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Home , Sipuncula, Sipunculidae, Themiste pyroides, Trochozoa

1

THE MOLECULAR AND DEVELOPMENTAL BIOLOGISK BASIS OF BODYPLAN PATTERNING IN INSTITUT AND THE EVOLUTION OF

SEGMENTATION

Alen Kristof | Københavns Universitet 2

DEPARTMENT OF BIOLOGY FACULTY OF SCIENCE UNIVERSITY OF COPENHAGEN

PhD thesis Alen Kristof

The molecular and developmental basis of bodyplan patterning in Sipuncula and the evolution of

Principal supervisor

Associate Prof. Dr. Andreas Wanninger

Co-supervisor

Prof. Dr. Pedro Martinez, University of Barcelona

April, 2011 3

Reviewed by: Assistant Professor Anja Schulze Department of , Texas A&M University at Galveston Galveston, USA

Professor Stefan Richter Department of Biological Sciences, University of Rostock Rostock, Germany

Faculty opponent: Associate Professor Danny Eibye-Jacobsen Natural History Museum of Denmark, University of Denmark Copenhagen, Denmark

______Cover illustration: Front: Frontal view of an adult specimen of the sipunculan pyroides with a total length of 13 cm.

Back: Confocal laserscanning micrograph of a Phascolosoma agassizii pelagosphera showing its musculature. Lateral view. Age of the specimen is 15 days and its total size approximately 300 µm in length. 4

“In a world that keeps on pushin’ me around, but I’ll stand my ground, and I won’t back down.”

Thomas Earl Petty, 1989

5 Preface

Preface The content of this dissertation comprises three years of research at the University of Copenhagen from May 1, 2008 to April 30, 2011. The PhD project on the development of Sipuncula was mainly carried out in the Research Group for Comparative , Department of Biology, University of Copenhagen under the supervision of Assoc. Prof. Dr. Andreas Wanninger. I spent nine months working on body patterning genes in the lab of Prof. Dr. Pedro Martinez, Department of Genetics, University of Barcelona, Spain. Within these three years, research trips of altogether 16 weeks were made to collect, rear, and fix different sipunculan at the Sven Lovén Center for Marine Sciences in Kristineberg, Sweden (6 weeks), at the Espeland Marine Biological Station in Bergen, Norway (4 weeks), and at the Vostok Station of the A. V. Zhirmunsky Institute of Marine Biology in Vladivostok, Russia (6 weeks). Furthermore, PhD courses had a great impact on my scientific knowledge and skills. During my studies I attended the EMBO course Marine Models in Evolution and Development at the University of Gothenburg, Sweden, Comparative Embryology of Marine at the Friday Harbor Laboratories, University of Washington, USA, and Scientific Writing at the University of Copenhagen, Denmark. This PhD project was funded by the European Research Council (MEST-CT-2005-020542) and the Faculty of Science of the University of Copenhagen. This thesis is composed of four chapters. The first chapter gives a general introduction to the topics of the present study, its objectives, and the discussion of the results in a broader perspective, leading to conclusions and perspectives for future research. Chapters II to IV contain the major findings of this PhD project in three published papers.

Copenhagen, April 2011 Alen Kristof Acknowledgements 6

Acknowledgements I am heavily indebted to my principal supervisor Andreas Wanninger for more than three years of support, guidance, education, motivation, and invaluable help. His overall excitement for evolution, invertebrates, science, and finally had the greatest impact on me. Thank you Andi! Gratefully acknowledged is also my co-supervisor Pedro Martinez for a great time in his lab in Barcelona. Many thanks are going to his lab members Johannes Achatz, Alex Alsen, Eduardo Moreno, Elena Perea, and especially to Marta Chiodin who introduced me to molecular techniques as well as to the city of Barcelona. Furthermore, I am grateful to José María Martín-Durán from the “planarian group” also from the University of Barcelona for helping me establishing an in situ protocol for my . I thank Prof. Dr. Danny Eibye-Jacobsen that he agreed to act as the head of my defence committee at the University of Copenhagen, as well as Prof. Dr. Stefan Richter and Prof. Dr. Anja Schulze for kindly reviewing this thesis. Thanks to the working group for Comparative Zoology for a wonderful time in Copenhagen. All these years of the PhD project would not be the same without these members and associates. Therefore, special thanks are going to my friends Tim Wollesen, Uwe Spremberg, Judith Fuchs, and my crazy office mate Jan Bielecki. Special thanks are also going to Anders Garm who helped me with the Danish abstract. Thanks man! In addition, I would like to thank Dany “Sahne” Zemeitat, Ricardo Neves, Nora Brinkmann, Lennie Rotvit, Andreas Altenburger, Birgit Meyer, Henrike Semmler, Christopher Grubb, Tommy Kristensen, Jens Høgh, Lisbeth Haukrogh, and Åse Jaspersen. A huge thank-you goes to Christiane Todt (University of Bergen, Norway) for hosting and taking care of me during my visit at the Marine Station of Bergen. In addition, I want to thank Anastassia S. Maiorova (Marine Biology Institute, Vladivostok, Russia) for being helpful, reliable, and fun to work with during my time at the Vostok Station. I am grateful to Andrey V. Adrianov (Marine Biology Institute, Vladivostok, Russia) and the staff of the Vostok Station for their help and hospitality. For comments, important discussions and collaboration, I highly acknowledge Mary Rice (Smithsonian Research Center, Florida, USA) and Katrine Worsaae (University of Copenhagen, Denmark). 7 Acknowledgments

