ) SEM of the and the blood corpuscle mass ( BC ) has not yet dissoci- juvenile tentacle rudiments ( AT ) attached to the proximal- ated. This morphotype was collected from the plankton of basal side of the larval tentacles ( LT ) of Actinotroch C. Tampa Bay, Florida, USA, and its identity is likely ( C ) Musculature of Actinotroch C stained with phalloidin Phoronis hippocrepia . (I ) First stage of metamorphosis in ( red ). (D ) Musculature (red ) and serotonergic nervous Phoronopsis harmeri (collected from the plankton outside system ( green ) of an actinotroch morphotype with similar Los Angeles Harbor, CA, USA), showing the numerous anatomical traits to that of Phoronis muelleri (but a sepa- serotonin-like cell bodies and fi bers (green ) in the larval rate species), showing the apical organ (AO ), frontal organ and juvenile nervous systems. ( J ) SEM of the juvenile ( SO ), and numerous serotonin-like cell bodies in the tissue lophophore of an unidentifi ed phoronid species. Scale of the juvenile trunk sac (TS ). Tentacle muscles, elevators bars in A , C , D , and I equal 200 μm, scale bar in B equals ( TE ), and depressors (TD ) underlie the frontal and abfron- 10 μm, and scale bars in F and H equal 100 μm tal sides of each larval tentacle. (E ) Magnifi ed apical view 10 Phoronida 241

apical organ include groups of cells that express for Phoronopsis harmeri (Temereva 2012 ), the serotonin (Hay-Schmidt 1990 ; Santagata 2002 ; cytoarchitecture of these neural fi bers has very Temereva and Wanninger 2012 ; Sonnleitner et al. little in common with the ventral nerve cords of 2013 ), catecholamines (Hay-Schmidt 1990 ; molluscan or annelid larvae in that phoronid lar- Santagata 2002 ), FMRFamide (Hay-Schmidt vae do not have centralized ventral nerve cords 1990 ; Sonnleitner et al. 2013 ), and a cardioactive with repeated neuronal cell bodies along them B-like peptide (Sonnleitner et al. 2013 ). (Nederbragt 2002 ; Nederbragt et al. 2002 ; Serotonin-like immunoreactive cells include Denes et al. 2007 ; Meyer and Seaver 2009 ). numerous fl ask-shaped cells that overlie other Overall, there are several structural and neuro- bipolar or multipolar neuronal cells (Hay- chemical aspects of the larval nervous system Schmidt 1990 ; Santagata 2002 ; Temereva and that vary among species (Santagata and Zimmer Wanninger 2012 ; Sonnleitner et al. 2013 ). 2002; Sonnleitner et al. 2013 ; Temereva and Neuronal cells and fi bers that have catecholamine- Tsitrin 2014 ). However, it should be noted that like immunoreactivity are situated at the periph- some of this variation in the larval nervous sys- ery of the apical organ (Hay-Schmidt 1990 ; tem may be due to unrecognized cryptic specia- Santagata 2002 ). Only a few to perhaps ten bipo- tion when comparing what are believed to be the lar neuronal cell bodies on dorsolateral sides of same larval types from geographically distant the apical organ have FMRFamide-like immuno- populations. One example can be found in reactivity, depending upon the species and larval Phoronopsis harmeri, a species whose type stage (Hay-Schmidt 1990 ; Temereva and locality is near Vancouver Island, Canada (Pixell Wanninger 2012 ; Sonnleitner et al. 2013 ). The 1912 ). Near its type locality, this species’ FMRFamide-like immunoreactive cells do not competent larval form typically has 18–20 ten- appear to be sensory, as they lack cilia and their tacles and four blood corpuscle masses fi ber projections go to the apical neuropil and the (Fig. 10.3; see also Zimmer 1964), but compe- tentacular neurite bundles, leading into the major tent Phoronopsis harmeri larvae collected from tentacle nerve rings. the plankton of Vostok Bay, Sea of Japan, are Beyond the apical organ, signifi cant larval competent at 24 tentacles (Temereva and nerves include the main hood nerve and two Wanninger 2012 ) and may have variable num- dorsolateral nerves that run down into the collar bers of blood corpuscle masses (Temereva and (mesosome) region of the larval body and merge Neretina 2013 ). For these reasons, care must be with the minor and major tentacle nerve rings. taken when assessing the phylogenetic context From these nerve rings, the larval tentacles (and and functional importance of characters in the sometimes the juvenile tentacle rudiments; phoronid larval nervous system (Santagata Santagata and Zimmer 2002 ) are innervated on 2011 ). frontal and abfrontal sides (Hay-Schmidt 1989 , Some of the more dramatic features that 1990 ; Lacalli 1990 ; Santagata 2002 ; Temereva develop in the larval nervous system near the and Wanninger 2012 ; Sonnleitner et al. 2013 ; time of metamorphic competence are the three Temereva and Tsitrin 2014 ). Basiepithelial median hood nerves that emanate from the apical fi bers also project from these nerve rings organ and merge into a secondary hood sense through the larval trunk epithelium and merge organ (also called the frontal organ) that has its with a telotrochal nerve ring at the posterior end own neuropil (Santagata and Zimmer 2002 ; of the larva (Hay-Schmidt 1990 ; Santagata and Santagata 2002 ; Temereva and Wanninger 2012 ; Zimmer 2002; Temereva and Wanninger 2012 ; Sonnleitner et al. 2013 ). Depending on the mus- Sonnleitner et al. 2013 ). Although a possible culature of the hood, the frontal organ may be vestigial ventral nerve cord has been reported cone-shaped and eversible (Santagata and 242 S. Santagata

Zimmer 2002 ) or be composed of a fi eld of sen- coelom and formation of the blind-ended capil- sory cells at the distal portion of the hood laries are similar to that of P. muelleri (Santagata (Santagata and Zimmer 2002 ; Temereva and 2002 ). In this species, the larval tentacle muscles Wanninger 2012 ). More commonly, the frontal (both elevators and depressors) are broken down organ includes several bipolar, serotonergic and new juvenile tentacle muscles are formed in sensory cells (Fig. 10.3F, G ; Santagata and a day (Santagata 2002 ). However, in Phoronopsis Zimmer 2002 ; Temereva and Wanninger 2012 ) or harmeri , only the tentacle depressors are broken another neurotransmitter type (Santagata 2002 ; down (Temereva and Tsitrin 2013 ), and this dif- Sonnleitner et al. 2013). Evolutionary modifi ca- ference may be linked with the fact that the ten- tions to the functions of the frontal organ’s neu- tacle capillaries are already formed and expanded rons may confer different degrees of settlement in the competent larva’s comparatively larger behavior specifi city (Santagata 2004a ). tentacles. Other muscles carried over to the Actinotroch larval metamorphosis is rapid and juvenile body in this species are found in the cataclysmic. Within a span of 15–30 min, a larva esophagus (Temereva and Tsitrin 2013 ). In all with a straight gut and radiating tentacles trans- phoronid species investigated thus far, the forms into a juvenile with an elongate vermiform muscular and neural tissues of the preoral trunk with a bulbous ampullar posterior end and hood undergo histolysis (Herrmann 1979 ; an oval or U-shaped array of tentacles. At its Bartolomaeus 2001 ; Santagata 2002 ; Temereva midpoint, the larval gut is drawn into the cavity and Tsitrin 2013 ). In contrast to some Phoronis of the trunk sac by the midventral mesentery spp., the hood coelom of Phoronopsis harmeri (Fig. 10.3H ). During this process, the cells of the has a true epithelial lining, at least a portion of blood corpuscle mass dissociate and stream into which may be carried over to form the coelomic the vessels of the juvenile circulatory system lining of the juvenile epistome (Temereva and (Zimmer 1991 ; Santagata 2002 ; Temereva and Malakhov 2006 , 2011 ). Tsitrin 2013 ). Depending on how the juvenile tentacles develop (Santagata and Zimmer 2002 ), there are differences in (1) the formation of the OPEN QUESTIONS lophophore blood vessels and (2) the larval tis- sues that undergo cell death. In species that have • Are similar axial patterning mechanisms a separate set of juvenile tentacles in the compe- present in phoronids, brachiopods, and per- tent larva (e.g., Phoronis muelleri), the larval ten- haps even ectoprocts? tacles are completely lost. During metamorphosis, • What are the expression profi les of key devel- the compacted lining of the collar coelom inside opmental genes such as Hox and ParaHox the juvenile tentacles expands (Bartolomaeus genes in phoronids? 2001 ), and post-metamorphosis, the blind-ended • Are the expression patterns for genes such as capillaries of the juvenile tentacles form in a day. Six3 / 6 , NK2.1 , Homeobrain , FoxQ2 , and Otp For those species that form the juvenile tentacles in embryos and larvae of rhynchonelliform from basal tentacle rudiments attached below the brachiopods (e.g., Terebratalia ) shared by the proximal ends of the larval tentacles (e.g., embryos and larvae of phoronids? Phoronis psammophila ), similar tissue modifi ca- • What is the evolutionary basis and importance tions occur, but the fate of the frontal portion of of both radial-like and spiral-like cleavage the larval tentacles overlying the juvenile rudi- patterns among species of phoronids? ments has not been studied in great cytological detail (Veillet 1941; Herrmann 1979 ). Species of Acknowledgments Kelly Ryan and Russel Zimmer both, Phoronopsis and Phoronis, remodel the lar- provided excellent comments on previous versions of this val tentacles into the juvenile form, but in slightly chapter. A portion of the data in this book chapter was gathered at the Smithsonian Marine Station (Fort Pierce, different ways. During metamorphosis, the distal FL) and is designated contribution number 988. Faculty portions of the larval tentacles of Phoronis pal- Research Grants provided by Long Island University-Post lida are cast off and expansion of the tentacle also supported this work. 10 Phoronida 243

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Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger.

S. Santagata Biology Department , Long Island University-Post , Greenvale , NY 11548 , USA e-mail: [email protected]

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 247 DOI 10.1007/978-3-7091-1871-9_11, © Springer-Verlag Wien 2015 248 S. Santagata

INTRODUCTION then particles are gathered in a muscular pharynx. Ingested food particles are then processed in a First considered to be plantlike (moss) animals U-shaped gut. Ectoprocts do not have nephridia, similar to cnidarians, the evolutionary origin and but solid waste is expelled by an anus situated out- affi nities of the ectoprocts or bryozoans have side the tentacles. Feeding zooids, called autozo- been enigmatic subjects of research since the oids, can transfer nutrients to other non-feeding sixteenth century. The term Bryozoa originally polymorphic zooids (e.g., avicularia, kenozooids, encompassed both the entoprocts (or kampto- vibracula, etc.) in the colony via a network of tis- zoans, Chapter 6) and the ectoprocts (Ehrenberg sue cords and communication pores called the 1831 ); however, these animal groups were later funicular system (Banta 1969 ; Best and Thorpe separated (Nitsche 1869 ) and eventually orga- 1985). Neuronal cell bodies of the intraepithe- nized into different phyla (Cori 1929 ). Ectoprocts lial nervous system are mainly concentrated in are aquatic invertebrates that can form elaborate a single ganglion positioned between the mouth and occasionally large colonies (>1 m) composed and anus (Schwaha et al. 2011 ) and are capable of numerous individual zooids, each typically no of integrating sensorimotor information between more than a millimeter in length. Zooids in a col- zooids (Thorpe et al. 1975 ). Collectively, the ony may be one of several different polymorphic lophophore, gut, nervous system, musculature, forms specialized for various functions such as and funicular tissue are grouped together as the feeding, reproduction, or defense. Current esti- polypide. Surrounding the polypide is an outer mates of ectoproct diversity range from 4,000 stratifi ed epithelial layer known as the cystid to 8,000 extant species (Ryland 2005 ), many of that may have a tubular, ovoid, or boxlike shape. which are broadly distributed throughout fresh- Gymnolaemate ectoprocts dominate in brackish water, brackish, and marine environments. More and marine environments and are comprised of than 15,000 fossil species that trace their origins two orders, the uncalcifi ed Ctenostomata and back to the Ordovician period approximately 483 the varyingly calcifi ed Cheilostomata. Further million years ago have been described (Xia et al. taxonomic subdivision of the Ctenostoma has 2007 ). This period of origin is much later than been based mainly on colony-level traits such that of many other animal phyla that arose dur- as whether zooids bud from tubular stolons ing or before the Cambrian (Erwin et al. 2011 ). (Stolonifera, e.g., Bowerbankia and Walkeria ) Although one Cambrian ectoproct fossil has been or form fl eshy sheet-like or globular colonies described (Landing et al. 2010 ), this morphotype (Carnosa, e.g., Alcyonidium , Hislopia , Nolella , has been reinterpreted as a type of octocoral and Sundanella ). Family-level relationships (Taylor et al. 2013 ). Whether the relatively late under these categories inferred from morphologi- geologic origin of the ectoprocts is correct or cal characters differ among authors (e.g., Jebram merely the result of preservational bias against 1992 ; Todd 2000 ). some as-yet-unknown soft-bodied form remains Although no longer considered monophy- an open question, but all extant morphological letic clades (Waeschenbach et al. 2012), differ- grades of ectoprocts with and without mineral- ent morphological grades of cheilostomes ized zooids were clearly present by the Jurassic (Anasca, Cribimorpha, and Ascophora) were (McKinney 1995; Taylor and Ernst 2004 ; previously grouped according to the structure Ostrovsky et al. 2008 ). of the frontal cystid membrane covering the Ectoprocts are currently arranged into three polypide and its role in eversion of the lopho- classes, the Stenolaemata, Gymnolaemata, and phore. The frontal membrane may be a fl exible Phylactolaemata (Bock and Gordan 2013 ). All structure that can be depressed by parietal mus- ectoprocts feed with a lophophore, which is a cles, thus increasing pressure in the coelom circular or U-shaped ring of ciliated tentacles surrounding the polypide, resulting in the surrounding the mouth. Ciliary currents direct eversion of the lophophore (as found in the suspended food particles toward the mouth, and anascans Bugula, Flustra, and Membranipora). 11 Ectoprocta 249

Alternatively, the frontal membrane may be evolutionary hypotheses about ectoproct evolution elaborated with a series of fused calcifi ed spines (Waeschenbach et al. 2009, 2012 ). Strong (e.g., Cribrilina ). Numerous species of cheilo- support exists for the morphological characters stomes have completely calcifi ed their frontal separating the three main classes: Stenolaemata, membranes into a wall with pores of varying Gymnolaemata, and Phylactolaemata. However, numbers and sizes. These species have a mem- some family-level (and higher) divisions based branous sac (ascus) below the frontal wall that is on zooidal frontal wall morphology in cheilo- open to the surrounding seawater (Dick et al. stomes and colony budding types in ctenostomes 2009 ). In these so-called ascophorans, parietal and phylactolaemates have not been completely muscles expand this sac to evert the lophophore congruent with molecular data (Waeschenbach (e.g., Celleporaria , Schizoporella , and et al. 2012 ). Homoplasy among characters asso- Watersipora ). Stenolaemates had their greatest ciated with skeletal features and colonial growth species radiation in the Paleozoic Era, but their strategies is not unexpected as predation and com- diversity was signifi cantly reduced by the petition for space are convergent selective pres- Permian-Triassic and Cretaceous-Paleogene sures acting on all forms of ectoprocts (McKinney mass extinction events (Lidgard et al. 1993 ). 1992 , 1995). Living members of this class are represented How ectoprocts are related to other forms of only by the order Cyclostomata, whose cylindri- bilaterian animals has long been a debated sub- cal calcifi ed zooids lack any specialized closing ject. Based on the morphology of their feeding structure (e.g., an operculum or muscular con- structures, body cavities, and embryonic cleav- striction). Similar to the other morphological age patterns, early work united ectoprocts with grades of ectoprocts, the validity of morpholog- the phoronids (Chapter 10 ) and brachiopods ical classifi cations within the cyclostomes has (Chapter 12 ) under the superphyletic assemblage been questioned (Taylor and Weedon 2008 ). Lophophorata (e.g., Hyman 1959 ), all of whose Cyclostomes occur strictly in marine environ- members were believed to share affi nities with ments, and their colonies can be relatively small deuterostome animals. This interpretation was (such as Crisia or Tubulipora ) in comparison to challenged by the burgeoning fi eld of molecular those of other ectoproct classes. Perhaps the phylogenetics, based largely on 18S ribosomal most divergent morphological, and evolution- DNA (Halanych et al. 1995 ), placing the ecto- arily controversial, zooidal traits are found in procts in a new superphyletic assemblage of pro- the strictly freshwater phylactolaemates whose tostome animals called the Lophotrochozoa, in colonies are often described as being either which the evolutionary position of the ectoprocts gelatinous and saclike or a series of chitinous, relative to phoronids, brachiopods, and spira- branched tubes (Wood and Lore 2005 ). Unlike lians was unresolved. Increased taxon sampling other ectoprocts, phylactolaemate zooids have a bolstered with data from more rDNA genes, full hollow neuronal ganglion (Gruhl and mitochondrial genomes, and select nuclear genes Bartolomaeus 2008 ; Schwaha and Wanninger continued to support these evolutionary scenar- 2012 ), bud from a commingled fl eshy mass ios (Passamaneck and Halanych 2004 , 2006 ; (some of which can “crawl” along the bottom Jang and Hwang 2009 ). More recent phyloge- like Cristatella ), and have a U-shaped lopho- nomic approaches supported the resurrection of phore. The latter trait has occasionally been the Polyzoa concept in which ectoprocts are regarded as a possible synapomorphy with linked with entoprocts (Hausdorf et al. 2007 , another member of the lophophore-bearing ani- 2010; Helmkampf et al. 2008; Hejnol et al. mals, the phoronids (Chapter 10 ; Hyman 1959 ). 2009). Interestingly, the latest phylogenomic A robust molecular phylogeny of the ectoprocts analysis supports reuniting all the lophophorates with extensive taxon sampling is still lacking, but in one monophyletic group, where the phoronids signifi cant progress toward this goal has been are a sister taxon to the ectoprocts (Nesnidal made, allowing for the evaluation of some macro- et al. 2013 ). 250 S. Santagata