I am deeply grateful for the never ending support from my , in particular my parents. Za sve što ste napravili da krenem ovim uspješnim putem, jednostavno se zahvaljulem vama tisuču puta. Puno hvala mama! Puno hvala tata i puno hvala mom burazu Jožefu! Finally, thousand thanks to my lovely girlfriend Maria Eracleous for being the one. Maria mou, ise i zoi mou! Also, I want to thank all members of her family, especially Despina, Iraclis, and Georgia for treating me as one of them. For financial support I would like to thank Friday Harbor Laboratories, Washington, USA, the European Research Council (MEST-CT-2005-020542), and the Faculty of Science, University of Copenhagen, Denmark.

Content 8

Content

Preface…………………………………………………………..…………………...... 5 Acknowledgements………………………………………………..……………....6 – 7 Content……………………………………………………….………………………..8

Chapter I……………………………………………………..…………………...9 – 46 Danish abstract……………...…………………………………………………9 Abstract……….…………….………………………………………………..10 Short abstract………………....………………………………………………11 General introduction.…………....……………………………………....12 – 15 Sipuncula – a neglected taxon with affinities...... 13 – 15 Thesis objectives..…………………………………………………....…15 – 16 Material and methods…………………………………………………...16 – 22 RNA isolation and cDNA synthesis...... 17 Degenerate PCR...... 17 – 19 Rapid amplification of cDNA ends (RACE)...... 19 – 21 Whole mount in situ hybridisation...... 21 – 22 Results...... …………………………………………...... 22 – 31 Cloning of homeobox genes in ...... 22 – 23 Establishing of an in situ hybridisation protocol...... 23 – 24 Serotonergic and FMRFamidergic nervous system development in sipunculan worms (chapters I-III)...... 25 – 30 Myogenesis and cell proliferation during sipunculan development (chapter IV)...... 30 – 31 General discussion...... 31 – 36 Early neurogenesis in ...... 31 – 32 Establishment and loss of segmentation during sipunculan neurogenesis...... 32 – 33 Comparative trochozoan neurogenesis...... 33 – 34 Sipunculan growth zone(s)...... 34 – 35 Myogenesis in sipunculans and ...... 35 – 36 Conclusions and future perspectives…………………………………....37 – 38 References…………………………………………………………...... 38 – 46

Chapter II Kristof A, Wollesen T & Wanninger A. 2008. Segmental mode of neural patterning in Sipuncula. Current Biology 18: 1129-1132...... ….….....47 – 51 Chapter III Wanninger A, Kristof A & Brinkmann N. 2009. Sipunculans and segmentation. Communicative & Integrative Biology 2: 56-59.………..52 – 56 Chapter IV Kristof A, Wollesen T, Maiorova AS & Wanninger A. 2011. Cellular and muscular growth patterns during sipunculan development. Journal of Experimental Zoology Part B: Molecular and Developmental Evolution 316B: 227-240...... ……..………………………………...... 57 – 71

9 Chapter I – Danish abstract

Chapter I

Danish abstract Pølseorme (Sipuncula) har gennem tiderne været regnet som nære slægtninge til mange forskellige dyregrupper, men for nyligt har molekylære data placeret dem indenfor ledormene (Annelida) på trods af sipunculidernes ikke leddelte krop. For at undersøge om sipunculide orme har spor af en leddelt krop i deres ontogeni, blev nerve- og muskel-dannelsen sammen med fordelingen af celledelinger undersøgt hos 3 arter af sipunculider. Resultaterne viser, at rudimenter af mange ringmuskler i kropsvæggen og den længdegående retraktormuskel dannes samtidigt i den tidlige trochophor-larve. Ydermere, gennem hele udviklingen dannes nye ringmuskler via spaltning af allerede eksisterende muskelceller. I modsætning til det, så følger nervedannelsen et leddelt mønster gennem dannelsen af parrede serotonin-holdige cellekroppe langs den anteriøre-posteriøre akse. Cellekroppene er associeret med en parret ventral nervestreng og er arrangeret i fire stringent gentagne grupper. Dette leddelte mønster forsvinder inden metamorfosen, og den parrede ventrale nervestreng fusionerer til en enkelt nervestreng i det voksne dyr. Fordelingen af mitotiske celler viser sammenstemmende en lighed med en annelid-ligende posteriør vækstzone, som også forsvinder hos metamorfiserende pelagosphera larver. Disse udviklingsmæssige og morfologiske data understøtter således de nye molekylære analyser og antyder, at sipunculider stammer fra en leddelt forfar. Tabet af en leddelt krop hos disse relativt store og fritlevende dyr indikerer desuden, at leddeling forsvinder lettere gennem evolutionære processer end tidligere antaget. Initialiseringen og efterfølgende (sekundære) tab af leddeling gør Sipuncula ideel til evolutionære og udviklingsmæssige studier af leddelingsprocesser indenfor Metazoa.

Abstract 10

Abstract worms (Sipuncula) have been variously related to a number of animals in the past, but, recently, and despite their non-segmented adult body, molecular phylogenetic analyses place them within Annelida. In to contribute to the question whether or not sipunculan worms show traces of a segmental pattern in their ontogeny, neuro- and myogenesis as well as the distribution of proliferating cells were analysed in three different sipunculan species. The data show that the rudiments of numerous circular muscles of the body wall musculature and the longitudinal retractor muscles appear at the same time in the early larva. In addition, throughout development newly formed ring muscles emerge along the entire anterior- posterior axis by from already existing myocytes. In contrast to that, neurogenesis does follow a segmental pattern by subsequently emerging pairs of serotonergic perikarya along the anterior-posterior axis, which are associated with a paired ventral nerve cord and are arranged in four distinct repetitive units. This metameric pattern, however, disappears prior to and the ventral nerve cords fuse to form the single ventral nerve of the adult. Congruently, the distribution pattern of mitotic cells show similarities to an annelid-like posterior growth zone that also disappears in metamorphic competent pelagosphera larvae. Accordingly, these developmental and morphological data corroborate recent molecular analyses and show that sipunculans stem from a segmented ancestor. Furthermore, the loss of segmentation in these relatively large, free living animals indicates that body segmentation may easier be lost during evolution than previously assumed. The ontogenetic establishment and (secondary) loss of segmentation renders Sipuncula ideal for developmental and evolutionary studies on the segmentation process in Metazoa.

11 Short abstract

Short abstract In this PhD thesis, three sipunculan species were investigated by immunocytochemistry in conjunction with confocal laserscanning microscopy and 3D reconstruction software, in order to clarify whether or not cryptic segmentation can be found during sipunculan ontogeny. The results show that sipunculan myogenesis does not follow a segmental manner, but for a short period of time neurogenesis and the distribution of mitotic cells show transitional stages of segmentation during sipunculan development, thus supporting a sipunculan/annelid affiliation. Moreover, the establishment of an in situ hybridisation protocol for the model sipunculan Themiste pyroides for gene expression analyses pave the way for future studies on the molecular processes underlying sipunculan segmentation that might give important insights into the evolution of segmentation.

General introduction 12

General introduction Annelids and are highly successful animals that have a “segmented” body plan. Therefore, understanding the origin of segmentation as well as the mechanisms of segment formation and diversification has been of scientific interest for decades. Segmented animals exhibit repeated units at the cellular, tissue or organ system level along the anterior-posterior axis (Wilmer 1990, Scholtz 2002). While many animal clades show serial repetition of organs along the anterior-posterior axis (i.g., serially arranged shell plates in polyplacophoran mollusks, ring muscles in platyhelminths, or commissures in numerous worm-shaped invertebrates), only the above mentioned animal clades display the definition of “true” segmentation, i.e., multiple organ systems that are generated successively along the anterior-posterior axis (Couso 2009, Chipman 2010). Recently, our view on animal interrelationships has changed dramatically, splitting them roughly into three mojor lineages (Deuterostomia, , and ) and placing the segmented animals all in separate clades (e.g., Aguinaldo et al. 1997, Halanych 2004, Dunn et al. 2008, Hejnol et al. 2009). Consequently, the “new” animal tree of life has led to two equally possible and hotly debated scenaria: either, the common ancestor of the so-called (bilaterally symmetrical animals) was very simple and segmentation evolved independently in each superclade; or, their last common ancestor was more complex and segmented, with segmentation having been lost in the vast majority of animal clades (Arendt 2005, De Robertis 2008, Couso 2009, Chipman 2010). Accordingly, the monophyly of segmentation is in conflict with the long-standing argument that segmentation is an evolutionary key invention in metazoan (multicellular animals) evolution that is unlikely to get lost (Seaver 2003). Interestingly, recent interpretations of gene expression data on representatives of the segmented phyla have revealed remarkable similarities, proposing a segmented ancestor of all Bilateria (e. g., Arendt 2005, De Robertis 2008, Couso 2009). Since these data largely come from a very limited number of model system animals such as Drosophila (fly), Tribolium (beatle), Capitella (), Hirudo (leech), Cupiennius (spider), and so forth, developmental studies of non-model system taxa are needed in order to either verify or falsify this hypothesis. Moreover, comparative molecular, developmental, and morphogenetic approaches, namely gene expression, cell proliferation, and organ system formation, have proven useful in elucidating the evolution of body patterning 13 General introduction processes (e.g., Hessling and Westheide 2002, Denes et al. 2007, Hejnol and Martindale 2008, Maxmen 2008, Wanninger 2009, Boyle and Seaver 2010, Brinkmann and Wanninger 2010a, b).