EARLY DEVELOPMENT and equal, resulting in an eight-cell stage in which all of the pigmented cytoplasmic granules are Ectoproct reproductive patterns and brooding contained in the four vegetal cells. Fourth cleav- styles (where applicable) have been reviewed age is through the AV axis resulting in a bilater- extensively (Reed 1991 ; Temkin 1994 , 1996 ; ally symmetric 16-cell embryo, composed of two Santagata and Banta 1996; Ostrovsky et al. 2009 ; rows of four equally sized cells in the animal half Ostrovsky 2013 ) and will be only briefl y men- of the embryo directly stacked on the eight cells in tioned here in the context of particular species the vegetal half of the embryo. However, fourth for which early development has been followed cleavage in the vegetal cells is not equal, and as a in some detail. The vast majority of information result the four middle blastomeres of the eight on the early development of ectoprocts comes vegetal cells are larger than the adjacent cell pairs from a select set of species that hardly repre- (Fig. 11.1H). Third and fourth cleavage planes sents the rich diversity of the three major clades. and cell arrangements of 8- and 16-cell stages are These data are entirely descriptive and based on also entirely radial. Fifth cleavage results in a the cell arrangements found in living and fi xed 32-cell stage comprised of 4 tiers of cells consist- embryos, but as of yet no detailed cell lineage ing of 8, 12, 8, and 4 cells from the animal to the study using modern cell tracing techniques exists vegetal pole. Gastrulation begins with the inter- for any ectoproct species. For now, this review nalization of the four vegetal cells as the embryo of ectoproct early embryology is based on a few nears the 64-cell stage (Fig. 11.1J, K ). According key species and compares these accounts to data to Zimmer (1997 ), representative species of gym- gathered using cytological fl uorescent probes and nolaemates are consistent with respect to the con- confocal microscopy. tributions made to larval and presumptive juvenile Anascan cheilostome ectoprocts, such as tissues by the four tiers of cells at the 32-cell Membranipora spp., that broadcast spawn fertil- stage. The top animal tier of eight cells is respon- ized oocytes into the water column (Temkin 1996 ) sible for forming the apical disc composed of the and develop into a characteristic feeding larval sensory and neuronal cells of the larval apical form called the cyphonautes (Kupelweiser 1905 ) organ (see Santagata 2008a for details) and (if are generally the easiest model species for the present) the epidermal blastemal cells that will study of early development due to their numerous form ectodermal and endodermal tissues of the gametes, ease of collection, and relatively fast ancestrular polypide. developmental times (Fig. 11.1 ). Many of the In species with a feeding cyphonautes larva recent developmental observations of this genus (e.g., Membranipora ), the second animal tier of come from genetically distinct clades and perhaps eight cells will form hundreds of multiciliated three different species (Schwaninger 2008 ) occur- coronal cells used for locomotion. However, ring on kelp growing on the fl oating docks adja- there is considerable variation in the number and cent to Friday Harbor Laboratories (University of morphology of coronal cells among species. In Washington, WA, USA). Once activated by expo- some non-feeding larval forms, these cells may sure to seawater, the fl attened disc shape of a undergo cleavage arrest at an early stage, forming Membranipora-fertilized primary oocyte (approx. a row of only 32 or 40 cells in the coronal band 60 μm in diameter) will round up, produce fi rst (such as species of Tanganella , Watersipora , and second polar bodies, and enter fi rst cleavage Amathia , and Bowerbankia; also see Corrêa within approximately 1–2 h (12–14 °C). First 1948). There are hundreds of strap-like coronal cleavage is equal and through the animal-vegetal cells in Bugula neritina (Woollacott and Zimmer (AV) axis. Second cleavage is also equal and 1975 ), but the larvae of Sundanella sibogae and through the AV axis, but rotated 90° to that of the Nolella stipata have numerous corona-like cells fi rst cleavage plane. Pigmented cytoplasmic gran- covering the aboral half of the larva (Santagata ules are segregated to the vegetal side of cells by 2008a ). The third tier of 12 cells is more vari- the four-cell stage. Third cleavage is equatorial able in its contribution to larval and presumptive 11 Ectoprocta 251

A BCD

E FG H

I JK L

M NOPQ

RST

Fig. 11.1 Spawning and development of Membranipora nucleic acids, red fi brous actin). (K ) 64-cell stage. ( L ) membranacea . (A ) Fertilized zygotes being released from Early gastrula. (M ) Midgastrula stage where the clear the intertentacular organ ( ITO ). ( B ) Close-up of the ectodermal cells spread over the embryo. ( N) Late gas- spawning zooid in ( A ). (C ) Rounded-up zygote sur- trula stage with the initial invagination of the vestibule. rounded by the wrinkled fertilization envelope 1–2 h post ( O ) The beginning of larval development during which spawning. ( D) Two-cell embryo. (E ) Vegetal view of an the vestibule ( V) elongates. (P ) Early trapezoidal-shaped eight-cell embryo. ( F) Eight-cell embryo undergoing cell larval stage hatching out of the fertilization envelope; note division to the 16-cell stage. (G ) Vegetal view of a 16-cell the ciliation on the coronal cells (C ). (Q ) Hatched feeding embryo. ( H ) Partial volume rendering of the 16-cell stage larval stage with a defi ned apical disc ( AD ). (R ) Late- embryo showing the four larger central macromeres stage larva with a pyriform complex ( PO), ciliated ridges ( arrow ) in the vegetal half of the embryo (green nucleic ( R), and internal sac (IS ). ( S) Confocal z-projection of a acids, red fi brous actin). (I ) Vegetal view of a 32-cell late-stage larva stained for muscles ( red) and acetylated embryo showing the four macromeres ( arrow ) that will be α-tubulin (green ) for cilia and nervous system. (T ) Larval internalized during gastrulation at the 64-cell stage. ( J ) stage that is competent to metamorphose; note growth of Optical section through the four internalized macromeres internal sac ( IS ) and the relatively narrow gut (G ). Scale (one of which is indicated by the arrow) surrounded by bars in A – C and I – T equal 50 μm, scale bars in D – H the blastomeres that will become the coronal cells ( green equal 30 μm 252 S. Santagata

juvenile tissues. In all species investigated, these proceeds creating a two-cell-layered primary cells form the larval pyriform complex consist- embryo that will grow in size as the ovary degen- ing of a superior and an inferior glandular fi eld erates. The enlarged primary embryo will then of cells bordering four specialized multiciliated produce other genetically identical (Hughes et al. cells that constitute the vibratile plume. The 2005 ; Pemberton et al. 2007 ) secondary embryos pyriform complex has numerous other sensory through polyembryony. In Crisia pugeti secondary and neuronal cells (Reed et al. 1988 ; Santagata embryos are produced from bilayered projections 2008b ) used during crawling behaviors to that bud off the primary embryo (Dimarco-Temkin probe various substrates before metamorpho- and Temkin, pers. comm.). In this species, second- sis. Another derivative of this cell tier is the oral ary embryos consist of a central group of eight epithelium that consists of ciliated cells whose cells surrounded by an outer layer of 32 cells. The primary function is to capture food particles (the outer epithelium continues to develop, forming an ciliated ridges and cell lining of the vestibule) in invaginated aboral epithelium and an internal sac feeding larvae. In non-feeding larval forms these (Barrois 1877 ). These tissues form the cystid at cells make an oral fi eld of ciliated cells that can metamorphosis, leaving the internal group of cells be used in both crawling and swimming behav- to form the polypide. iors. In some ctenostome larval forms lacking The development of freshwater ectoprocts is the epidermal blastemal cells in the apical disc, the least understood and, despite their ancestral a portion of the oral epithelium remains undiffer- position in most current phylogenetic analyses, entiated and is the source of the ectodermal and may have the most derived developmental pat- endodermal tissues of the ancestrular polypide terns found among all ectoproct clades. What little (Zimmer and Woollacott 1993 ). The last struc- information that exists comes from a few sources ture derived from this cell tier is the internal sac, (Braem 1897 ; Brien 1953), and much of what a simple or complex eversible epithelium that has been reviewed (see Reed 1991 ; Zimmer and is used for temporary or permanent attachment Woollacott 1993; Zimmer 1997; d’Hondt 2005 ) to the substrate at metamorphosis. Larval types focuses on one species, Plumatella fungosa . Early with a complex internal sac form all or part of the cleavages have been described as holoblastic and ectoderm of the cystid from it (Lyke et al. 1983 ). irregular, producing an elongated, hollow blastula The fourth tier of four (vegetal) cells internalized consisting of a single layer of cells. Mechanisms during gastrulation will make the endodermal of gastrulation are unclear (possibly ingression), cells of the larval gut in feeding larvae and either but cells invade the blastocoel (approx. 72-cell a partial nonfunctional gut or nutrient-rich cells embryo) and form an early gastrula consisting of in non-feeding larvae. The fourth cell tier also two cell layers (note similarity to primary embryo makes the larval mesodermal tissues (striated formation in cyclostomes). The embryo then grows and smooth musculature; see Santagata 2008a ) due to the nutrients provided by the maternal zooid as well as those blastemal cells situated near the via the tissue connection between the ovisac and apical disc that will form the ancestrular meso- the equatorial region of the embryo. The bilay- dermal tissues. It should be made clear that varia- ered embryo then forms one, two, or four differ- tion in the formation of and contribution to these entiated polypides and sometimes polypide buds latter cell groups was discussed by Prouho (1892 ) (depending on the species). These polypides will and also that future investigations of ectoproct eventually be covered by the proliferating mantle embryonic cell lineages are likely to fi nd more tissue, except for a small pore at the lagging end differences as data for more species are gathered. of the larva. The outer epithelium of mantle tissue Stenolaemate (cyclostome) ectoprocts brood becomes ciliated, and the leading (swimming) end their embryos in specialized enlarged gonozooids. of the ciliated epithelium develops an apical neu- Fertilization occurs within the ovary, and the ronal plate (Franzén and Sensenbaugh 1983 ) with resulting early cleavage stages are thought to be unclear homology to the apical disc (apical organ) irregular in cell number (Borg 1926 ). Cleavage of other ectoproct larvae. 11 Ectoprocta 253

LATE DEVELOPMENT internal sac and are used to spread this adhesive secretory epithelium over the substrate at meta- Most gymnolaemate larval types can be grouped morphosis. During this time the internal sac will into one of three categories, a thin pyramidal- be joined with the pallial epithelium to form the shaped feeding cyphonautes, a squat pyramidal- cystid (Stricker 1988). Blastemal cells that will shaped non-feeding pseudocyphonautes, and eventually form the polypide are pulled into the various forms of oblate spheroid-shaped non- center of the pre- ancestrula by retractor muscles feeding larvae (the most common being the coro- positioned between the apical disc and the pyri- nate larva). The cell types and anatomy of the larval form complex. Internalized larval tissues (e.g., nervous system and musculature have been better corona and pyriform organ) will undergo histoly- studied in selected species of gymnolaemates, and, sis during metamorphosis. Divisions between collectively, these data support some broad-scale lateral regions of the ancestrula are evident in homology with the larval neuromuscular systems Membranipora , producing a twinned ancestrula of spiralian protostomes (Santagata 2008b ; Gruhl of two complete autozooids. Developmental time 2009 ; Wanninger 2009 ) but differ from the larval for the ancestrula to become a functional feeding morphologies of phoronids and brachiopods (cf. zooid varies with temperature, but in general it Chapters 10 and 12 ; Santagata and Zimmer 2002 ; occurs within two days. Altenburger and Wanninger 2009 ; Santagata 2011 ; Non-feeding (coronate) larval forms found in Temereva and Wanninger 2012 ). ctenostome and cheilostome ectoprocts some- All gymnolaemate embryos have a late gas- what retain the morphology of their late gastru- trula stage with a comparatively large ring of lae (Fig. 11.2 ). Major differences occur in the coronal cells equatorially arranged between the position of the apical disc relative to the pyri- more cell-rich oral and aboral larval fi elds. In spe- form complex that correlates with the orienta- cies with a cyphonautes larva, the entirety of the tion of the larvae during swimming and crawling oral cell fi eld will be internalized as the embryo behaviors (Fig. 11.3 ; see also Santagata 2008a , fl attens laterally and takes on a more trapezoidal b for review). The positions of blastemal cells shape (Fig. 11.1N ). As a consequence of this pro- that eventually make the polypide can also cess, the outer pallial epithelium now covers the vary considerably. These cells may be found in surface of the embryo except for where it meets or below the apical disc, in centralized bands the border of the developing apical disc and the inside the larva, and also bordering the oral cili- coronal cell band. At this stage the vestibular cav- ary fi eld (Zimmer and Woollacott 1989 , 1993 ). ity inside the larva is formed. The vestibule con- Considerable differences are found in the mecha- tains the ciliated ridges and other ciliated cells, nisms of morphogenetic movements of the larval the function of which is to bring food particles and presumptive juvenile tissues at metamorpho- to the apically positioned mouth (Fig. 11.1R ). sis as well as the contributions made to the ances- Substantial circular musculature lines the mouth trula by specifi c types of presumptive juvenile and esophageal regions (Stricker et al. 1988a ; tissues in the larva. Santagata 2008a , b ) that lead into the gut and ter- Some general differences in anatomy among minate at the anus positioned at the posterior end phylactolaemate, cyclostome, ctenostome, and of the coronal cell band. The pyriform complex, a cheilostome larval types are summarized in sensory and glandular organ that includes a cen- Fig. 11.4 . Although there are convergent mor- tral group of four cells that make fused ciliary tufts phologies present between vesiculariform (e.g., collectively called the vibratile plume, and also Amathia , Bowerbankia , and Zoobotryon ) and the presumptive juvenile tissues (blastemal cells buguliform ( Bugula and Scrupocellaria ) lar- and the internal sac, see Stricker et al. 1988b ) are val types, each has divergent structure in the not present at the early feeding stage. Late-stage presumptive juvenile tissues, musculature and metamorphically competent larvae develop involved in metamorphosis, and blastemal con- large, paired retractor muscles that insert on the tributions to the ancestrula. For instance, larval 254 S. Santagata metamorphosis in Bowerbankia imbricata begins gle tubular zooid that will bud asexually from with the eversion of the internal sac via constric- basal stolons. Similar to Bowerbankia larvae, the tion of the equatorial ring muscle. The larva eversion of the internal sac in Bugula larvae is temporally adheres to the substrate using cellu- muscle mediated. Evagination of the small pal- lar secretions of the internal sac, but the sac is lial epithelium and involution of the coronal cells degraded later in metamorphosis. The coronal are also accomplished by the reversal of coronal cell epithelium involutes from the force exerted cell ciliary beat (Reed and Woollacott 1982 ). In by the reversal of ciliary beat against the pellicle, contrast to Bowerbankia larvae, the raising of the and at the same time the expansive pallial epithe- walls of the internal sac and contraction of the lium spreads over the entire surface of the pre- pallial epithelium in Bugula larvae are completed ancestrula, forming the entire cystid epithelium by the actions of bands of microfi laments in the (Reed and Cloney 1982 ). The internal band of pallial epithelium (Reed and Woollacott 1982 ). blastemal cells in Bowerbankia larvae is already The cystid of the ancestrula of Bugula neritina centralized within the ancestrula by these tissue is composed solely of the internal sac, and the rearrangements and does not require the actions pallial epithelium is largely vestigial, forming of larval musculature, eventually forming a sin- only a portion of the tentacle sheath (Woollacott

A BC

DEF

Fig. 11.2 Early development of gymnolaemate ecto- embryo. ( D) Optical section though a 64-cell stage of procts with a non-feeding coronate larva (confocal z-pro- Dendrobeania lichenoides, showing the four internalized jections, specimens stained for fi brous actin (red ) and macromeres ( M) at gastrulation. (E ) Late gastrula stage of nucleic acids ( green )). ( A) Animal view of a 32-cell stage Dendrobeania lichenoides; note the fewer and larger cells of Dendrobeania lichenoides (embryos dissected out of that will eventually make the coronal band ( C ). ( F ) Early ovicells from colonies collected from the fl oating docks at larval stage of Dendrobeania lichenoides in which the Friday Harbor Laboratories, University of Washington, coronal cell band ( C) is differentiated and demarcates the WA, USA). A median band of cells along the equator of boundary between the aboral and oral cellular fi elds, both the embryo will make the coronal cell ( C) ring. (B ) of which have many more cells. The approximate position Animal view of a 32-cell stage of Hippodiplosia insculpta of the developing apical disc (AD ) is labeled. Scale bars with polar bodies ( PB ). ( C ) Side view of the 64-cell stage equal 50 μm of Rhynchozoon rostratum; note bilateral symmetry of the 11 Ectoprocta 255 and Zimmer 1971 ; Reed and Woollacott 1982 ). of metamorphosis in the numerous and morpho- In this species, the bilayered groups of blaste- logically divergent species of ascophoran-grade mal cells below the apical disc are centralized cheilostomes and most families of ctenostome by axial muscles running between the apical ectoprocts. Preliminary work on these larval disc and the roof epithelium of the internal sac. types supports some aspects of previously dis- Overall, very little is known about the process cussed processes, but more diversity exists in

A B

CD

Fig. 11.3 Aspects of larval morphology among non-feed- Parasmittina spathulata , an ascophoran-grade cheilo- ing forms of gymnolaemate ectoprocts. ( A ) Scanning stome. This species has a larger apical disc and several electron micrograph (SEM) of the larva of Aeverillia pigmented ocelli ( OE ). (F ) SEM (lateral view) of the setigera , a ctenostome that is elongated in the aboral-oral coronate larval type of Schizoporella fl oridana , another axis and fl attened in the anterior-posterior axis; observe ascophoran-grade cheilostome; note the relative differ- the small apical disc ( AD). Coronal cells (C ) cover the ences in the size of the ciliary fi elds of the coronal and oral aboral hemisphere of the larva, and the expansive oral cili- ciliated cells. ( G ) Fluorescent light micrograph of a ated cells ( OC ) cover the aboral hemisphere. This species Bugula stolonifera larva, an anascan-grade cheilostome also has a long oral ciliated groove. (B ) SEM of the fused that is elongated in the aboral-oral axis, has an expanded ciliary bundles of the vibratile plume cells (VP ). (C ) Light coronal cell fi eld and has a long oral ciliated groove ( OG ) micrograph of the larva of Nolella stipata, another cteno- bordered by ocelli. The larva is stained with bisbenzimide stome that is elongated in the aboral-oral axis. The homol- for nucleic acids. ( H) Fluorescent light micrograph of a B. ogy of the multiciliated cells ( MC) that cover the larval stolonifera larva stained with DASPEI, a live mitochon- epidermis to the coronal and oral ciliated cells of other drial probe that shows the morphology of the ciliated ray species is unclear (see Santagata 2008a for details). As cells (neurons) ( CR ) in the apical disc surrounded by the found in the larva of A. setigera , the aboral and oral ciliary dark, unstained pallial epithelium ( PE ) that surrounds the fi elds are separated by an equatorial band of sensory neu- apical disc. Scale bar in A equals 100 μm, scale bars in rons with small ciliary tufts ( IMC ). (D ) SEM of N. stipata . B – D equal 10 μm, scale bars in E – H equal 50 μm ( E) Light micrograph of the coronate larval type of 256 S. Santagata

EF

GH

Fig. 11.3 (continued) larval anatomy, the structure of presumptive with the internal sac and collectively forms the juvenile tissues, and tissue rearrangements at cystid. How the polypide is formed from largely metamorphosis. undifferentiated cells inside the larva is not well The larval anatomy of cyclostome ectoprocts understood. A central group of blastemal cells has been interpreted as simplifi ed or degenerate, (that may include both epidermal and mesoder- since their larvae are the only type to lack an apical mal cells; see Nielsen 1970 ) between the aboral disc (and associated larval apical sense organ) and epithelium and the internal sac are certainly pre- a pyriform organ and their nervous system con- cursors of the polypide, but whether or not these sists of a basiepithelial network of sensory cells cells are the only contributors to the polypide is and neurons (Santagata 2008b ). It is not clear if the unclear. As discussed earlier, the larvae of fresh- cuboidal multiciliated cells that make up the lar- water ectoprocts exhibit a form of adultation, as val epidermis are homologous to the coronal cells the polypide(s) are already formed in the body and/or oral ciliated cells of gymnolaemate larvae. of the larva before metamorphosis (Fig. 11.4 ). Metamorphosis of cyclostomes has been In contrast to all other described ectoproct lar- best studied in species of Crisia (Nielsen 1970 ; val forms, phylactolaemate larvae settle, probe, d’Hondt 1977 ). Similar to gymnolaemate lar- and initially attach to the substrate using an api- vae, the eversion of the internal sac at meta- cal plate comprised of both sensory neurons morphosis is muscle mediated, but in Crisia it and glandular cells (Franzén and Sensenbaugh is accomplished through the actions of small 1983 ; Sensenbaugh and Franzén 1998 ; Gruhl longitudinal muscle fi bers in the body wall 2010 ). For these reasons it has been hypoth- (d’Hondt 1977 ; Santagata 2008a ). The invagi- esized that the glandular cells are homologous nated aboral epithelium (likely homologous to to the internal sac of other ectoproct larval the pallial epithelium of gymnolaemates) fuses forms, particularly those with vesiculariform 11 Ectoprocta 257

Fig. 11.4 Evolutionary trends in larval anatomy and the ( VC, only a subset are fi gured) found in species with a arrangement of the presumptive juvenile tissues that form planktotrophic larva or are the oral ciliated cells ( OC ) of the ancestrula among various systematic groups of ecto- some non-feeding larval types. Structures found in most procts. Tissues shaded in blue (pallial epithelium) and larval forms include the apical disc ( AD) and vibratile black (internal sac) contribute the cystid epithelium (here plume cells ( VP ). Cyphonautes larvae are covered by a shown as either a tubular or rectangular box ) at metamor- bivalved shell (CS ). It is unclear if the multiciliated cells phosis. In species where one of these tissues does not con- ( MC) that cover the larvae of cyclostomes and phylac- tribute to the ancestrula at metamorphosis, either the tolaemates are homologous to the coronal and/or oral cili- pallial epithelium ( PE ) or internal sac (IS ) is unshaded. ated cells of gymnolaemate ectoprocts. Furthermore, the Tissues shaded in yellow are groups of blastemal cells that axial properties of phylactolaemate larvae are not well form the polypide (internal components of the lophophore established, and it is not clear if the apical disc ( AD ) and and gut) after metamorphosis. Cells shaded in red consti- pallial-like epithelium (PE? ) are homologous to these tute the coronal cell ring ( C ), mainly used for ciliary pro- structures in gymnolaemates (© Scott Santagata 2015. All pulsion. Cells shaded in green either form the ciliated Rights Reserved) ridges ( CR) and other ciliated cells within the vestibule

(ctenostome) types. However, the presence of ides form the walls of the cystid (Sensenbaugh sensory neurons along with the glandular cells and Franzén 1998 ). Forming an expansive cystid complicates this comparison, especially since epithelium from one epithelial source is similar both neuronal and glandular fi elds already exist to what is observed in the Vesiculariidae (Reed in the pyriform complex of most other ecto- and Cloney 1982 ) and at least one other cteno- proct larval forms. During the initial phases of stome taxon, the Aeverrilliidae (see Santagata metamorphosis, the mantle tissue that underlies 2008a ). the ciliated outer epithelium of the larva invo- lutes. This tissue rearrangement internalizes the residual larval tissues inside the pre-ancestrula GENE EXPRESSION (serving as a nutrient source) and also exposes the inner epithelium (starting at the point of the Gene expression studies on ectoprocts are terminal pore) of the vestibular chamber around in the early stages of exploration. No data the internal polypides. In this way the vestibular exist on developmental-related genes during epithelium and other tissues around the polyp- embryogenesis, but some limited information 258 S. Santagata has been published on the larvae and metamor- the cell fates of the blastemal cells of Bugula phic stages of Bugula neritina . One prevalent neritina larvae are determined before metamor- question deals with the potency and differen- phosis and support homologous cell differentia- tiation of the presumptive ancestrula tissues in tion mechanisms among metazoan adult body the body of the larva. Clearly, the pallial epi- plans. However, addressing the pluripotent or thelium and internal sac are formed of largely multipotent potential of blastemal cells will undifferentiated cells during the larval stage, as require detailed tracking of these cells’ expres- their functions are mainly to secrete and attach sion patterns and cell fates through metamor- the pre-ancestrula to the substrate (depend- phosis. Some of these data have been gathered ing on the species), and are not in their fi nal in Bugula neritina for select genes in the WNT state of differentiation (cystid wall epithelium signaling pathway and support the interpreta- composed of ectodermal cells) until a feeding tion that blastemal cell fate is determined before zooid stage is reached. Blastemal cells eventu- metamorphosis. Interestingly, WNT10 and sFRP ally form all the neural, muscular, coelomic, (secreted frizzled-related protein) are expressed and endodermal tissues of the polypide. So, col- in spatially opposite cell domains of the epider- lectively, all of the presumptive juvenile tissues mal and mesodermal blastemal cells at the lar- are, at the very least, composed of multipotent val stage and the differentiating polypide at the cells. However, since all of the variously posi- early ancestrula stage in Bugula neritina (Wong tioned blastemal cells of ectoproct larval types et al. 2012 , 2014 ), suggesting that these genes are either epidermal and/or mesodermal in ori- play a role in defi ning the spatial polarity of the gin (Fig. 11.4), it has been suggested that only polypide (see Fig. 11.5 ). these germ tissue types form the entirety of the adult body (Woollacott and Zimmer 1971 ; Reed and Cloney 1982 ; Reed 1991 ; Zimmer 1997 ). OPEN QUESTIONS Essentially, this hypothesis interprets ances- trula (polypide) formation at metamorphosis • How do detailed cell lineages of various as homologous to the process of forming a new species of ectoprocts compare with those zooid by asexual budding, in which the new of brachiopods, phoronids, and spiralian polypide is differentiated from cells originat- protostomes? ing from the frontal cystid epithelium. If this • How are ectoproct embryos, larvae, and various hypothesis is correct, one prediction would be kinds of zooids patterned during development that blastemal cells should not express endo- by differential gene expression (e.g., Hox, dermal differentiation factors during the larval Parahox, and other key developmental genes)? period. Furthermore, these factors should only • Are any larval tissues retained or remodeled be expressed during the later stages of ances- during metamorphosis when the adult body trula formation. Developmental genes such as plan is formed? Cdx , FoxA , and GATA456 are expressed in the • What other aspects of larval morphological endoderm of other invertebrate animals (Hejnol diversity and metamorphic patterns exist in the and Martindale 2008 ; Boyle and Seaver 2010 ). numerous unexplored species of ectoprocts? In the larva of Bugula neritina , Cdx and FoxA are expressed in the internal sac and most of the epidermal and mesodermal blastemal cells, but Acknowledgments Kelly Ryan and Russel Zimmer pro- GATA456 is only expressed in a limited sub- vided excellent comments on previous versions of this set of epidermal blastemal cells (Fuchs et al. chapter. A portion of the data in this book chapter was 2011). These expression patterns along with gathered at the Smithsonian Marine Station (Fort Pierce, FL) and is designated contribution number 989. Faculty others gathered from neural- and mesodermal- Research Grants provided by Long Island University-Post related genes (Fuchs et al. 2011 ) suggest that also supported this work. 11 Ectoprocta 259

A B

Fig. 11.5 Select gene expression cell domains in the nucleic acids; green , acetylated α-tubulin). (B ) Shaded larva of Bugula neritina . ( A ) Frontal optical section illustration showing select gene expression domains in the through the larval body showing the epidermal ( EB ) and blastemal cells and internal sac. Gene expression data mesodermal ( MB) blastemal cells as well as the complex from Fuchs et al. ( 2011 ) and Wong et al. (2012 ). Scale bar internal sac ( IS) that forms all of the cystid (pallial epithe- in A equals 50 microns lium forms only the tentacle sheath in this species; red ,

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Chapter vignette artwork by Brigitte Baldrian. © Brigitte Baldrian and Andreas Wanninger.

S. Santagata Biology Department , Long Island University-Post , Greenvale , NY 11548 , USA e-mail: [email protected]

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 263 DOI 10.1007/978-3-7091-1871-9_12, © Springer-Verlag Wien 2015 264 S. Santagata

INTRODUCTION musculature among rhynchonellids, craniids, and lingulids that may be homologous (Altenburger Enclosed in shells with ventral and dorsal and Wanninger 2009 ; Santagata 2011 ). Both valves, extant brachiopods (meaning “arm” and craniiform and linguliform adult brachiopods “foot”) are classifi ed into three major subphyla: have through guts, but the anus is lacking in the Rhynchonelliformea, the Linguliformea, rhynchonelliforms. This absence is generally and the Craniiformea (Williams et al. 1996 ). considered a secondary loss, possibly due to the Rhynchonelliform brachiopods encompass what reduction of water fl ow in the posterior part of the were once referred to as the “articulate” brachio- mantle cavity associated with the presence and pods, so named for the mineralized hinge that position of the hinge (LaBarbera 1981 ; Williams connects the calcite valves of their shells. No et al. 1997 ). such hinge is found in members of the other two Currently, the oldest known brachiopods in subphyla, rather their valves are held together the fossil record are from the lower Cambrian, only by various muscles and connective tissues. belonging to an extinct stem group called the Craniiform brachiopods (e.g., Novocrania ) also paterinids (Topper et al. 2013 ). Paterinid brachio- have calcitic shells, but the shells of linguliform pods had organophosphatic shells linking them to brachiopods (such as the lingulid Glottidia and linguliform brachiopods, but their shell morphol- the discinid Discinisca ) are composed of apa- ogy was more similar to that of rhynchonelliform tite, a phosphatic mineral, with an outer layer of brachiopods. Paterinid shell ultrastructure dif- chitin (Williams et al. 1997 ). Most brachiopod fered from all other crown fossil brachiopods but morphotypes have a smaller dorsal and a larger shared some characteristics with the organophos- ventral valve, the latter of which often bears a phatic sclerites of the extinct metazoans known muscular or rigid attachment structure called as tommotiids (Larsson et al. 2013 ). Whether or the pedicle. Rhynchonelliform brachiopods not particular tommotiid morphotypes that may are often attached to hard substrata by the rigid have been sessile fi lter feeders (such as pedicle with their ventral valves oriented upward Eccentrotheca; see Skovsted et al. 2011 ) repre- (Richardson 1997 ). The shells of linguliform bra- sent the stem lineage for crown phoronids, pateri- chiopods such as Glottidia and Lingula generally nids, and crown linguliform brachiopods remains have equally sized valves and their pedicles are unclear (Murdock et al. 2014). Molecular esti- long, muscular structures modifi ed for burrowing mates place the origin of the crown brachiopods into soft sediments. Craniiform brachiopods have in the Middle Cambrian, but the phoronid- lost the pedicle and cement directly to hard sub- brachiopod root is older, possibly originating as strates (Emig 1997 ). far back as the Ediacaran (Sperling et al. 2011 ). Beyond differences in their mineralized parts, Although molecular evidence is generally there are also morphological and functional dif- congruent with the paleontological origin of bra- ferences in brachiopods’ main feeding and respi- chiopods, discrepancies do exist, leaving open ratory structure, the lophophore, which can be the question of whether phoronids should be con- composed of looped, spiraled, or U-shaped fi elds sidered as an ancestral sister taxon to all brachio- of ciliated tentacles (Emig 1992 ). Only rhyn- pods (Sperling et al. 2011 ; although see Thomson chonelliform brachiopods support the lopho- et al. 2014 ) or instead as a subtaxon within the phore with calcifi ed support structures called brachiopods as a whole (Cohen and Weydmann brachidia, whereas all other brachiopods use only 2005 ; Santagata and Cohen 2009 ; Cohen 2013 ). musculature and connective tissue. Types and Molecular phylogenetic data support the evolu- arrangement of adult musculature typically sup- tionary affi nities of brachiopods with spiralian port a closer relationship between craniids and protostomes such as nemerteans, annelids, and linguliform brachiopods (Williams et al. 1996 ); mollusks (Halanych et al. 1995 ; Dunn et al. 2008 ; however there are also subsets of early juvenile Hejnol et al. 2009 ; Hausdorf et al. 2010 ; 12 Brachiopoda 265