Sipuncula – a neglected taxon with annelid affinities Sipuncula is a small, worm-shaped, exclusively marine lophotrochozoan taxon that uniformly exhibits an unsegmented adult body, which is subdivided into a posterior trunk and a retractable anterior introvert. Sipunculans are filter or deposit feeders that live in soft sediments, rock crevices, dead corals, or vacant mollusc shells, and are widespread throughout the oceans (Rice 1975a). Their fossil record is generally sparse but a description of six well preserved sipunculan specimens found in southwest dates their origin to more than 520 million years ago (Huang et al. 2004). Furthermore, the fossilised outer and inner morphology resembles extant sipunculan species, suggesting only limited changes since the Early Cambrium (Huang et al. 2004). Cladograms based on morphological characters divide sipunculans into two classes, and , comprising four orders and six families (, , , , Themistidae, and Sipunculidae) (Cutler and Gibbs 1985, Cutler 1994). Today, the use of DNA sequence data has changed this morphology-based view of the relationships among the currently recognised 147 species of Sipuncula (Maxmen et al. 2003, Schulze et al. 2005, 2007). The two classes have not been recognised by Maxmen et al. (2003) and Schulze et al. (2005), whereas the Phascolosomatidae was recovered in the by far most comprehensive analysis on sipunculan phylogeny of Schulze et al. (2005). Congruently, all analyses strongly support as the sister group to the remaining Sipuncula, while most of the previously recognised families and orders were not recovered as monophyletic (Maxmen et al. 2003, Schulze et al. 2005, 2007). However, based on morphological characters and DNA sequence data, the monophyly of Sipuncula is strongly supported (Maxmen et al. 2003, Schulze et al. 2005, 2007). In general, sipunculans are dioecious (separate sexes), but there are a few known hermaphroditic and parthenogenetic species (asexual reproduction) (Åkesson 1958, Rice 1970, Rajalu and Krishnan 1969, Pilger 1978, Cutler 1994). Four different, one direct, and three indirect developmental pathways have been described for Sipuncula (Fig. 1; Rice 1975b, c). Since sipunculans share certain developmental features such as spiral , that gives rise to a trochophore larva with an apical General introduction 14 tuft and a ring of cilia used for locomotion, they have commonly been placed within the clade , which, among others, also comprises annelids and mollusks (Jägersten 1972, Rice 1985, Cutler 1994, Rouse 1999).

Figure 1. The four distinct developmental modes in Sipuncula. Development may be direct, whereby the embryo emerges from the egg as a crawling worm, which eventually transforms into the juvenile stage without an intermediate larval form (black line and arrows). In some species the lecithotrophic trochophore, after a brief swimming period, transforms into a vermiform stage succeeded by the juvenile (red line and arrows). In the lecithotrophic indirect developmental pathway the trochophore larva starts to elongate and metamorphoses into a second larval stage, the pelagosphera, which swims in the for a short time, settles, and metamorphoses into a juvenile (blue line and arrows). In the planktotrophic indirect developmental mode the lecithotrophic trochophore transforms into a pelagosphera, which remains in the plankton for up to several months before it settles and metamorphoses into a juvenile (green line and arrows). Drawing modified from Rice 1975b, c.

Until they were considered a distinct taxon among the spiralians, the phylogenetic position of Sipuncula has long been an enigma. Accordingly, they were placed close to cucumbers (holothurian ), as derived annelids, or as an in-group of Gephrea (sipunculans, echiurans, and priapulids), then as relatives of , bryozoans, and (Prosoygii), until developmental studies proposed a close relationship to spiralians (reviewed in Rice 1985). From there on, 15 General introduction they have been variously seen as either a sister taxon to Annelida or to (e.g., Åkesson 1958, Rice 1975b, c, Scheltema 1993, 1996, Cutler 1994, Zrzavy et al. 1998, Giribet et al. 2000). Nowadays, a growing number of morphological and molecular analyses strongly suggest a close relationship to annelids, leaving the only question whether sipunculans cluster within Annelida or constitute their sister taxon (e.g., Wanninger et al. 2005a, Tzetlin and Purschke 2006, Struck et al. 2007, Dunn et al. 2008, Mwinyi et al. 2009, Shen et al. 2009, Sperling et al. 2009, Zrzavy et al. 2009, Dordel et al. 2010, Hausdorf et al. 2010, Struck et al. 2011). Accordingly, the inclusion of Sipuncula within Annelida suggests a secondary loss of a previously segmented body plan in Sipuncula rather than an initial evolutionary step towards segmentation in these animals (Dordel et al. 2010, Struck et al. 2011).

Thesis objectives In the light of the proposed sipunculan-annelid assemblage, it was the aim of the present PhD thesis to elucidate sipunculan body plan patterning and to deal with the following scientific issues:

1. Neuro- and myogenesis were described in three different sipunculan species, Phascolosoma agassizii, Themiste pyroides and Thysanocardia nigra (Fig. 2). Since nervous and muscle systems follow a segmental development in annelids, these experiments should clarify whether or not sipunculans show cryptic segmentation during their ontogeny. 2. The distribution of proliferating cells in T. pyoides and T. nigra were described throughout development to assess whether sipunculans possess a posterior growth zone similar to the segmented annelids. 3. Comparative analysis of the acquired developmental data with those available for annelids and other lophotrochozoans (e.g., Mollusca) were carried out in order to infer shared and possible ancestral neuromuscular features. 4. The establishment and application of an in situ hybridisation protocol for analyses of tempo-spatial expression patterns of selected candidate genes known to be involved in body patterning (e.g., neural, muscular and so-called “segmentation” genes (hox1-9, even-skipped, hairy)).

Thesis objectives 16

In the following, the main results are summerised and a general dicussion is presented, dealing with these key issues of the present thesis. More detailed discussions are found in the chapters II to IV, which comprise three published manuscripts. Finally, future perspectives of this project are proposed.

Figure 2. Adults of investigated sipunculan species in the present PhD study. All in lateral view. , which are at the top, mark the anterior end of animals. A. Phascolosoma agassizii, total length approximately 15 cm. B. Themiste pyroides, total length approx. 10 cm. C. Thysanocardia nigra, total length approx. 12 cm.