Mallatt et al. 2012). Interestingly, a recent called the metacoel). The gonads of both phylogenomic study supports the monophyly of craniiforms and rhynchonelliforms develop all lophophorate phyla in which brachiopods are inside mantle coelomic canals positioned in the still closely related to both phoronids and ecto- mesothelial linings that underlie both shell procts (Nesnidal et al. 2013 ). In general, higher- valves. Many brachiopods have separate sexes, level systematic groups within extant brachiopods but as more reproductive information has been are supported by molecular evidence (Cohen gathered, it is clear that select species of rhyn- et al. 1998 ; Cohen 2000 ), but superfamily-level chonelliform brachiopods are hermaphroditic relationships based on morphology within the (Kaulfuss et al. 2013 ). Furthermore, although rhynchonellid brachiopods are not congruent populations of Glottidia pyramidata from Tampa with molecular phylogenetic data (Cohen and Bay, Florida, are mostly gonochoristic (1:1 ratio Bitner 2013 ; Schreiber et al. 2013 ). of sexes), when isolated from other conspecifi cs, Fossil and molecular data aside, modern a small proportion of the population (0.7 %) may brachiopod species represent only a fraction of become hermaphroditic (Culter and Simon their former morphological and species diversity 1987 ). Similar patterns of gonochorism and her- (approx. 30,000 described fossil species). Current maphroditism also occur in Lacazella caribbea- assessment of brachiopod species diversity rec- nensis (Seidel et al. 2012 ). When spawned, sperm ognizes only 25 linguliforms, 18 craniiforms, and and eggs are released into the coelomic cavity, 348 species of rhynchonelliforms (Emig et al. and, once there, the ciliary beat of the coelomic 2013). The disparity in species number among lining transports the gametes to the paired meta- the systematic groups may be due in part to the nephridial gonoducts. In broadcast spawning reliance in many species descriptions on shell species such as Lingula anatina (Yatsu 1902 ), and brachidia characteristics that in rhynchonelli- Glottidia pyramidata (Paine 1963 ), and forms are more distinct than in (or that are absent Novocrania anomala (Nielsen 1991 ), gametes in) the other two subphyla. Some cosmopolitan are expelled via the gonoducts and are released species of Lingula may actually be cryptic spe- into the surrounding seawater by ciliary currents cies complexes (Yang et al. 2013 ). Comparatively produced by the lophophore. Among the rhyn- few taxonomists work on living brachiopods (but chonelliform brachiopods for which data are see Bitner 2006), but hopefully some of the work available, gametes are either broadcast spawned reviewed here will inspire a new generation of as in Terebratalia transversa (Long and Stricker evolutionary developmental biologists to study 1991) or the eggs are retained in various brood these intriguing animals that once dominated sites (mantle, lophophore, or a pouch), where ancient seas (Rong and Cocks 2013 ). they are cross-fertilized, or in the case of some hermaphroditic species may also be self-fertilized (Kaulfuss et al. 2013 ). EARLY DEVELOPMENT In general, linguliform species with planktonic feeding stages such as Discinisca strigata , Gametogenesis Glottidia pyramidata , and Lingula anatina pro- duce more numerous and smaller (65, 90, and Reproductive patterns in brachiopods have been 100 μm in diameter, respectively) mature oocytes reviewed in some detail (Long and Stricker 1991 ; than craniiform or rhynchonelliform species Kaulfuss et al. 2013 ), and so only pertinent (Yatsu 1902 ; Paine 1963 ; Freeman 1995 , 1999 ; aspects are discussed here. Gonads develop from Kaulfuss et al. 2013 ). Craniiform and rhyn- mesothelial folds surrounding the gut. chonelliform species that have non-feeding larval Linguliforms develop two pairs of gonads in the types develop from more variably sized oocytes, mesenteries adjacent to the gut, which when ripe approximately 100–300 μm in diameter among expand into the main body cavity (typically species (Kaulfuss et al. 2013 ). 266 S. Santagata

A

B

C

Fig. 12.1 Variation in early cleavage stages of lingulid, Hemithiris ) have open radial cell arrangements. (C ) The craniid, and rhynchonellid brachiopods. (A ) Eight-cell less common forms of 16-cell stages among brachiopods, stages of various brachiopods. Stacked and radial unipla- including the four-tier cell stacking found in Novocrania nar cell arrangements are found in Terebratulina and (All stages redrawn from Nielsen 1991 ; Freeman 1995 , Terebratalia . ( B ) The more prevalent forms of 16-cell 1999 , 2003 ). Two polar bodies at the site of the animal stages among brachiopods. Lingulids typically have a pole are shaded in black bilateral cell arrangement and rhynchonellids (except for

Cleavage, Gastrulation, and Germ 2000 ) with, interestingly, some embryos having Layer Formation blastomeres positioned in four-cell tiers (Fig. 12.1 ; Nielsen 1991 ). Despite variability at Early cleavage among brachiopods is typically the eight-cell stage, the 16-cell stages of both considered to be radial and holoblastic (Zimmer Terebratulina and Terebratalia have the open 1997 ). There is more uniformity among cleavage radial cell arrangement, but irregular 16-cell planes of early stages of brachiopods in lingulid stages are known from Hemithiris (Freeman and craniid species, which produce eight-cell 2003 ). Collectively, early cleavage stages of stages composed of four blastomeres stacked brachiopods share features found in the early directly on top of each other (Fig. 12.1 ). However, cleavage stages of both ectoprocts and phoronids eight-cell stages of rhynchonellid species can be (see Chapters 10 and 11 ). variable in their cell arrangement, producing the Beyond the similarities of cleavage stages, more prevalent stacked arrangement or arrange- there are also some intriguing early developmen- ments in which all the blastomeres are in a ring tal patterns among brachiopods that in some on a single plane (Fig. 12.1; Freeman 2003 ). cases are linked to the morphology of their plank- Variability in the cleavage plane continues, as lin- tonic stages. The fi rst cleavage through the gulid species such as Glottidia and Discinisca anterior-posterior (AP) axis corresponds to the have 16-cell stages with the more prevalent bilat- plane of bilateral symmetry only in species of lin- erally aligned cells while also having some gulids such as Glottidia and Discinisca that have embryos with an open radial arrangement bilateral cleavage (Table 12.1 ). Novocrania and (Freeman 1995 , 1999 ). The 16-cell stages of at least three species of rhynchonellids form the Novocrania anomala have been described as anterior ectoderm of the late gastrula from the having the open radial arrangement (Freeman area around the animal pole (Freeman 2003 ); 12 Brachiopoda 267

Table 12.1 Select developmental processes among six brachiopod species Glottidia Discinisca Novocrania Developmental event pyramidata strigata anomala Rhynchonellidsa First cleavage = plane of bilateral symmetry Yes Yes No No Site of apical lobe specifi cation Lateral Lateral Animal pole Animal pole Stage of animal half specifi cation AL: blastula AL: blastula AL, ML: AL, ML: unfertilized egg blastula-gastrula Stage of vegetal half specifi cation Eight-cell Blastula-gastrula Eight-cell Gastrula Stage of AP axis Specifi cation 16-cell Before cleavage Before cleavage Gastrula Bilateral symmetry is established with AP axis? Yes Yes No Yes Data taken from Freeman (2003 ) Abbreviations : AL apical lobe, ML mantle lobe a Data from three rhynchonellid species

Fig. 12.2 Embryonic germ tissue specifi cation among the mesoderm is shaded red, and the endoderm is shaded various species of brachiopods. All stages shown are late green (All stages redrawn and modifi ed from Freeman gastrulae. Two polar bodies (shaded black) mark the site ( 2003 ) ) of the animal pole. The anterior ectoderm is shaded blue , however Glottidia and Discinisca both form the pore forms the anus (see section on gene expres- anterior ectoderm from the tissue lateral to the sion). The AP axis is specifi ed before cleavage in animal pole (see Fig. 12.2 ). In general, the animal both Discinisca and Novocrania but at the 16-cell half of the embryo is specifi ed during the blastula stage of Glottidia and at the gastrula stage of the to gastrula transition, except in Novocrania rhynchonellids (Table 12.1 ). Furthermore, the AP where this occurs before the egg is fertilized and left-right (LR) axes are specifi ed at the same (Freeman 2000 ). Vegetal half specifi cation is embryonic stage in all but Novocrania . As the more variable, occurring at the eight-cell stage in evolutionary trends discussed here represent Glottidia and Novocrania but later in the devel- information from only six species, more com- opment of Discinisca and three rhynchonellids parative experimental data are required to test the (Freeman 1999 , 2003 ). In all species considered robustness of these developmental patterns. here, the vegetal pole is the initial site of gastrula- The origin and proliferation of mesoderm in tion, and a portion of the blastopore forms the select species of brachiopods have been studied mouth, except in Novocrania where the blasto- mainly through histological preparations of 268 S. Santagata

discrete embryonic stages and some limited cell 1902; Chuang 1977 ; Freeman 1995 , 1999 ). The marking experiments. During gastrulation, the gut of Discinisca larvae is complete with an embryos of Glottidia pyramidata , Lingula ana- esophagus, stomach, intestine, and anus (Chuang tina , and Discinisca strigata all form mesoderm 1977 ). Lingula and Glottidia larvae lack an anus from the anterior end of the archenteron from in the early feeding stages, but a complete gut cells that ingress into the blastocoelic space develops in late planktonic stages transitioning (Yatsu 1902 ; Freeman 1995 , 1999). The embryos into the benthic form (Yatsu 1902 ). Early feeding of Novocrania anomala and Terebratalia trans- stages of Lingula and Glottidia larvae have a versa have multiple archenteric ingression sites semicircular embryonic shell that takes on a cir- along the AP axis (Long and Stricker 1991 ; cular shape in the mid-larval phase (Fig. 12.3A, Nielsen 1991 ). At the very least, mesoderm B ) and elongates into an oblate shape in the late, originates from the endoderm in these brachio- planktonic post-larval phase (Yatsu 1902 ; Chuang pod species. Since the blastopore forms the anus 1977 ; Santagata 2011 ). Discinisca larvae with in Novocrania as well as in other protostome two or three pairs of cirri have not yet developed embryos such as those of the priapulid worm mantle tissue and so are shell-less in the earliest Priapulus caudatus (Martín-Durán et al. 2012 ) feeding stages bearing four different kinds of and the polychaete Owenia collaris (Smart and long embryonic “setae” (Fig. 12.3D ). For reasons Dassow 2009 ), it is possible that both of these discussed in Santagata (2011 ), the latter struc- deuterostomic developmental features were tures will be referred to as “chaetae” (Gustus and shared with the last common ancestor of the Cloney 1972 ; Lüter 2000). Embryonic chaetae of Bilateria (Martín-Durán et al. 2012 ). However, Discinisca are produced by two sacs positioned since mesodermal cell ingression occurs at the laterally adjacent to the gut (Chuang 1977 ). The boundary between the archenteron and the out- long embryonic chaetae are lost (likely shed) in side ectodermal epithelium, it is plausible that later planktonic stages as the circular larval shell some mesodermal cells originate from ectoder- grows, and during this time the mantle tissue mal ingression. Forming mesoderm from dual produces both curved and fl exible chaetae of endodermal and ectodermal sources would lend varying lengths (Fig. 12.3E ; Chuang 1977 ). support to the spiralian affi nities of brachiopods, Although eyespot-like structures have been but detailed cell lineage information would be described for Discinisca larvae (Chuang 1977 ), it needed from brachiopods to test this hypothesis is not clear that photoreception is the function of (see Santagata 2004 for review). these pigmented tissues. Discinisca , Glottidia , and Lingula larvae have a pair of statocysts. In general, the mid-larval fea- LATE DEVELOPMENT tures of Glottidia and Lingula with four to fi ve pairs of cirri and a short pedicle are quite similar Past the late gastrula stage, brachiopod embryos to the late (metamorphic competent) planktonic develop either two or three body regions. Lingulid stages of Discinisca (Nielsen 1991 ). When tran- and discinid brachiopods form the two most ante- sitioning to the benthos, Discinisca larvae extend rior regions fi rst, the apical and mantle lobes. their comparatively short pedicles and attach to Late larval stages of lingulid and discinid bra- hard substrates. However, the late planktonic chiopods are typically considered to be forms of (post-larval) stages of Glottidia and Lingula planktotrophic juveniles (Yatsu 1902 ; Chuang develop the dual-brachial arms of the juvenile 1977 ; Long and Stricker 1991 ; Santagata 2011 ) lophophore, lengthen the muscular pedicle, and but over the complete developmental period extend it beyond the valves for burrowing exhibit different degrees of embryonic, larval, (Yatsu 1902 ; Santagata 2011 ). and precociously developed juvenile traits. The All rhynchonelliform species investigated earliest feeding stages of Lingula anatina , thus far have a trilobed non-feeding larva. Similar Glottidia pyramidata , Discinisca strigata , and morphological features have been observed in other Discinisca spp. have one median tentacle several different rhynchonellid species (Conklin and two pairs of lateral tentacles or cirri (Yatsu 1902; Chuang 1996 ; Pennington et al. 1999 ; 12 Brachiopoda 269

A B C

DE F

Fig. 12.3 Embryonic, larval, and post-larval features of from the plankton on the Pacifi c side of Panama; note the linguliform brachiopods. ( A ) Mid-larval stage of a long embryonic chaetae ( EC ) (Photograph taken by Glottidia larva with seven pairs of cirri collected from the Richard Strathmann. © Richard Strathmann 2015. All plankton of Tampa Bay, Florida (see Santagata 2011 for Rights Reserved). ( E) Late-stage discinid larva with four details). The boundaries of the semicircular embryonic pairs of cirri that has shed the embryonic chaetae and has shell ( ES) are evident next to the circular larval shell. (B ) the full complement of curved mantle chaetae ( CC ) and a Preserved specimen of a post-larval stage of a Glottidia short pedicle ( PD ) (Photograph taken by Richard larva with 11 pairs of cirri collected near St. George Strathmann. © Richard Strathmann 2015. All Rights Island, Florida. Note the elongated and still internal pedi- Reserved). ( F ) Relative size and morphological differ- cle ( PD ). ( C) Relative size and morphological differences ences between the larval ( blue) and post-larval ( white ) among the embryonic (black ), larval (blue ), and post-larval shells of discinid brachiopods (Redrawn and modifi ed (white ) shells of lingulid brachiopods (Redrawn and modi- from Chuang 1977). Other abbreviations: gut ( G ), median fi ed from Chuang 1977 ). (D ) Shell-less stage larva of a tentacle ( MT). Scale bars A and E equal 50 μm and scale discinid brachiopod with three pairs of cirri collected bars B and D equal 100 μm

D’Hondt and Franzén 2001 ; Altenburger and sively from posterior to anterior, leaving only a Wanninger 2009 ), but development is perhaps small opening in the apical lobe that leads into best described in Terebratalia transversa the blind-ended gut. The morphology of the (Stricker and Reed 1985a ; Long and Stricker anterior portion of the apical lobe changes into a 1991 ). During the transition from the late gas- rounded dome that sits on a wider, cylindrically trula to the early trilobed larval stage of shaped base that includes the anterior transverse Terebratalia transversa , the dorsal side of the ciliated band. Late-stage larvae have pigmented embryo fl attens and ventral tissues near the slit- ocelli on the dorsal side of the apical lobe and like blastopore move toward the midline, curving vesicular bodies that border the posterior margin inwardly (see Fig. 12.4 and Santagata et al. 2012 of the apical lobe. The asymmetric mantle lobe for review). The three body regions of the early extends further on its ventral side and partially trilobed larva develop (apical, mantle, and pedi- covers the pedicle lobe. Other features of the cle lobes), and the blastopore closes progres- mantle lobe are the paired dorsal and medial 270 S. Santagata

A BC D

E FG

HI J K

Fig. 12.4 Larval development of Terebratalia transversa of the mantle lobe. (G ) Volume projection of a competent ( A – G ; micrographs by S. Santagata) and Novocrania larva in which the vesicular bodies and some secretory anomala ( H – K ; scanning electron micrographs provided cells in the pedicle lobe are nonspecifi cally stained ( blue ). by C. Nielsen). (A ) Early trilobed larval stage in which the ( H) Late gastrula stage of Novocrania . The blastopore boundaries of the apical ( AL), mantle ( ML), and pedicle ( BP ) will become the anus. (© Claus Nielsen, 2015. All ( PL ) lobes are being formed. ( B ) The mantle lobe enlarges Rights Reserved). ( I ) A ventral view of a late larval stage as the chaetal sacs with short chaetae develop. The slit- with uniform and simple ciliation. The rounded apical like blastopore is closing from posterior to anterior. (C ) lobe ( AL) sits on top of a more posterior secondary lobe As the apical lobe takes on a dome-shaped appearance, ( SL ) that may be homologous to only the mantle lobe of only a small portion of the blastopore ( BP ) remains. other brachiopod larvae or to both the mantle and pedicle Vesicular bodies (VB ) mark the boundary between the api- lobes (© Claus Nielsen, 2015. All Rights Reserved). ( J ) cal and mantle lobes. ( D) Later in larval development, the Dorsal view of the specialized epithelial region centrally chaetae lengthen and a row of red-pigmented ocelli ( OE ) located between the three pairs of chaetal bundles that will develops on the dorsal surface of the apical lobe. (E ) Late- secrete the dorsal valve ( DV ) of the juvenile shell at meta- stage larvae have a transverse ciliated band ( CB) at the morphosis (© Claus Nielsen, 2015. All Rights Reserved). base of the apical lobe, and near the time of metamorphic ( K ) Lateral view of a metamorphic competent larva competence, the dorsal and medial chaetae ( DC ) grow (© Claus Nielsen 2015. All Rights Reserved). All scale beyond the length of the pedicle lobe. ( F) A mid-ventral bars equal 50 μm band of cilia ( VC ) develops on the larger ventral extension 12 Brachiopoda 271 chaetal sacs and a mid-ventral band of cilia. Near boundaries of the periostracal layer are evident, the time of metamorphic competence, the poste- in contrast to the more elongate form of subse- rior pedicle lobe narrows and divides into muscu- quently shaped valves (Fig. 12.5 ) that open and lar and glandular portions (Fig. 12.4 ). close through the actions of the abductor and As discussed previously, the remaining por- adductor muscles (Santagata 2011 ). tion of the blastopore in the late gastrula of Different interpretations exist with respect to the Novocrania anomala is found at the posterior end metamorphosis of Novocrania anomala . According of the embryo and becomes the anus (see to Nielsen ( 1991), competent larvae use ventral Fig. 12.4 ; Nielsen 1991 ; Freeman 2000 ). Larval musculature in the mantle (posterior) lobe to bend development results in two body regions, a more the larval body. As a result, both the anterior por- rounded apical lobe and an elongate posterior tion of the apical lobe and a posterior region of the lobe that bears three pairs of chaetal bundles on mantle (posterior) lobe are in contact with the sub- the dorsal surface (Fig. 12.4 ). Late-stage strate and with their cellular secretions serve as the Novocrania anomala larvae have a specialized attachment area. Over time the larval body fl attens circular epithelium centrally located between the and the dorsal shell fi eld expands, pushing the dorsal chaetal bundles that secretes the dorsal chaetal bundles to the periphery of the juvenile valve at metamorphosis (Nielsen 1991 ). Since body (see Fig. 12.5 ). Altenburger et al. (2013 ) dis- adult craniiform brachiopods lack a pedicle and agreed with these interpretations based on the fol- cement directly to the substrate at metamorpho- lowing reasons: (i) Initial larval attachment to the sis, the posterior larval lobe that bears chaetae substrate was achieved by the posterior tip of the has typically been interpreted as the mantle lobe mantle (posterior) lobe similar to other rhynchonel- (Nielsen 1991 ). However, this interpretation has lid larvae (Stricker and Reed 1985a ), and (ii) the been challenged, as cell types of both the mantle ventral attachment area included a cuticle but nei- and perhaps the pedicle lobes may be present in ther mantle tissue nor a ventral valve until 17 days the posterior larval lobe of Novocrania postmetamorphosis. Further implications of (Altenburger et al. 2013). Ciliation on the apical Nielsen’s interpretations pertain to the features of lobe is uniform and simple (Nielsen 1991 ), and the ancestral brachiopod in which both the dorsal although a pair of anterolateral red-pigmented and ventral valves at metamorphosis are derived spots has been observed on the apical lobe, it is from different anterior and posterior regions of the not known if these structures are ocelli. larval dorsal epithelium (Nielsen 1991 ). Some Metamorphosis of rhynchonelliform bra- aspects of this hypothesis are based on the develop- chiopods has been most closely studied in ment of Lingula anatina , in which the initial Terebratalia transversa (Long and Stricker embryonic shell fi eld is a circular domain that is 1991). Initially, the competent larva attaches to split and folded during development to make two the substrate using secretions produced by the valves (Yatsu 1902 ). Subsequently called the posterior tip of the pedicle lobe (Stricker and “Brachiopod Fold Hypothesis” (Cohen et al. 2003 ), Reed 1985a ). Once attached, the edges of the this concept has been used to explain body plan dif- mantle lobe are extended over the apical lobe ferences among fossil and living brachiopods and through the actions of mantle and pedicle mus- phoronids. Whether the metamorphosis of culature (Stricker and Reed 1985c ; Altenburger Novocrania anomala supports or refutes this and Wanninger 2009 ; Santagata 2011 ). In com- hypothesis will require more information regarding petent larvae, a periostracum layer is secreted the development and metamorphic modifi cations underneath the mantle epithelium and on top of of the axial properties (anterior-posterior and the anterior portion of the pedicle. After meta- dorsoventral) of larval and juvenile brachiopod morphosis, this layer covers the mantle tissue bodies. Gene expression studies that include exper- and when calcifi ed forms the dorsal and ventral imental perturbations of embryonic axes (see valves of the juvenile shell (Stricker and Reed Röttinger and Martindale 2011 ) should help resolve 1985b ). In young juvenile stages, the circular some of these questions. 272 S. Santagata

A B

CD

Fig. 12.5 Juvenile morphology of Terebratalia trans- muscles ( ABD ). (C ) Early stage of metamorphosis of versa (A , B : myoanatomy; Santagata 2011 ) and Novocrania in which the dorsal valve fi eld ( DV ) has Novocrania anomala (C , D: gross morphology; Nielsen expanded, pushing the chaetal bundles (CH ) to the periph- 1991 ). ( A ) Juvenile with six tentacles; note the boundaries ery (© Claus Nielsen, 2015. All Rights Reserved). ( D ) of the periostracal layer (PS ) and subsequent shell growth Three-day-old juvenile showing the mineralized dorsal that lacks this appearance. PD pedicle. (B ) Juvenile with valve. Scanning electron micrographs in C and D pro- eight tentacles, each of which has intertwined striated vided by C. Nielsen (© Claus Nielsen, 2015. All Rights muscles ( TM ) also showing the large paired abductor Reserved). All scale bars equal 50 μm

GENE EXPRESSION the apical lobe (a more dorsal apical organ and a second ventral cell cluster), with a third neu- Although no current gene expression studies ral domain located mid-ventrally on the mantle have investigated the specifi cation of embry- lobe (Fig. 12.6A). Additional details about the onic axes in brachiopods, there have been sev- histaminergic cells of the larval nervous system eral recent accounts of the differentiation of the of the competent larva are described in Santagata larval nervous system and sense organs of the ( 2011 ), but in general the larva has a compara- rhynchonellid Terebratalia transversa. Late- tively broad apical organ that contains numer- stage trilobed larval stages have a basiepithe- ous monociliated sensory neurons consisting of lial nervous system with at least three distinct at least two morphological types that send axo- neural cell domains, two of which are found in nal fi bers into a central neuropil (Fig. 12.6B ). A 12 Brachiopoda 273

A B CDE

F GHIJ

K

Fig. 12.6 Gene expression patterns in the larval nervous All Rights Reserved). ( E) Expression of c-opsin in the system and sense organs of Terebratalia transversa . ( A ) dorsal larval ocelli of a mid-trilobed larval stage Lateral view of the histaminergic nervous system (yellow ) (Passamaneck et al. 2011 ) (© Yale Passamaneck, 2015. of a late trilobed stage larva showing the broad neuronal All Rights Reserved). ( F) Expression of otp in a group of cell domains in the apical organ (AO ), a ventral cell cluster central fl ask-shaped cells in the developing apical organ in the apical lobe ( AVO ), and a mid-ventral band of hista- of a late gastrula stage. (G – K) Expression of several minergic cells on the mantle lobe ( MVC ). Phalloidin neural-related genes in the trilobed larval stage; views are ( blue) was used as a general cell marker. DC dorsal chae- ventral, dorsal, and lateral, showing the remaining portion tae, MC medial chaetae, PD pedicle lobe. ( B) Frontal opti- of the blastoporal opening ( M), the apical tuft (AT ), and cal section through the apical organ that contains a central the region of the transverse ciliated band ( CB) on the api- group of sensory neurons (SN1 and SN2) as well as other cal lobe ( AL). All of these genes are expressed in overlap- ciliated cells that are labeled with acetylated α-tubulin ping ectodermal domains in the apical lobe, a few are ( yellow). Nuclei are stained blue. NP neuropil. ( C , D ) expressed in the anterior endoderm (*), but only one gene, Transmission electron micrograph of a larval ocellus and otp , is expressed in a subset of cells in the mid-ventral a composite drawing of the lens ( LS ) and pigmented gran- band of cilia on the mantle lobe (ML ). None of these genes ules ( PG) inside the photoreceptive cells, both of which were expressed in the pedicle lobe ( PD ) at this stage have folded ciliary membranes (CM ) (Micrograph and (Santagata et al. 2012 ). Scale bar in (A ) equals 50 μm, drawing by N. Furchheim and C. Lüter, from Passamaneck scale bars in (B ) and (E – J ) equal 25 μm, and scale bars in et al. (2011 ). © Nina Furchheim and Carsten Lüter, 2015. ( C and D ) equal 10 μm 274 S. Santagata subset of these sensory neurons within the apical bilaterian animals express Six3/6 , Homeobrain , organ can be labeled with acetylated α-tubulin and NK2.1 (see, e.g., Meyer and Seaver 2009 ; (Santagata 2011 ) and serotonin (Altenburger Steinmetz et al. 2010; Ferrier 2012), the cytoar- et al. 2011 ). Other sensory cells located near the chitecture of larval apical organs is both struc- apical organ are the red-pigmented ocelli on the turally and neurochemically diverse (Byrne dorsal side of the apical lobe. Each ocellus con- et al. 2007 ; Wanninger 2009 ; Santagata 2011 ). sists of a lens cell and a shielding pigment cell Genes involved in the specifi cation of ante- (Fig. 12.6C, D ), and the cilia of both photorecep- rior neuroectoderm (Six3/6 ) and the apical tuft tive cells have folded and enlarged membranes ( FoxQ2 ) are expressed in similar larval domains (Passamaneck et al. 2011). The onset of phototac- in brachiopods and other bilaterians (Santagata tic behavior, however, precedes the development et al. 2012 ). At the late gastrula stage, a cen- of larval ocelli, as it has been demonstrated that tral group of otp-positive cells in the develop- the asymmetric middle gastrula stage (those in ing larval apical organ (see Fig. 12.6F ) bears a which the anterior end has been specifi ed) exhib- striking resemblance to the fl ask-shaped, sero- its a positive phototactic response (Passamaneck tonergic sensory neurons in the apical organs et al. 2011 ). This stage also expresses both cili- of several spiralian larval types (Wanninger ary and Go-class opsins in an anterior medial 2009 ). However, as discussed in the previous ectodermal domain as well as other genes section, the fully developed larval apical organ related to the regulation of phototransduction of Terebratalia transversa is a relatively wide (Passamaneck and Martindale 2013 ). Similar to neuronal cell domain with several structural and the gene expression patterns during the differen- neurochemical differences from the larval api- tiation of cerebral eyes in other protostome and cal organs of other spiralians and ambulacral deuterostome animals (Hill et al. 1991 ; Arendt deuterostomes (Santagata 2011 ). Interestingly, et al. 2002 ), early trilobed larval stages express the proliferation of neuronal cells in the api- both Otx and Pax 6 in bilaterally symmetrical cal organ of Terebratalia transversa during bands in the apical lobe where the ocelli even- larval development corresponds to broadened tually develop. By the mid-trilobed larval stage, expression domains of FoxQ2 , otp, hbn, and the expression of c-opsin is mainly limited to the NK2.1 (Fig. 12.6F–K ; see Santagata et al. 2012 ) dorsal ocelli (Fig. 12.6E ). Since the photorecep- relative to their more restricted expression tor cells of larval cerebral eyes of related spira- domains in the larval apical organs of spiralians lian animals are rhabdomeric rather than ciliary (Nederbragt et al. 2002 ; Fröbius and Seaver (Arendt et al. 2002 ), it is unclear whether these 2006 ; Tessmar-Raible et al. 2007 ). photoreceptive cell types are homologous or have evolved independently (Passamaneck et al. 2011 ). However, as discussed in Passamaneck OPEN QUESTIONS et al. (2011 ), diverse members of protostome and deuterostome animals have larval cerebral • What were the larval and adult characteristics of eyes with both ciliary and rhabdomeric photore- the stem and crown lineages of brachiopods? ceptive cells, and thus it is plausible that ciliary • What is the phylogenetic position of various photoreceptors are plesiomorphic for the Spiralia clades of extant brachiopods relative to the and perhaps the Bilateria. phoronids, ectoprocts, and related spiralians? Broadscale homology has been proposed • Can any evolutionary inferences be drawn from for other aspects of the larval nervous system, the variation in the blastomere arrangements of in particular the larval apical organ (Tessmar- early cleavage stages of brachiopods? Raible 2007 ) and subsets of centrally located • How are the embryonic axes specifi ed and cells responsible for making the long apical cili- modifi ed in the various clades of brachiopods? ary tuft (Yaguchi et al. 2010 ). Although anterior • How are the neurogenic domains specifi ed in regions of larval neuroectoderm from divergent linguliform and craniiform brachiopods? 12 Brachiopoda 275

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A Apatite , 264 Abdomen , 219 Apical cap , 112, 123 Abdominal-A , 219 Apical lobe , 267, 269–274 Abboral , 63, 250, 252–256 Apical organ , 36, 38, 80, 84, 90, 96–100, 109, 111, 116, Acanthella , 9–11 117, 119–121, 123, 124, 126–128, 130, 131, Acanthocephala , 2, 3, 9 140–143, 146, 168, 175, 179, 188, 200, 202, Acanthocephalan , 7, 9, 11 207, 211–213, 237 Acanthor , 9, 11 Apical rosette , 92, 94–96 Acoela , 50 Apical sensory organ , 122 Acoelomate , 14, 18, 80, 90 Apical tuft , 98, 112, 117, 119, 120, 123, 130, 165, Acrosome , 234 170–172, 174–176, 186, 202–208, 213, 214, Actin , 83, 94, 120, 132, 180, 199, 235, 251, 254 237, 238, 273, 274 Actinotroch , 235, 239, 240, 242 Aplacophora , 139 Aculifera , 109, 133, 136, 139, 142, 144, 146 Aplacophoran , 95, 109, 123, 131, 133, 134, 136 Adultation , 256 Apomorphic , 156–159 Adult shell , 123, 135, 136, 138 Apomorphy , 249 Agassiz , 212, 213 Appendage , 15, 146, 194 Amnion , 173, 174 Aquaculture , 106, 107 Amphistomy , 201 Archenteron , 167, 174, 268 Ampulla , 242 Architomy , 45 Ampullary , 97, 130, 131, 232 Archoophora , 22–24, 44 Anasca , 248, 250, 255 Arm , 120, 141, 142, 264 Anatomy , 30, 120, 131, 132, 156–159, 212, 239, 253, Arthropod , 11, 22, 124, 216–218, 220 256, 257 Articulata , 216 Ancestral , 18, 34–38, 43, 57, 63, 65, 109, 133, 142, 166, Ascophora , 248, 249, 255 171, 184, 198, 207, 208, 212, 215, 217, 218, Asexual , 23, 33, 34, 43, 45, 48, 57, 80, 81, 84–86, 220, 233, 252, 264, 271 254, 258 Animal-vegetal axis , 15, 25, 33, 36, 162, 165, 200, 234 Asexual reproduction , 33, 45, 48, 57, Annelid , 3, 94, 95, 109, 185, 194, 197–204, 207, 208, 99–100, 197 212, 215–218, 220, 222, 223, 241 Asymmetric cell division , 36 Annelida , 42, 163, 175, 193–223 Asymmetry , 106, 107, 144, 200 Annelid cross , 3, 95, 200 Asynchronous , 33, 34, 92 Antalis , 104, 112, 113, 117–119, 121, 130–132, 136, 138, 141 Atria , 90, 156 Antenna , 194, 206, 220 Atrium , 90, 99 Anterior ectoderm , 17, 237, 238, 266, 267 Attachment disc , 80, 82, 85, 90 Anterior endoderm , 273 Autozooid , 248, 253 Anterior-posterior (AP) axis , 5, 26, 29, 30, 36, 44, 48, Axial mesoderm , 127, 128 53, 61–66, 96, 113, 124, 125, 131, 133, 142, Axial organ , 36, 62 161, 165, 171, 174, 194, 213, 216, 217, 220, Axis , 5, 9, 15, 17, 23, 25, 30, 33, 36, 44, 46, 51, 53, 237, 255, 266–268, 271 60–66, 96, 113, 120, 122, 123, 125, 131, 133, Antibodies , 30, 80, 119, 124, 131, 176, 187 142, 161, 162, 165, 166, 168, 171, 174, 186, Antp , 125, 126, 128, 140, 142, 218 194, 199–201, 216, 217, 220, 234, 237, 250, Anus , 2, 23, 90, 91, 96, 99, 122, 123, 125, 127, 139, 156, 255, 266–268 160, 177, 181, 182, 194, 201, 205, 215, 232, Axis elongation , 217 237, 238, 248, 253, 264, 267, 268, 270, 271 Axon guidance , 56, 65

A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia) 279 DOI 10.1007/978-3-7091-1871-9, © Springer-Verlag Wien 2015 280 Index