Material and methods In the present PhD project, several sipunculan species were investigated by developmental-morphological markers such as immunolabelling, confocal microscopy, and 3D reconstruction software (Fig. 2). In addition, developmental stages of Themiste pyroides were used for cloning genes known to be involved in the formation of segmentation in arthropods and annelids, and for establishment of an in situ protocol for gene expression analyses (see below). An overview of the species investigated by morphological methods is given in Table 1. Further details of the respective techniques are provided in the chapters II to IV.

17 Material and methods

Table 1. List of species investigated in the course of the PhD project by fluorescence markers; Serotonin and FMRFamide – neurotransmitters/peptides; Phalloidin – F-actin of the musculature; Dapi (4’, 6-diamindino-2-phenylindole) – cell nuclei marker, EdU (5-ethynyl-2’-deoxyuridine) – proliferating cells.

Species (Family) Neurogenesis Myogenesis Cell nuclei, cell proliferation Themiste pyroides Serotonin, F-actin Dapi, EdU (Themistidae) FMRFamide (chapter IV) (chapter IV) (chapter I) Thysanocardia Serotonin, F-actin Dapi, EdU nigra (Golfingidae) FMRFamide (chapter IV) (chapter IV) (chapter I) Phascolosoma Serotonin, F-actin - agassizii FMRFamide (chapter IV) (Phascolosomatidae) (chapters I-III)

RNA isolation and cDNA synthesis Total RNA was purified from Themiste pyroides embryos at 15, 28, 40, 48, 62, 72, 84, and 111 hours post fertilizaton (hpf) (miRCURY RNA isolation kit, Exiqon, Vedbaek, Denmark). cDNA samples were synthesised by reverse transcription (RETROsript, Ambion, Woodward St. Austin, TX, USA), and stored at -20 °C until use.

Degenerate PCR To clone homeobox genes, a range of degenerate primers were designed referring to Martinez et al. (1997), Nederbragt et al. (2002), Seaver et al. (2006), and Paps et al. (2009). The sequences of the oligonucleotides used are given in Table 2. A touchdown PCR was performed with the degenerate primers given in Table 2. The amplification parameters were: 3 min at 94 °C, 10 cycles of 45 sec at 94 °C, 45 sec at 52 °C (every cycle -1 °C), 30 sec at 72 °C, and 30 cycles of 45 sec at 94 °C, 45 sec at 42 °C, and final 30 sec extension at 72 °C. The PCR products were purified by a gel extraction kit (QIAquick, QIAGEN, Copenhagen, Denmark), and on 1 µl of this reaction another PCR with the same parameters was performed. The samples were displayed by electrophoresis on a 1% agarose gel, purified, concentrated by speedvacuum (GENEVAC EZ-2plus, Ipswich, UK), resuspended in 10 µl distilled water, and directly ligated into pGEM-T Easy vector using a ligation kit (Promega, Material and methods 18

Nacka, Sweden). The inserted DNA fragments were sequenced using BigDye Terminator 3.1 Cycle Sequencing kit with an ABI Prism 377 DNA sequencer (Applied Biosystems, Carlsbad, CA, USA). Degenerate primers were also designed to clone a number of muscular and neural genes for establishment of a whole mount in situ hybridisation protocol (Table 2). Subsequently, all recovered gene fragments were identified by the similarity of their nucleotide sequences to already known genes that are placed at the public online source of the National Centre for Biotechnology Information (NCBI) using basic local alignement searches (BLASTs).

Table 2. Nucleotide sequences of degenerate primers and their melting temperatures (Tm) used to clone homeobox, muscular and neural genes in Themiste pyroides.

Primers Sequence Tm Homeobox genes (Martinez et al. 1997) CT 77 5’-CGGATCCYTIGARYTIGARAARGART 42°C CT 78 5’-GGAATTCATICKRTTYTGRAACCAIATTY Engrailed (Seaver et al. 2006, Nederbragt et al. 2002) EnEH2 5’-TGGCCTGCITGGGTNTAYTGYAC 42 °C en2-2 5’-TGRTTRTANARNCCYTGNGCCAT Eng1 5’-ATGGAATTCCNGCNTGGGTNTWYAC 42 °C Eng4 5’-TGGAAGCTTRTANARNCCYTSNGSCAT Myosin heavy chain type II (Ruiz-Trillo et al. 2002) Mio7 (F) 5’-TGYATCAAYTWYACYAAYGAG 53 °C Mi6 (R) 5’-CCYTCMARYACACCRTTRCA Tropomyosin (Paps et al. 2009) TropoF 5’-ATYRAGAAGAARATGNBKGVCATG 53 °C TropoR 5’-GTHYGRTCCARTTGNYCACT Intermediate filaments (Paps et al. 2009) FilF1 5’-TACATCGAGAAGGTGCGTTTCCTGG 48 °C FilR1 5’-CCTCACCCTCCAGCAGCTTTCTGTA FilF3 5’-TACATCGAGAAGGTGCGTTTCCTGG 50 °C FilR3 5’-CYTCNCCYTCCAGCAGYTTYCTGTA β-Actin (Matsuo and Shimizu 2006) 19 Material and methods