B Buccal , 80, 82–85, 97, 123, 124, 126, 128, 131, 132 Barentsia , 91, 92, 97 Buccal cavity , 80 Basal , 23, 34, 50, 58, 90, 91, 96, 109, 111, 112, 122, 130, Bud , 68, 99, 179, 180, 201, 238, 248, 249, 252, 254 135, 138, 139, 142, 175, 177, 185, 186, 194, Budding , 80, 99, 100, 137, 249, 258 197, 222, 223, 240, 242, 254 Bugula , 248, 250, 253–255, 258, 259 Basic helix-loop-helix (bHLH) , 56 Burrowing , 104, 111, 264, 268 Bdellonemertea , 156, 157 Beta (β)-catenin, 30, 36, 45, 53, 61–64, 66, 128, 185, 186, 200, 220 C Bilaterality , 2, 4, 14, 30, 49, 53, 60, 61, 116, 139, 169, Caenorhabditis , 53, 64, 198, 222 173, 177, 183, 184, 188, 200, 201, 210, 211, Caenorhabditis elegans , 53, 64, 198, 210, 216 250, 254, 266, 267, 274 Calyx , 90, 99 Bilateral symmetry , 61, 169, 201, 254, 266, 267 Cambrian , 105, 194, 233, 248, 264 Bilateria , 50, 144, 198, 222, 268, 274 Canonical Wntá , 36, 63 Bilaterian , 80, 142, 143, 145, 218, 220, 222, 249, 274 Cap , 112, 114, 122, 123, 179, 180, 201, 202, 216 Bivalvia , 104, 106, 111, 123, 125, 146 Capitella , 201, 210, 215–222 Blastema , 25, 27, 45–48, 53, 56, 57, 60 Catastrophic metamorphosis , 99, 169, 174 Blastemal cells , 250, 252–259 Catecholamine , 50, 241 Blastocoel , 94, 164, 167, 168, 185, 201, 237, 238, Catenulida , 22, 24, 34, 38, 44, 50, 52, 59 252, 268 Caudal , 15, 17, 50, 139, 170, 171, 182, 215 Blastomere , 3–7, 9, 15–17, 23, 25–30, 33–38, 63, 84, 92, Caudofoveata , 104 96, 107, 113, 114, 116–118, 122, 124, 125, Cdx , 126, 140, 185–187, 220, 222, 258 144, 145, 161–169, 177, 186, 199, 200, Cell , 3, 14, 23, 42, 80, 90, 107, 156, 197, 232, 248, 266 234–236, 250, 251, 266, 274 Cell adhesion , 53, 55, 64, 67 Blastoporal , 169, 171, 174, 273 Cell division , 3–5, 9, 15, 17, 25, 27, 34, 36, 92, 107, 199, Blastopore , 5, 7, 16, 17, 96, 121, 122, 125–128, 139, 200, 210, 217, 236, 237, 251 171, 172, 174, 181, 201, 202, 222, 237, Cell internalization site , 5, 250 267–271 Cell labeling , 237 Blastula , 94, 106, 144, 168, 169, 237, 238, 252, 267 Cell lineage , 9, 11, 18, 36, 60, 92, 96, 100, 107, 113, Blood , 33, 156, 158, 159, 177, 182, 185, 188, 232, 233, 115, 116, 121, 124, 130, 138, 139, 146, 238–242 166–169, 237, 250, 252, 258, 268 Blood cell , 232, 239 Cell type , 18, 28, 30, 33, 37, 42, 43, 47, 48, 51, 121, 167, Blood corpuscle , 238–242 184, 197, 220, 239, 253, 271, 274 Blood vessel , 33, 158, 159, 185, 232, 233, 239, 242 Cement , 264, 271 BMP , 23, 29, 30, 60–66, 127, 128 Central nervous system (CNS) , 5–23, 28, 49–51, 57, 62, Bmp2 , 127, 188 65, 66, 130, 131, 141, 142, 215, 220, 221 Bmp2/4 , 127, 188 Centrosome , 35 Body plan , 24, 29, 30, 35, 37, 38, 80, 91, 99, 100, 104, Cephalic appendage , 146 109, 123, 135, 143, 158, 166, 176–177, 194, Cephalic ganglia , 44, 45, 49, 51, 53–58, 62 218, 258, 271, 275 Cephalopoda , 104, 107, 108, 128 Body wall , 31, 37, 80, 84, 90, 97, 133, 134, 156–158, Cerebral , 5, 7, 44, 59, 84, 90, 116, 117, 119, 120, 124, 177, 179, 185, 206, 212, 232, 237, 238, 256 126, 130, 131, 140–142, 158, 171, 173, 175, Bone morphogenetic protein , 64 183, 184, 186, 187, 194, 221, 274 Bowerbankia , 248, 250, 253, 254 Cerebral orga , 158, 171, 173–175, 183, 184, 186, 187 Brachidia , 264, 265 Chaetae , 194, 203, 204, 206, 207, 268–271, 273 Brachiopod , 233, 264, 265, 267–271 Chaetodermomorpha , 104, 109, 124, 144, 146 Brachiopoda , 175, 194, 263–274 Chaetopleura , 113, 117, 118, 121 Brachiopod fold hypothesis , 271 Chaetopterus , 196, 198, 199, 202, 203, 211, 215, 216, Brachyury , 125, 128, 139, 222 218, 220, 222 Brain , 2, 4, 5, 9, 14, 17, 18, 26, 28–30, 32, 33, 37, 38, 44, Change , 5, 23, 35, 36, 45, 69, 90, 106, 107, 49–51, 53, 55–59, 61, 62, 84, 105, 108, 132, 109, 123, 138, 141, 169, 187, 207, 236, 142, 156–159, 175–178, 183, 184, 186–188, 239, 269 194, 195, 213, 217, 220–222, 233 Character state , 109 Branching , 22, 49, 50, 173, 175, 197, 222, 223 Cheilostomata , 248 BrdU , 48, 210 Chirality , 113, 143–145, 237 Brood , 90, 96, 135, 197, 233, 235, 238, 250, 252, 265 Chitin , 264 Brood care , 107, 112 Cholinergic , 50, 56 Brood chamber , 138 Chordate , 22, 64, 65 Brooding , 135, 197, 238, 250 Chordoid , 80, 82, 84–86 Bryozoa , 90, 232, 248 Chorion , 108, 160 Index 281

Cilia , 5, 14, 15, 32, 33, 59, 80, 96, 112, 119, 120, Cribimorpha , 248 122–124, 130, 166–168, 171, 172, 174, 186, Crisia , 249, 252, 256 202, 203, 205, 209, 238, 239, 241, 251, 270, Cross furrow , 162, 163, 166, 236, 237 271, 273, 274 Crustacea , 9, 85, 107, 108, 159 Ciliary band , 5, 112, 125, 172, 173, 202–205, 207, Crustacean , 9, 85, 107, 108, 159 209, 238 Cryptic species , 232, 265 Ciliary photoreceptor , 274 Ctenostomata , 248 Cilium , 212 Cut , 46–48, 109 Circulatory system , 51, 90, 242 Cuticle , 14, 80, 82–84, 90, 271 Cirri , 96, 97, 109, 171, 185, 194, 199, 202, 203, 268, 269 Cycliophora , 79–86, 91 Cirrus , 171, 206 Cycliophoran , 80, 82, 84–86 Cleavage , 3, 15, 23, 66, 84, 90, 106, 161, 199, 234, Cyclostomata , 249 249, 266 Cyphonautes , 250, 253, 257 Cleavage pattern , 3, 11, 15–17, 23–27, 32, 34, 38, 85, 92, Cyst , 82, 84 113, 114, 144, 163, 165, 199, 200, 234, 236, Cystacanth , 11 237, 242, 249 Cystid , 248, 252–254, 256–259 Cleavage program , 3, 116 Cytoplasm , 34, 42, 61, 67, 68, 113, 114, 122, 161–163, Cleave , 15, 25, 33, 84, 112, 122, 236 198, 199, 212, 250 Cleaving , 3, 5, 36, 113, 114, 163, 200, 206, 236 Clonal , 48, 80, 100, 165, 166, 187, 211 Cnidaria , 146 D Cnidarian , 61, 63, 163, 169, 194, 248 4d , 24–26, 35, 37, 92, 96, 113, 114, 116, 118, 122, 165, Cocoon , 28, 30, 197, 200 167, 200, 201, 210, 236 Coeloblastula , 95, 121, 168 Decapentaplegic (Dpp) , 28, 64, 125, 127, 143 Coelom , 4, 23, 90, 234, 236, 242, 248 Decapod , 124, 136, 139, 141 Coelomate , 180, 194 Decidula , 169, 177 Coelomic , 90, 185, 194, 215, 232, 234, 237, 242, Deformed , 145 258, 265 Delamination , 167, 173, 174, 178, 180 Coelomic cavity , 185, 194, 215, 232, 237, 265 Delay , 57, 166 Coelomic lining , 215, 242, 265 Delta , 221 Coleoid , 138, 139, 142 Delta-Notch pathway , 216 Collar , 233, 234, 237–242 Dendrobeania , 254 Collier , 55, 56, 142 Desor , 169, 174, 175, 178–180, 183 Collinearity , 218, 220 Determinant , 36, 144, 165, 166 Colonial , 90–92, 99, 105, 249 Deuterostome , 50, 65, 143, 144, 163, 222, 237, 239, Coloniales , 90 249, 274 Coloniality , 105 Deuterostomy , 201, 223 Colony , 32, 91, 248, 249 Deutocerebrum , 142 Column , 91, 93, 96, 122, 123, 134, 168, 172, 173, 250 Development , 2, 15, 23, 42, 80, 90, 105, 161, 197, 232, Commissure , 14, 18, 49, 51, 54, 97, 98, 120, 124, 126, 250, 265 130, 131, 194, 195, 215 Developmental biology , 23, 54, 61, 66, 85, 105, 111, Competent , 34, 43, 96, 97, 99, 128, 131, 233, 238–242, 144, 198 251, 253, 268, 270–272 Developmental gene , 11, 15, 100, 125, 146, 223, Compound cilia , 80, 96, 119, 122, 130, 202 242, 258 Conchifera , 109, 111, 123, 124, 130, 131, 133–139, 142, Dexiotropic , 3, 163 143, 146 Dfd , 218 Conklin, E.G. , 5, 113, 138, 200, 268 Differentiation , 5, 6, 15, 17, 18, 28, 33, 37, 43, 45–48, Connective , 194, 195, 213, 264 55, 56, 60, 64, 68, 122, 167, 182, 215, 221, Conus , 105 239, 258, 272, 274 Convergence , 57 Digestive system , 9, 32, 120, 212, 220 Co-option , 218 Digestive tract , 2, 5, 6, 9, 16, 18, 32, 90, 99, 116, 142, Cord , 28, 50, 56, 125, 156, 187, 194, 213, 215, 217, 220, 145, 222 221, 241 Dioecious , 14, 111, 112 Corona , 4, 250, 253 Dioecy , 112 Coronal cells , 250, 251, 253–257 Diplosoma , 160 Coronate , 253–255 Direct development , 4, 24, 34, 37, 112, 169, 207 Cortex , 161, 198 Discinisca , 264–268 Craniiformea , 264 Discoidal cleavage , 113, 114 Creeping-type larva , 90, 96–98 Disko Island , 3 Crepidula , 107, 113, 114, 116–118, 121, 124, 130 Distal-less , 60 282 Index

Division , 3–5, 9, 15, 17, 22, 24, 25, 27, 28, 32, 34, 36, 92, Endomesoderm , 35–37, 63, 96, 114, 118, 122, 167, 107, 139, 161–167, 186, 199, 200, 204, 210, 168, 188 211, 217, 232, 235–237, 248, 249, 251, 253 Endoplasm , 198, 199 Dlx , 59–60, 221 Endoplasmic reticulum (ER) , 161 Dorsal ectoderm , 5, 8 Engrailed , 125, 127, 142, 143, 188, 216–218 Dorsal pharyngeal muscle , 30 Enterocyte , 68 Dorsal valve , 264, 270–272 Entoproct , 80, 89–100, 109, 130, 201, 202, 212, 248, 249 Dorso-ventral (DV) axis , 26, 30, 36, 61, 63–66, 162, 165, Entoprocta , 80, 89–100, 212 166, 168, 201, 220, 270, 272 Epiboly , 24–27, 121, 202 Drosophila , 53, 59, 64, 65, 67, 68, 142, 187, 198, 216, Epidermis , 2, 3, 5, 9, 18, 23, 25–33, 35, 37, 47, 90, 166, 217, 220, 222 167, 169, 172–175, 177–184, 237, 239, Dwarf male , 4, 82, 84, 197 255, 256 Epimorphic , 45, 46 Epimorphosis , 45 E Epipod , 127 Eccentrotheca , 233, 264 Episphere , 96, 111, 122, 141, 202, 207 Ecdysozoa , 220, 222 Epistome , 232, 242 Echinoderm , 61, 143, 194, 232 Epithelial cell , 47, 68, 184 Echiura , 194, 200, 202, 203, 212, 213, 215 Epithelial lining , 232, 242 Ectoderm , 4, 5, 8, 14, 17, 25, 26, 30, 35, 65, 92, 96, 99, Epithelium , 34, 138, 143, 177, 179, 180, 215, 221, 232, 114, 116, 117, 121, 122, 124–128, 136–139, 233, 237, 239, 241, 252–259, 268, 271 158, 165, 167, 169, 171–173, 177–179, Equal cleavage , 199 181–183, 200, 201, 210, 211, 216–218, Equal division , 210 220–222, 237, 238, 250–252, 258, 266–268, Errantia , 194 273, 274 Esophageal , 121, 125–127, 132, 140–142, 171–173, 178, Ectomesoderm , 35, 37, 114, 116, 117, 121, 122, 167, 181, 194, 213, 253 168, 201 Esophagus , 80, 90, 117, 129, 156, 158, 165, 167, Ectoplasm , 198–199 171–174, 177, 178, 180–183, 242, 268 Ectoproct , 248–250, 252, 256–258 Euprymna , 124, 128, 129, 132, 141, 142 Ectoprocta , 80, 90, 91, 247–259 Eve , 216, 217 Ectoteloblast , 201, 210 Even skipped , 224, 229 Ediacaran , 233, 264 EvoDevo , 38, 106, 146, 222, 111143 EGFR , 53 Evolution , 3, 24, 26, 35, 36, 38, 43, 50, 66, 69, 104, 109, Egg , 3, 4, 6, 9, 11, 14–17, 22–28, 31–34, 106–108, 113, 133, 143, 144, 146, 169, 185, 198, 208, 218, 121, 137, 138, 159–163, 165, 169, 174, 175, 222, 249 198–200, 204, 207–210, 233, 234, 236–238, Evolutionary developmental biology , 23, 105, 111, 265, 267 144, 198 Egg shell , 6, 15, 16, 25, 28, 34 Evolutionary novelties , 111 Elav , 128, 129 Evolution of segmentation , 198, 218 Embryo , 3–11, 15, 17–18, 24–26, 28, 30, 32, 34–38, 63, Excretory system , 5, 14, 68, 116, 123 64, 85, 95, 113, 114, 122, 143, 144, 161, Extracellular matrix , 172, 177, 232 163–166, 168–170, 199, 200, 204, 206, 210, Exumbrella , 171–173, 237 216, 234–237, 250–254, 267, 269, 271 Eya , 59, 60 Embryogenesis , 26, 28, 30, 32, 34, 35, 37, 38, 61, 63, 66, Eye , 28, 30, 47, 53, 56–61, 117, 121, 132, 141, 165, 168, 69, 84, 92, 107, 236, 257 170, 171, 184, 186, 197, 204, 206, 221 Embryology , 6, 23, 24, 33, 38, 85, 86, 91, 93, 113, 139, 250 Eyeless , 60 Embryonic , 11, 23–30, 32–37, 42, 48, 56, 61, 63, 66, 67, Eye morphogenesis , 57 100, 107, 112, 122, 123, 132, 135, 136, 138, Eye spot , 58, 184, 197 139, 144, 146, 161, 163, 164, 166, 181, 185, 188, 207, 235, 236, 249, 252, 267–269, 271–272, 274 F Embryonic shell , 112, 123, 135, 136, 138, 139, 268, Fate map , 8, 18, 237 269, 271 Feeding , 28, 30, 35, 37, 80–86, 122, 123, 174, 175, 202, Endocrine , 51 205, 207–209, 222, 232, 233, 239, 248–255, Endoderm , 17, 26, 27, 30, 35, 96, 114, 116, 118, 125, 257, 258, 264, 265, 268 127, 129, 177, 181, 186, 201, 237, 238, 258, Feeding stage , 80–85, 205, 208, 253, 265, 268 267, 268, 273 Fertilization , 2, 14, 23, 84, 85, 91, 106, 107, 111, 113, Endodermal , 5, 7, 15, 17, 92, 99, 125, 139, 171, 222, 160, 161, 165, 182, 197–199, 234, 236, 250, 252, 258, 268 251, 252 Index 283

Fertilization envelope , 251 Germband , 28, 37, 201 Filter-feeding , 233 Germ cell , 8, 42, 118, 201 Fin , 139 Germinal vesicle breakdown (GVBD) , 161 Fission , 23, 33, 42, 45, 48 Germ layer , 30, 63, 114, 116, 127, 158, 185, 266 Flagellum , 160 Germ line , 8, 42, 116, 198 Flask cell , 98, 117, 119, 130, 131 Germovitellarium , 4–6, 8 Flask-shaped , 96, 97, 116, 119, 120, 130, 175, 176, 212, Giant neuron , 131 241, 273, 274 Glottidia , 264–269 Flatworm , 22–24, 26, 27, 29, 31–35, 37, 38, 42, 44–51, Gonad , 4, 14, 161 53, 56, 95, 111 Gonoduct , 265 Follicle cell , 113 Gradient , 61, 63, 124, 133, 213 Foot , 90, 91, 96–99, 109, 111, 112, 117, 118, 120, 121, Gradual , 62, 63, 156, 169, 171 123, 125–127, 131, 132, 135, 140–142, 264 Grid , 30, 37, 133 Foot gland , 96, 99 Ground pattern , 194, 195, 197, 201, 212, 222 Foregut , 127, 139, 156, 181, 185, 188, 201, 222 Growth , 5, 8, 23, 41–69, 99, 107, 109, 122, 133, 136, Forkhead , 139 169, 172, 181, 194, 204, 207, 210, 211, 213, Fossil , 14, 133, 136, 139, 233, 248, 264, 265, 271 215–217, 219, 232, 249, 251, 272 FoxA , 30, 35, 63, 127, 222, 258 Growth zone , 109, 133, 194, 204, 207, 210, 211, 213, FoxQ2 , 60, 212–213, 242, 274 216, 217, 219 Free-swimming , 26, 59, 80, 84, 105, 205, 207, 215 Gut , 2–4, 7, 9, 14, 15, 18, 23, 26, 28, 30–32, 48, 68, 80, Frontal organ , 96–99, 159, 183, 184, 186, 187, 240–242 83, 96, 116, 118, 126, 143, 156, 165, 172, 173, Fruit fl y , 66 179–181, 186, 208, 215, 216, 222, 232, 237, Funnel , 80–85, 128, 142, 215, 234–236 238, 242, 248, 251–253, 257, 265, 268, 269 Gymnolaemata , 248, 249