β-Actin F1 5’-TGGGAYGAYATGGARAARAT 50 °C β-Actin R1 5’-GCCATYTCYTGYTCRAA α-Tubulin (Zheng et a. 1998) AT16-F 5’-ATHGGHAAYGCNTGYTGGG 50 °C AT412-R 5’-RAAYTCDCCYTCYTCCATDCC β-Tubulin (Garant and MacRea 2009) BT-F 5’- 55 °C GGNCARTCNGGNGCNGGNAAYAAYTGGGCN BT-R 5’-NGGRAANGGNACCATRTTNACNGC

Rapid amplification of cDNA ends (RACE) Fragments of Themiste pyroides homeobox containing genes were isolated by degenerate PCR using cDNA of mixed larval stages as template. In addition, 5’ and 3’ cDNA were generated using the same template for rapid amplification of cDNA ends (RACE) with the SMARTer RACE cDNA amplification kit (Clonetech, Mountainview, CA, USA). For each of the recovered gene fragments, which ranged between 135 and 271 base pairs (bp) (Table 3), gene specific primers (GSP) were designed and used for RACE (Advantage 2 PCR Kit; Clonetech). Touchdown PCRs were performed with following amplification parameters: 5 min at 94 °C, 10 cycles of 45 sec at 94 °C, 45 sec at 57 °C (every cycle -1 °C), 3 min at 72 °C, and 25 cycles of 45 sec at 94 °C, 45 sec at 47 °C, and final 30 sec extension at 72 °C. Reactions were agarose-gel electrophoresed, stained with SYBRsafe (Invitrogen, Taastrup, Denmark), and photographed under UV light. Subsequently, 1 µl of RACE PCR products were used as a template for another RACE PCR with nested GSPs (see Table 3). The parameters were: 5 min at 94 °C, 35 cycles of 45 sec at 94 °C, 45 sec at 60 °C, 30 sec at 72 °C, and a final 10 min extension at 72 °C. Positive fragments were purified from the agarose gel, cloned into the vector pGEM-T easy, and sequenced as described above. However, I was able to amplify into the 3’ direction of two genes only, hox1 (47 bp) and hox8 (73 bp).

Material and methods 20

Table 3. Isolated DNA fragments of Themiste pyroides mixed larval stages. Gene abbreviations: lab, labial; scr, sex combs reduced; ftz, fushi tarazu; abd-A, abdominal-A. Other abbreviations: bp, base pairs; e-value, expected value (the similarity between a given sequence and orthologues from the database; high similarities are indicated by considerably low e-values, while high e-values indicate low similarity). Red and underlined parts within the sequences indicate gene specific and nested primers, respectively.

Gene Sequence bp/e-value hox1 CGGATCCCTGGAACTGGAGAAAGAATTCCACTTTA 136/10-10 ACAAATATCTCACAAGAGCCAGGCGGATAGAAATA (lab) GCTGCTGCACTTGGACTAAATGAAACACACAGGTT AAGATCTGGTTTCAGAACAGCCGCATGAATTCC hox3 GGATCCTTGGAACTGGAGAAGGAATTCCATTTTAAC 135/10-14 CGTTACTTGTGTCGGCCACGCCGTATTGAAATGGCA GCCATGCTCAACCTGACAGAAAGACAGATCAAGAT CTGGTTTCAGAATAGCAGCATGAATTCC hox5 GGATCCCTGGAACTGGAGAAAGAATTTCATTACAA 135/10-14 CAGATACCTAACAAGACGAAGGAGAATAGAAATAG (scr) CACATGCTTTGGGACTCACGGAACGCCAAATTAAG ATCTGGTTTCAGAATAGCAGCATGAATTCC hox5 GGATCCTTGGAATTGGAAAAAGAATTTCACTTCAAC 135/10-12 CGTTACTTGTCCAGCAAACGACGTACAGAGATAGC (ftz) CGAGTCACTGGCTCTGACAGATAGACAAGTTAAGA TCTGGTTTCAAAACAGCCGCATGAATTCC hox8 CGGATCCTTGGAATTGGAAAAAGAATTTCAGTTTAA 136/10-15 TCATTATTTAACCCGAAAAAGGAGAATAGAGATAG (abd-A) CGCACGCTTTGTGTCTAACAGAAAGACAGATAAAG ATCTGGTTTCAGAATCGCAGCATGAATTCC even- CGGATCCTTGGAGTTGGAAAAAGAATTTTACAGAG 252/10-9 AGAACTATGTTAGCAGACCAAGGAGGTGTGAACTG skipped GCTGCGGAACTCAACCTGCCAGAATCAACAATCAA GGTAAGAAGAAAATAGTGCGACCATTTATCACAGT TTCAGAGCCAACATTTTCAAAGTCGATTTTTCTAAT TTGGAATAAAATTGCAATAAATTAATGTTTTATTTT TTACTTTCAGATCTGGTTTCAGAACAGCAGCATGAA TTCC lox2 CGGATCCTTGGAGCTGGAGAAAGAATTCAAATTCA 136/10-14 ACAGATACTTAACTCGCAGAAGACGAATAGAACTG TCTCATATGTTGTGCTTAACTGAGCGCCAAATCAAG ATCTGGTTTCAGAACAGCAGCATGAATTCC caudal GGAATTCATGCTGCTGTTCTGAAACCAGATTTTGAC 136/10-12 CTGTCTCTCAGACAGACTGAGTGACTGTGCTAGTTC (cdx) TGCCTTTCTTCTGATTGTGATATAACGACTGTAATG GAATTCTTTCTCCAGTTCCAGGGATCCG xlox CGGATCCCCTGGAATTGGAAAAAGAATTTCATTTTA 137/10-14 ATAAATATATCTCCAGACCAAGGAGAATAGAATTA GCTGCCATGTTAAATCTAACAGAGAGACATATAAA GATCTGGTTTCAGAATAGCAGCATGAATTCC not CGGATCCTTGGAGTTGGAGAAAGAATTTCACTTCAA 271/10-11 CCGTTACTTGTCCAGCAAACGACGTACAGAGATAG 21 Material and methods

CCGAGTCACTGGCTCTGACAGATAGACAAGTTAAG ATCTGGTTTCAGAACAGCAGCATGAATTCCGGATCC CTGGAATTGGAGAAGGAATTCGAAAGGCAACAATA CATGGTTGGATCTGAAAGGTATTATCTGGCAGTATC GCTTAATTTATCCGAATCCCAAGTGAAGATCTGGTT TCAGAACCGCCGCATGAATTCC

Whole mount in situ hybridisation Embryos, larvae, and juveniles of Themiste pyroides were anesthetised by adding

drops of a 3.5 or 7% MgCl2 solution to the seawater, and were subsequently fixed in 4% paraformaldehyde (PFA) in 0.1 M phosphate-buffered saline (PBS) (pH 7.3) for 20-30 min at room temperature. Subsequently, the fixative was removed by three washes in 70% ethanol (15 min each), followed by the storage of the specimens at -20 °C. Fixed specimens were rehydrated in PBS, washed four times for 5 min in PTw (phosphate-buffer containing 0.1% Tween), and permeabilised with proteinase K (Sigma; 10 µg/ml in PTw for 15 min at room temperature). Digestion was terminated by two washes (5 min each) with PTw containing 2mg/ml of glycine and treated with triethanolamine (TEA, 0,1M pH 7.6, Fluka; 3 x 5 min), following addition of 4µl/ml and 8µl/ml of acetic anhydride without changing the TEA solution to block positive charges. After two 5 min washes in PTw, specimens were post-fixed in 3.7% PFA for 1 hr at room temperature. Subsequently, specimens were rinsed 5 x 5 min in PTw, incubated for 10 min in 50% pre-hybridisation buffer (50% formamide, 5x SSC, 1mg/ml yeast RNA, 0.1 mg/ml heparin, 0.1% Tween 20, 10mM DTT) in PTw and an incubation in 100% pre-hybridisation buffer overnight at 65 °C. Antisense and sense digoxigenin-labelled riboprobes were generated with a RNA labeling kit (SP6/T7; Roche; Copenhagen, Denmark), and used at a working concentration of 1 ng/µl for Tp-mhc and Tp-actin (fragment sizes ca. 700 and 500 bp, respectively). Riboprobes were warmed up in hybridisation buffer (50% formamide, 5x SSC, 0.1% Tween 20) for 10 min at 70 °C. Afterwards, specimens were added to that solution and incubated for 72 hr at 65 °C. After hybridisation, probes were recovered and specimens were washed at 65 °C (30 min in 100% hybridisation buffer, in 75% hybridisation buffer and 75% 2x SSC, in 50% hybridisation buffer and 50% 2x SSC, in 25% hybridisation buffer and 25% 2x SSC, 2 x 15 min in 2x SSC, and final 2 x 15 min washed in 0.2x SSC). Afterwards, specimens were washed two times 5 min in maleic buffer (MABTw; 100 mM maleic acid, 150 mM NaCl, 0.1% Tween 20, pH 7.5), blocked for Material and methods 22

1 hr in 1% blocking solution (Roche) and incubated overnight with anti-digoxigenin alkaline phosphatase (AP) conjugated antibody (Roche; 1:200 dilution) in blocking solution at 4 °C. Specimens were washed at least four times 15 min in MABTw and two times 5 min in AP buffer (0.1M NaCl, 0.1M Tris pH 9.5, 0.05M MgCl2, 0.5% Tween 20) and developed with NBT/BCIP (Roche) in the dark. The reactions were stopped with PTw, specimens were fixed in 3.7% PFA overnight at 4 °C and stored in 80% glycerol in PBS at -20°C. Specimens were analysed and photographed using DIC optics on a Zeiss Axiophot microscope (Zeiss, Jena, Germany) in conjunction with a Leica DFC300FX digital camera (Leica, Microsystems, Wetzlar, Germany).

Results In the course of the present PhD study, three different sipunculan species, which cover three out of six currently recognised families, were investigated. Phascolosoma agassizii was collected and reared at the Friday Harbor Laboratories on the San Juan Island (Washington, USA) whereas Themiste pyroides and Thysanocardia nigra were acquired at the Vostok-Marine-Station (Vladivostok, Russia). The muscle and nervous system development is described for all investigated species, the distribution of proliferating cells for T. pyroides and T. nigra, and cloning of homeobox genes as well as the establishment of an in situ hybridisation protocol for T. Pyroides. The main findings are summarised in the following section. Further details are given in the chapters II – IV.