G Gamete , 91, 250, 265 H Gametogenesis , 201, 265–266 Hairy , 216 Ganglia , 44, 45, 49, 51, 53–58, 62, 80, 90, 119, 120, Haliotis , 116, 124–128, 136, 142 122–128, 140–142, 194, 195, 220, 221 HAM. See Hypothetical ancestral mollusk (HAM) Gangliogenesis , 123, 124 Handedness , 143–145 Ganglion , 2, 5, 7, 59, 84, 90, 96, 98, 116, 117, 124, 126, Hatching , 4, 5, 17, 18, 32, 33, 37, 107, 108, 123, 132, 130, 140, 232, 248, 249 212–214 Ganglionic , 84, 130, 211 Hatching envelope , 251 Gap gene , 217 Hatchling , 3, 5, 28, 37, 108 Gastropoda , 104, 106, 107, 117, 118, 123, 125 Hbn , 274 Gastrotrich , 13–18, 37 Head larva , 200 Gastrotroch , 204, 208, 209 Heart , 67–69, 116–118, 121, 123 Gastrula , 94, 169, 172, 174, 181, 237, 238, 251–254, Hedgehog , 53, 139 266–270, 273, 274 Hedgehog signaling , 217 Gastrulation , 3–9, 15, 16, 24, 25, 27, 30–32, 35–37, 63, Hemoglobin , 232 94, 95, 100, 106, 121–122, 126, 138, 139, 161, Hermaphrodite , 107, 159, 234 167–169, 174, 181, 201–202, 210, 216, 237, Hermaphroditic , 91, 111, 233, 265 238, 250–252, 254, 266–268 Hermaphroditism , 112 GATA , 222 Heterochrony , 123, 197, 212 GATA456 , 30, 258 Heteronemertea , 156–158, 163, 165, 174, 175, 177, 179, Gene , 2, 8, 9, 11, 15, 18, 28–30, 35, 43, 45, 48, 51, 55, 183, 184 57, 59–60, 64, 66, 85, 86, 100, 106, 107, 121, Hexapoda , 104 125, 127–130, 139–143, 146, 186–188 Hh , 128, 129 Gene conversion , 9 Hidden trochophore , 169 Gene expression , 8, 11, 15, 18, 29, 30, 85, 86, 100, 107, Hindgut , 14, 90, 181, 217, 222 121, 125, 127, 129, 130, 139–143, 146, Hippo , 23, 66–69 185–188, 201, 215–223, 257–259, 267, Hippo signaling pathway , 66–69 271–274 Hirudinea , 210 Gene expression pattern , 18, 85, 215, 217, 273, 274 Histamine , 272 Gene function , 48, 139, 143 Histaminergic neuron , 273 Gene network , 185–187, 220, 221 Holoblastic , 92 Genome , 8, 48, 61, 65, 106, 108, 144, 186, 198, 218, 249 Holoblastic cleavage , 84, 113, 161, 252, 266 Germarium , 5, 7, 27 Homeobox , 55 284 Index

Homeobox-containing genes , 9, 51, 60, 100, 125–129, J 139, 185, 220, 222 Jelly , 199 Homeobrain , 242, 274 Joint , 184 Homolog , 34, 45, 51, 53, 55, 56, 60, 64–66, 68 Juvenile , 3, 5, 15–17, 26–28, 30, 32, 33, 37, 99, 123, Homologous , 5, 27, 37, 50, 135, 146, 163, 167, 175, 177, 131–133, 136, 145, 157, 166, 169–174, 177, 184, 186, 188, 215, 256–258, 264, 270, 274 181–187, 198, 205–207, 215, 216, 221, 239, Homology , 16, 53, 96, 100, 121, 123, 135, 143, 169, 175, 240, 242, 250, 252, 253, 256–258, 264, 268, 180, 185, 202, 218, 223, 252, 253 270–272 Homonomous , 194 Juvenile tentacle , 99, 207, 239–242, 272 Hoplonemertea , 156–161, 163–172, 175–185, 188 Host , 2, 9, 11, 31–34, 43, 45, 80, 82–85, 106 Hox , 100, 124–129, 139, 143, 144, 186, 188, 223, 258 K Hox cluster , 9, 186, 218–220 Kamptozoan , 90 Hox gene , 9, 62, 100, 124–130, 139–143, 186, 188, Knockdown , 68, 198, 217 216–221 Kowalevsky, M.A. , 113, 138 Hub , 67, 69 Hunchback , 217, 218 Hydra , 56 L Hyposphere , 96, 98, 111, 112, 123, 125, 202, 207, 214 Lab , 141, 198, 218 Hypothetical ancestral mollusk (HAM) , 109 Lacuna , 90 Lacunifera , 91, 109 Lamina , 3 I Lappet , 167, 170–174, 186 Identity , 17, 61, 63, 65, 66, 143, 240 Larva , 9, 24, 26, 30, 32–34, 36, 37, 80, 82–85, 90, Identity specifi cation , 28, 61–62 96–99, 111, 112, 119, 120, 122, 123, 133, 134, Idiosepius , 104, 108, 129, 132, 141, 142 140–142, 156, 159, 167, 169–176, 179–184, Ilyanassa , 106–107, 113, 114, 116–118, 121, 128, 202, 204, 207, 213, 234, 237–240, 242, 130, 138 250–259, 268–273 Imaginal disks , 170, 173, 174, 179, 184 Larval , 4, 24, 43, 80, 90, 106, 161, 197, 232, 250, 265 Immunocytochemical , 124, 130, 212, 213 Larval development , 122–123, 171–172, 179, 202, 208, Immunohistochemistry , 51, 178 217, 218, 237–239, 251, 270, 271, 274 Immunoreactivity , 119, 130, 178, 212–214, 241 Larval phase , 183, 184, 215, 268 Immunostaining , 49, 51, 54 Larval settlement , 123 Indirect development , 24, 31, 36–38, 106, 111, 112, 122, Larval shell , 112, 136, 268, 269 130, 136, 169, 198, 199 Lats , 67 Individuality , 3, 5, 6, 42, 44, 48, 80, 83–85, 96, 99, 105, Lecithotrophic , 90, 122, 169, 174, 197, 198, 202, 205, 107, 108, 112, 113, 116, 122, 124, 130, 207, 208, 212, 213, 215 134–136, 143, 145, 146, 198, 200, 201, 204, Lecithotrophy , 208 216, 234, 235, 248 Leech , 199, 200, 210–212, 215–218, 220, 221 Induction , 6, 8, 105, 116 Left-right axis , 65 Inductive interaction , 163, 167, 168 Lepidodermella , 15–18 Infection , 2, 11, 19, 31–33, 45 Life cycle , 9–11, 23, 31–34, 36, 43, 45, 80–86, 106, Ingression , 145, 169, 181, 182, 252, 268 198, 207 In situ hybridization , 29, 106, 108, 139, 187, 198 Ligament , 138 Instar , 66 Limb , 43–45, 47, 48, 56, 68 Integument , 5, 185 Limb bud , 68 Intercalation , 42, 177 Lineage , 9, 22, 24, 27, 36, 37, 43, 47, 53, 60, 92, 96, 107, Internal budding , 137, 138 109, 111, 113–116, 121, 124, 130, 138, 139, Internal fertilization , 2, 9, 14, 17, 84, 91, 106, 107, 160, 143, 166, 168, 169, 185, 198, 208, 210, 211, 161, 197, 234 215, 222, 233, 237, 250, 264, 268 Internal sac , 138, 139, 239, 251–259 Lineage tracing , 121 Interstitial , 2, 14, 111, 194 Lingula , 264, 265, 268, 271 Intertentacular organ (ITO) , 251 Linguliformea , 264 Intestinal stem cell , 68 Lipid , 198, 206 Invagination , 58, 59, 99, 121, 136–138, 169, 171–174, Lobster , 80, 83–85 178–184, 201, 237, 238, 251 Longitudinal muscle , 37, 80, 83, 90, 97, 98, 133, 156, Inversion , 64 179, 212, 232, 239, 256 Involution , 254 Longitudinal patterning gene , 100, 139 Irregular cleavage, 23, 24, 236, 252 Lophophorate , 175, 265 Index 285

Lophophore , 232–235, 240, 242, 248, 249, 257, 264, Metatrochophore , 204–207, 209 265, 268 Metazoan , 3, 4, 8, 35, 43, 61, 63–66, 69, 84, 113, 217, Lophotrochozoa , 84, 90, 91, 96, 97, 99, 109, 116, 124, 258, 264 131, 143, 201, 218, 222, 249 Micrognathozoa , 2, 3 Loxosoma , 91 Micromere , 3, 5, 9, 24–27, 33–36, 93, 96, 113, 114, 116, Loxosomatid , 91 121, 122, 128, 130, 138, 139, 144, 162–168, Lumen , 1, 3, 7, 18, 131, 215 199–202, 206 Midbrain/hindbrain boundary , 142 Midgut , 14, 116, 156–158, 171, 181, 182, 206, 222 M Midgut gland , 116 Macromere , 5, 9, 24–28, 32, 34, 35, 84, 94, 96, 113, 114, Midline , 62, 64, 65, 220, 269 116, 118, 121, 127, 139, 162–166, 199, 200, Mitochondria , 59, 113, 160, 198, 233, 249, 255 206, 251, 254 Mitochondrial genome , 249 Macrostomum , 26, 27, 35, 36, 43, 44, 47, 49, 52, 57, Mitotic spindle , 113, 199 68, 69 Mitraria , 204, 207 Main cord , 49, 50 Model organism , 26, 198, 222 Mandible , 194 Model species , 45, 47, 48, 66, 69, 131, 142, 198, 250 Mantle , 25, 26, 107, 112, 117, 120, 127, 128, 131, 132, Modifi cation , 23, 42, 166, 200, 242, 271 135, 136, 138–141, 143, 252, 257, 264, 265, Mollusca , 57, 103–146, 163, 175, 194, 200, 212, 241 267–272 Molluscan cross , 200 Mantle cavity , 112, 117, 128, 264 Mollusk , 33, 34, 43, 95, 96, 105, 109, 130, 237 Mantle lobe , 268–273 Molt , 194 Maternal determinant , 36 Molting process , 194 Maternal mRNA , 199 Monophyletic , 3, 22, 57, 88, 90, 91, 109, 156, 194, Mats , 67 248, 249 Median hood nerve , 240, 241 Monophylum , 15, 157 Median tentacle , 268, 269 Monophyly , 90, 96, 109, 157, 158, 233, 265 Medullary cord , 156, 157, 177, 178, 184, 186, 187 Monoplacophora , 104, 109, 110, 112, 116, 122–124, Meiofauna , 215 131, 133, 146 Meiosis , 113, 234 Monostilifera , 156–158, 168, 181, 183 Membranipora , 248, 250, 251, 253 Morphallaxis , 46 Meniscotroch , 202 Morphogen , 139 Mesenchymal , 100, 167, 173, 179, 237, 238 Morphogenesis , 46, 48, 57, 60, 67, 116 Mesenchyme , 23, 118, 168 Morpholino , 106, 198, 217 Mesendoblast , 3, 92, 96, 116, 118, 122, 126, 167, 179 Morphology , 28, 38, 57, 85, 90, 91, 96–97, 107, 111, Mesendoderm , 122 135, 158, 160, 161, 175, 182, 201, 202, 205, Mesenteron , 9 207, 233, 239, 249, 250, 253, 255, 264, 265, Mesocoel , 232 269, 272 Mesoderm , 17, 18, 25, 26, 30, 35, 42, 92, 96, 115, 116, Mosaic , 8, 237 121–122, 125–127, 165, 167, 179, 180, 201, Mosaic development , 8 217, 232, 237, 238, 267, 268 Mouth , 2, 5, 7, 14, 28, 38, 80, 84, 90, 96, 99, 112, 116, Mesodermal , 5, 17, 18, 96, 99, 122, 128, 139, 158, 167, 121, 122, 134, 156, 170, 171, 174, 178, 172–174, 177–180, 182, 183, 201, 210, 211, 181–184, 187, 201, 202, 204, 205, 209, 221, 215–217, 252, 256, 258, 259, 268 232, 237–239, 248, 253, 267 Mesoderm mother cell , 96 Msx , 188, 217, 221 Mesoteloblast , 201, 210 Multipotent , 258 Metacoel , 232, 265 Muscle , 11, 28–30, 32, 33, 37, 47, 58, 83, 90, 96–99, Metagenesis , 80 109, 120, 121, 131–135, 156–158, 168, 173, Metagenetic , 80 177, 179–181, 197, 201, 212, 232, 237, 238, Metamorphic , 96, 97, 99, 107, 112, 119, 207, 232, 241, 254, 256 258, 268, 270, 271, 275 Muscle founder cell , 92, 116 Metamorphosis , 30, 36, 37, 96–100, 105, 122, 123, 130, Muscle pioneer , 28 131, 133, 135, 136, 169, 174, 175, 177, 179, Muscle precursor , 135 181, 185, 205, 207, 215, 232, 238–240, 242, Muscle progenitor cell , 47, 48, 56, 184 252–258, 270–272 Musculature , 5, 11, 17, 18, 26, 30, 33, 37, 80, 83, 84, 90, Metanephridia , 123, 194, 215, 234, 235, 265 96, 97, 111, 120, 123, 132–135, 158, 168, 172, Metasoma , 239 173, 175, 179, 180, 185, 197, 211, 212, Metasomal sac , 239 239–241, 248, 252–254, 264, 271 Metatroch , 202–209 Mushroom bodies , 197 286 Index

MyoD , 139 Non-feeding , 208, 239, 248, 250, 252–255, 257, 265, 268 Myoepithelial cell , 232 Notch/Delta , 221 Myogenesis , 131–135, 139, 212 Notch pathway , 216, 217, 221 Myosin , 30, 35 Nototroch , 204 Myzostomida , 194, 201, 203 Novelties , 109, 111, 143 Novocrania , 264–268, 270–272 Nuclei , 9, 11, 92–94, 120, 132, 234, 235, 251, 254, 255, N 259, 273 Nacre , 136, 143 Nucleus , 42, 61, 67, 160, 161 Nanos , 8, 60, 128 Nautilus , 138, 142 Nectochaete , 205–207 O Nectosoma , 207 Ocelli , 58, 59, 121, 171, 255, 269–271, 273, 274 Nematode , 64, 105 Ocellus , 273, 274 Nemertea , 155–188, 194 Octopamine , 53, 56 Nemertean , 91, 95, 156–161, 165, 167–169, 174–176, Octopod , 105, 112, 123, 132, 136, 137, 139 180–182, 184, 185, 188, 199–201, 232, 233, 264 Ontogenetic , 27, 31, 33, 38, 42, 105, 106, 121, 122, 130, Nemertodermatida , 50 135, 136, 210, 213 Neoblast , 23, 29–30, 38, 42–43, 47, 48, 53, 56–57, 68, 69 Ontogeny , 11, 37, 99, 136–137, 139, 204, 213 Neodermata , 22, 24, 31–32, 34, 35, 37 Oocyte , 3, 8, 23, 27, 32, 33, 36, 82, 84, 85, 113, 162, Neomeniomorph , 96, 104, 109–112, 115–116, 120, 123, 188, 234–236, 250, 265 131, 133, 134, 139, 144–146 Oogenesis , 106, 161, 207 Neomeniomorpha , 104, 109, 144–146 Operculum , 123, 126, 141, 249 Neonemertea , 158–159, 163, 176–177, 179, 183, 184 Optic lobe , 132, 141 Neoophora , 22, 23, 27, 29, 32–38, 50 Oral-aboral axis , 63 Nephridia , 5, 158, 182–183, 185, 194, 205, 215, 236 Oral ciliated cell , 255–257 Nerve , 2, 4, 37, 49, 56, 59, 84, 90, 96, 98, 116, 117, 121, Oral fi eld , 252 125, 131, 158, 172, 178–179, 183, 186, 188, Ordovician , 248 195, 213, 214, 232–233, 240 Organizer , 36, 116 Nerve cord , 28, 30, 49–51, 53–57, 84, 97, 111, 117, 120, Organogenesis , 5, 11, 18, 25, 29, 30, 32, 33, 37–38, 86, 121, 124, 125, 158, 194, 213, 215, 217, 220, 96, 100, 222 221, 241 Organophosphatic sclerite , 233, 264 Nerve net , 49, 132 Orthogon , 30, 37, 49–50, 195, 199 Nerve ring , 49, 96, 213, 233, 241 Ortholog , 8, 43, 212–213, 216, 218, 221 Nervous system , 18, 23, 28, 30–33, 38, 49–62, 64, 65, Oscillation , 161 90–91, 97, 98, 109, 111, 120, 122–124, Otd , 60 127–131, 141, 142, 156, 165, 168, 169, 171, Otp , 51, 127, 142, 220, 242, 273, 274 172, 177–179, 194–195, 197, 198, 212–217, Otx , 51, 59–60, 127, 129, 142, 185–187, 220, 221, 274 220–223, 232, 239–241, 248, 251, 253, 256, Ovarian , 161 272–274 Ovary , 32, 165, 234, 236, 252 Netrin , 55, 56, 60, 221 Oviduct , 5 Neural marker , 83–84 Neural progenitor cell , 56, 220 Neurite , 55, 120, 131, 177 P Neurite bundles , 14, 84, 98, 172, 241 Paired , 2–4, 14, 18, 30, 33, 34, 49, 50, 55, 59, 60, 82, 90, Neuroanatomy , 80 96–98, 111, 116, 125, 126, 131, 133, 135, 139, Neuroblast , 37 156, 161, 163, 164, 167–174, 176–178, 180, Neuroectoderm , 126, 178, 220, 221, 274 183, 184, 194–195, 201, 205, 210, 211, Neurogenesis , 30, 38, 53, 56, 123–131, 142, 212–217, 213–218, 220, 234–236, 238, 239, 250, 253, 220–222 265, 268–272 Neuropeptide , 52, 57, 60, 213 Pair rule gene , 216–218 Neuropil , 49, 51, 55, 56, 98, 130, 132, 168, 172, Pallial epithelium , 253–259 239–241, 272, 273 Palliovisceral , 125, 128 Neurotransmitter , 57, 124, 185, 186, 242 Palp , 194, 197, 206 Neurotroch , 202–205, 207–209 Pandora , 80–85 Nidamental gland , 233 ParaHox genes , 100, 124–126, 130, 139–145, 188, Nitric oxide (NO) , 131, 161 218–220, 222, 223, 242, 258 NK2.1 , 127, 142, 220, 221, 242, 274 Paralog , 216–218, 220 NK4 , 129, 139, 217 Parapodia , 194, 197, 204–207, 211 Nodal , 127, 143–145 Parasite , 34, 105, 159, 194 Noggin-like genes (nlg) , 65 Parasitic , 2, 3, 9, 22, 23, 31–34, 37, 42–43, 45, 51, 59, 156 Index 287

Paratomy , 45, 57 Plasticity , 23, 64, 91, 109, 135 Paratroch , 204, 206 Plate , 26, 38, 91, 112, 123, 131, 133, 136, 138, 139, 202, Paraxial mesoderm , 127, 128 210–213, 237, 252, 256 Patella , 95, 104, 113, 116–118, 121, 123, 126–128, 135, Platyhelminthes , 21–38, 41–69, 175, 212 138, 139, 142, 185 Platynereis dumerilii , 186, 197–199, 201, 203, 206, 210, Paterinid brachiopod , 264 212, 213, 215–218, 220–222 Pattern , 3, 9, 11, 15–18, 22–30, 32, 34, 38, 47–50, 53, 56, Plesiomorphic , 23, 130, 133, 156, 207, 237, 274 57, 61, 63–66, 68, 84–85, 92, 94–95, 100, 108, Plumatella fungosa , 252 113, 114, 121, 125, 133, 136, 138–140, 142, Pluripotent , 42–43, 48, 210, 258 144–145, 158, 163–165, 168, 194, 195, 197, Poecilogeny , 197, 210 199–201, 210, 212–218, 220–222, 233, 234, Polar bodies , 4–7, 9, 11, 15, 17, 85, 93, 113, 114, 161, 236, 237, 242, 249, 250, 258, 265–267, 273, 274 162, 165, 234, 235, 238, 250, 254, 266, 267 Pax , 127, 129, 141–142 Polarity , 6, 23, 29, 36, 48, 57, 61, 63, 66, 67, 198, Pax3/7 , 220–221 216, 258 Pax6 , 30, 53, 59, 60, 129, 142, 220–221, 274 Polar lobe , 113, 114, 116, 199 Pedal , 97, 98, 117, 124–128, 131, 132 Polyclad , 23–27, 34–38, 43, 44, 59, 95, 175 Pedal nerve cord , 97, 120 Polyembryony , 252 Pedicellina , 91–92, 96 Polyp , 80 Pedicle , 264, 268, 269, 271, 272 Polypide , 248, 250, 252, 253, 256–258 Pedicle lobes , 269–271, 273 Polyplacophora , 104, 109, 117–118, 125, 139, 144–145 Pelagic , 112, 122, 132, 136–138, 156, 160, 168–170, Polystilifera , 156–158, 183, 185 174, 185, 188 Polyzoa , 80, 91, 249 Pelagosphaera , 207 Postembryonic development , 3, 11, 37 Perikarya , 97, 98, 213–215 Posterior ectoderm , 127, 201, 237 Perinotum , 136 Posterior growth zone , 109, 133, 194, 203–204, 207, Periostracum , 136, 138, 271 210, 211, 213, 216, 217 Peripheral nervous system , 195 Posterior lobe , 172, 271 Peristomial , 202 Posterior mesoderm , 126 Peristomium , 194, 207, 219, 221 Postmaxillary Phalloidin , 83, 120, 132, 134, 211, 240, 273 POU , 127, 129, 141–143 Pharynx , 2, 3, 5, 14, 15, 18, 25, 26, 28–30, 35–37, 42, Precambrian , 104–105 44–45, 49, 54, 62–63, 181, 248 Primary germ cell , 201 Phenotype , 55, 60, 62–66, 68, 104, 143 Primordial germ cell , 42, 201 Phoronid , 91, 163, 194, 231–242, 249, 253, 258, Primordium , 3, 25, 26, 28–30, 33, 37, 57, 59 264–266, 271, 274 Proboscis (pb) , 7, 11, 157, 158, 187, 218, 235, 254 Phoronida , 194, 231–242 Proctodeum , 15–17, 206, 222 Phoronis , 232–236, 238–240, 242 Prodissoconch , 136 Phoronopsis , 232–234, 238–242 Programmed cell death , 33 Phosphatic , 264 Proliferation , 5, 17, 18, 25, 26, 28, 33, 42, 43, 45–48, 56, Photoreceptor , 23, 30, 47, 57–61, 97, 171, 185, 186, 197, 57, 67–69, 124, 169, 174, 177, 179, 215, 220, 220, 274 252, 267–268, 274 Phototactic behavior , 274 Prometheus , 80, 82, 84 Phylactolaemata , 248, 249 Prostomium , 194, 197, 207, 221 Phylogenetic analysis , 2, 3, 14–15, 22, 23, 34, 44, 49, 50, Protoconch , 112, 136, 138, 139 64, 80, 106, 109, 157–158, 160, 169, 175, 184, Protonephridia , 84, 123 194, 200, 216, 218, 232, 233, 241, 249, 252, Prototroch , 117 264, 265, 274 Pseudocyphonautes , 253 Phylogenetic tree , 64 Phylogenomic , 14–15, 80, 110, 194, 233, 249, 265 Phylogeny , 2, 34, 37, 104, 109–111, 159, 175–176, 194, R 195, 197, 213, 249 Radial cleavage , 163, 234, 237 Pigment cell , 58–60, 97, 171, 197, 274 Radula , 110–111, 126, 127, 131, 141, 142 Pilidiophora , 156–159, 161, 164–167, 169, 170, Reconstruction , 8, 18, 85, 92–94, 105, 109, 113–114, 172–175, 177–179, 181–185, 188 124, 130, 134, 135, 143, 144, 207–208 Pilidium , 156, 159, 166–175, 180, 181, 183, 186 Regeneration , 23, 28, 30, 41–69 Planarian , 28–30, 35, 37, 42, 43, 46, 48, 51, 53, 55–66, Regionalization , 105, 156 68–69 Regulation , 9, 30, 35, 42, 43, 56, 60, 61, 64–68, Planktonic , 37–38, 96, 106, 111–112, 123, 169, 265, 139–140, 143, 165–168, 187, 188, 217, 220, 266, 268 234, 274 Planktotrophic , 96, 107, 136, 197–199, 202, 207, 208, Regulative capacities , 166 212, 213, 215, 257, 268 Remodelling , 46 288 Index

Reptantia , 156 Sexual , 44, 48, 69, 80, 84, 86, 106, 108, 197, 198 Respiration , 232 Sexual dimorphism , 197 Respiratory organ , 90 Shavenbaby , 60 Retina , 185–188 Shell , 4, 6, 15, 16, 25, 28, 34, 104–107, 109, 111–113, Retinal determination gene network (RDGN) , 185, 187 121, 123, 131, 133, 135–140, 142–145, 257, Retractor , 10–11, 96, 97, 99, 117, 118, 120, 121, 131, 264, 265, 268–271 132, 135, 158, 180, 197, 212, 253 Shell fi eld , 121, 125–128, 136–139, 143 RFamide , 52 Shell gland , 117, 121, 138 Rhabdomeric photoreceptor , 59, 197 Shell sac , 137–139 Rhomboid , 175 Shield , 274 Rhynchocoel , 156–159, 177, 180, 183, 185, 188 Shrimp , 232 Rhynchodeum , 156, 158, 177, 180–182 Signal , 25, 34–36, 43, 57, 63–64, 83–84, 116, 131, 143–144, Rhynchonelliformea , 264 161, 186, 213, 216, 217, 219, 221, 236, 237 Ring muscle , 83, 90, 96–99, 133, 134, 253–254 Signalling pathway , 23, 30, 43, 53, 61–69 Ring vessel , 232 Sine oculis , 30, 51–52, 60 RNAi , 8, 34, 48, 55, 57, 60–66, 68, 106, 139 Single minded (sim) , 55, 56, 221 Robo homolog , 66 Sinusoida , 90–91, 109 Rostraria , 207 Sipuncula , 194, 203 Rotifer , 4–9, 11, 80 Six3 , 212–213, 220, 221 Rotifera , 2, 4, 5 Six3/6 , 129, 185, 242, 274 Roundabout , 53, 55 Skeletogenic cell Row , 5, 18, 114, 158, 166–167, 172, 250, 270 Skeleton Rudiment , 96, 99, 113, 122, 126, 127, 133, 134, 137, 166, SL. See Secondary lobe (SL) 168–171, 173–176, 178–185, 187, 212, 240–242 Slit , 55, 56, 65–66, 90, 237, 269, 270 Run , 49, 98, 241 Small intestine , 32 Runt , 216, 217 Snail , 30, 31, 33, 63, 104–106, 122, 128, 131, 139, Rx factor , 220 143–145 Solenogastres , 104 Solitaria , 90 S Solitary , 90–92, 97, 99 18S , 249 Somatic mesoderm , 180 Saccostrea kegaki , 106, 113, 121, 125, 139 Spawning , 105–107, 111, 113, 159–161, 197, 198, 233, Salivary gland , 105, 118 251, 265 Scalloped , 105 Specifi cation , 3, 18, 23, 28–32, 35–38, 43–44, 47, 48, 51, Scaphopoda , 104, 117, 118, 146 56–66, 69, 85, 90, 97, 113, 116–117, 123, 129, Schizocoely , 180, 185 131, 139, 142, 143, 146, 163, 167–169, 185, Schmidtea , 28, 29, 35, 37, 42–44, 46–48, 51–54, 61–62, 186, 188, 198, 200–202, 210, 217, 220, 221, 64–66, 68 242, 253, 267, 272, 274 Scr , 218 Spengel, J.W. , 197 Secondary lobe (SL) , 270 Sperm , 22, 113, 160–161, 199, 234, 265 Sedentaria , 194 Spermatophore , 234 Segment , 33, 109, 131, 156, 204–208, 210–213, Sperm transfer , 160 215–219, 222, 223 Spicule , 104, 117, 121, 123, 136, 138 Segmental , 156, 194–195, 204, 207, 210, 213, 215–217 Spine , 15, 249 Segmentation , 194, 197, 198, 204, 210–211, 213, Spiral , 3, 4, 11, 23–27, 34–36, 38, 84–85, 90, 92, 93, 216–218 113, 114, 116, 145, 161, 163–166, 168, Segmentation gene , 216, 223 199–201, 234–237, 242 Segment polarity , 216 Spiral cleavage , 3, 4, 11, 23–27, 34–36, 38, 84–85, 90, Segment polarity gene , 216 92, 93, 113, 114, 116, 164–166, 168, Seison , 9 199–201, 234 Sensory organ , 57, 82, 84, 90, 96, 122, 123, 129, 158, Spiralia , 2, 3, 5, 14–15, 22–24, 27, 35–38, 92, 94–96, 183–185, 197 107, 116, 121, 133, 144, 156, 165–168, 175, Sepia , 107–108, 129, 139 180, 185, 187, 201, 233, 236, 237, 249, 253, Seriality , 133, 144 258, 264, 268, 274 Serotonergic , 50, 51, 53, 55–57, 84, 97, 120, 132, 172, Sponge , 65 175, 176, 178, 213–215, 233, 240, 242, 274 Stalk , 80, 82, 85, 90, 99, 162, 165 Serotonin , 38, 50–52, 57, 83–84, 96, 98, 106, 119, 120, Statocyst , 117, 121, 123, 127, 140, 142, 143, 268 124, 128, 130, 131, 176, 212–214, 240, 241, 274 Stellate cell Sessile , 207, 264 Stem cell, 23, 28, 38, 42–43, 46, 48, 66–69, 201, 210, 211 Settlement , 84–86, 99, 123, 131, 136, 242 Stenolaemata , 248, 249 Index 289

Sterroblastula , 121 Twist , 15, 30, 35, 63, 113, 139, 188, 235–237 Stolon , 99, 248, 254 Stomodeum , 15–17, 127, 141, 178, 181, 183, 206 Stripe , 59, 187–188, 217 U Stylet , 156, 158, 159, 170, 180 Unequal cleavage , 163, 199 Subumbrella , 171–173, 183, 237, 238 Unequal division , 210 Swimming-type larva , 96, 98 Unpaired , 173, 174 Symbion , 80–86 Urbilateria , 50, 53 Symmetry , 61, 143–144, 169, 254, 266, 267 U-shaped , 80, 90, 232, 235, 239, 242, 248, 249, 264 Synapomorphy , 249 Synapsin , 84 Synchronous , 133, 163 V Syncytium , 25, 28, 31, 33–35, 114, 122 Vasa , 8, 28, 43, 125, 128, 139 Syndermata , 2, 3 Vein , 31 Syndermate , 80 Veliger , 105, 107, 111, 116, 120, 123, 125–128, 135, 141 Ventral midline , 64, 128, 220 Ventral nerve cord , 28, 30, 49, 51, 53–57, 84, 194, 213, T 215, 217, 220, 221, 241 Talpina , 233 Ventral valve , 264, 271 Target gene , 67 Vermiform , 90, 111, 233, 242 Teleoconch , 123, 136, 138, 139 Vertebrate , 9, 11, 32–34, 43, 45, 47, 48, 50, 53, 57, 63–65, Teloblast , 201, 202, 210–211, 216, 217 69, 124, 143, 163, 202, 215–216, 220–222 Teloblastic growth zone , 210 Vesicular bodies , 269, 270 Telotroch , 112, 117, 121, 123, 125, 174, 202–206, 208, Vestibule , 251–253, 257 209, 238, 240, 241 Vibratile plume , 252, 253, 255, 257 Tentacle , 32, 90, 99, 104, 116, 117, 123, 127, 142–143, Visceral , 97, 117, 120, 121, 123–126, 128, 136, 141 205, 207, 232–235, 238–242, 248, 254–255, Vitellaria , 23, 27 259, 264, 268, 269, 272 Vitelline , 33, 34, 160–161, 234 Tentacle nerve , 241 Viviparity , 197 Terebratalia , 242, 265, 266, 268–274 Viviparous , 160 Test , 267, 268 V-shaped sperm , 234 Test cell , 123, 144, 146 Tetraneuralia , 90–91, 109, 130 Tetraneurous , 90–91, 97, 109, 111, 124 W TGF , 34, 64 Wart , 67 TH , 238, 240 Wingless , 216 Thoracic , 219 Wirenia argentea , 109, 112, 133, 134 Tommotiid , 233, 264 WntA pathway , 217 Transcription , 42, 64–68, 217 Wnt gene , 216, 217 Transcription factor , 18, 47, 53, 59–61, 129, 139, 143, Wnt pathway , 23, 29, 30, 36, 53, 55, 61–66, 68, 127, 186, 216–218, 220, 222 216, 217, 220 Transcriptome , 48, 59, 100, 144 Wnt8 pathway , 217 Transgenesis , 69, 106 Wnt10 pathway , 258 Transmission , 36, 84, 130, 212, 273 Wnt signaling , 258 Tree , 22, 44, 64, 80, 91, 109, 194, 208 Trilobed larva, 269, 270, 272–274 Trochoblast , 92, 96, 111, 115, 116, 121, 122, 125, 127, Y 165–167, 169, 177, 182, 185, 186, 200–202 Yap , 67–68 Trochophore , 80, 86, 106, 107, 111, 112, 119, 120, 122, Yolk cells , 23, 25–31, 33–35, 37, 63, 113, 114, 118, 121, 123, 125–128, 131, 141, 142, 144, 167, 169, 122, 132, 137, 138, 163, 169, 185, 188, 174, 182, 186, 197, 201–207, 209, 212, 215, 198–202, 207–209 222 Y-organ , 18 Trunk , 80, 82–85, 171, 173–175, 178–180, 183, 194, Yorkie , 66–69 206, 207, 232–234, 236–242 Trunk sac (TS) , 238–240, 242 Tuft , 82, 96, 98, 112, 117, 119, 120, 122, 123, 130, 165, Z 166, 170–172, 174–176, 186, 202–208, Zinc fi nger , 59–60, 217 212–214, 237, 238, 253, 255, 273, 274 Zooid , 45, 91, 99, 248, 249, 251–254, 258 Turbanella , 15–18 Zygote , 11, 15, 28, 33, 42, 106, 111–113, 122, 162, 166, Twin of eyeless , 60 234, 235, 251