Cloning of homeobox genes in Themiste pyroides With the pair of degenerate primers CT77 and CT78, which were designed to the conserved regions within the homeobox, the orthologues for the hox1 (labial), hox3, hox5 (sex combs reduced), hox5 (fushi tarazu), hox8 (abdominal-A), even-skipped, lox2, caudal, xlox, and not were isolated. The isolated gene fragments of Themiste pyroides cDNA were generated from mixed embryonic and larval stages; their lengths were between 135 and 271 bp (Table 3). Unfortunately, gene specific primers and RACE PCRs were not able to extent the recovered fragments neither in 5’ nor in 3’ direction. The only exceptions were the genes hox1 (labial) (47 bp) and hox8 (abdominal-A) (73 bp), which included the stop codon and a poly-A tail, but the 23 Results obtained fragments were too short to generate riboprobes for successful in situ hybridisation experiments.

Establishment of an in situ hybridisation protocol Using a set of degenerate primers (Table 2), gene fragments of 700 bp of myosin heavy chain (Tp-mhc) and 500 bp of beta-actin (Tp-actin) were recovered by PCR. Tp-mhc is expressed from early trochophore stages (ca. 2 days after fertilisation (dpf)) onwards in the four developing retractor muscles (Fig. 3A-C). First Tp-actin expression is also in the early trochophore larvae in a prominent U-shaped band in the area of the forming retractor muscles (Fig. 3D). In pelagosphera larvae the Tp-actin is expressed in distinct areas of the paired dorsal and ventral retractor muscles (Fig. 3E, F). Results 24

Figure 3. Expression of myosin heavy chain (Tp-mhc) (A-C) and beta-actin (Tp-actin) (D-F) during larval stages of Themiste pyroides. Anterior is to the top in all panels and scale bars equal 50 µm. Black represents areas of gene expression – Tp-mhc in A-C and Tp-actin in D-F, respectively. (A) Dorsal view of an early trochophore larva showing Tp-mhc expression (arrowheads) in the anlagen of the retractor muscles. (B) Dorsal view of a late trochophore larva. Tp-mhc expression is restricted to the retractor muscles. Note that only three out of four are in focus; mt marks the ciliated metatroch. (C) Lateral view, ventral to the right, pelagosphera larva with Tp-mhc expression in the retractor muscles; pt marks the ciliated prototroch. (D) Dorsal view of an early trochophore larva with Tp-actin expression in the area of the retractor muscles (arrows). (E) Dorsolateral view of an early pelagosphera larva, ventral to the left, showing Tp-actin expression in the retractor muscles. (F) Late pelagosphera larva, dorsal view with strong Tp-actin expression in the retractor muscles.

25 Results

Serotonergic and FMRFamidergic nervous system development in sipunculan worms (chapters I – III) In the planktotrophic species Phascolosoma agassizii, the first detectable serotonergic and FMRFamidergic nervous structures are visible in the anterior region of the trochophore larvae. A pair of neurites emerging from the apical organ contributes to the ventral nerve cords. During further development, interconnecting commissures and pairs of perikarya (four serotonergic and one pair of FMRFamidergic perikarya) appear subsequently along the ventral nerve cords from anterior to posterior, resulting in a rope-ladder like ventral nervous system. Due to migration of serotonergic perikarya and fusion of serotonergic ventral nerve cords this metameric arrangement is lost towards metamorphosis, and gives rise to the single non-metameric adult ventral nervous system (chapters II and III). In the lecithotrophic species Themiste pyroides and Thysanocardia nigra, neurogenesis is similar to Phascolosoma agassizii (Figs. 4-6). The first serotonergic and FMRFamidergic cells are in the apical organ and a pair of neurites contributes to the ventral nerve cords (Figs. 4A, 6A). In contrast to P. agassizii, a first, second, and a third pair of serotonergic perikarya appear already in the trochophore larvae of T. pyroides and T. nigra in an anterior to posterior progression, while later the fourth pair appears in the pelagosphera larva (Figs. 4A-D, 5E, H). Results 26

Figure 4. Confocal micrographs of the serotonergic nervous system in trochophore larvae of Themiste pyroides. A and C are in ventrolateral left view, whereas B and D are in ventral view. Anterior faces upwards and scale bars equal 50 µm in all aspects except the insert in B where it is 10 µm. Age of larvae is given in days post fertilisation (dpf). (A) Early trochophore larva (2 dpf) showing an apical organ comprising three cells (arrowheads), two neurites of the developing ventral nervous system (vn) that is interconnected by a commissure (arrow), and a pair of perikarya (asterisks). (B) Slighly later (2 dpf), at the base of the apical organ (ao), the first cells of the adult cerebral organ are formed (cg), the two ventral neurites are stronger developed than in the stage before, and the first perikarya of the second pair appears. Insert is a magnification of the apical pole of B showing two flask-shaped and two round cells in the apical organ. (C) Trochophore larva (2.5 dpf) with two pairs of perikarya associated with the ventral neurite bundles and the first perikarya of the third pair already formed. (D) Slightly later (2.5 dpf) the second perikarya of the third pair is established posteriorly to the second pair. Note that the cerebral ganglion is more elaborated than in the stage before.

During subsequent development a second pair of FMRFamidergic perikarya appears along the ventral nervous system in T. pyroides and T. nigra, while there is only one in P. agassizii (Fig. 6D). All three species show four serotonergic and two to three FMRFamidergic flask-shaped cells in their apical organ, as well as a serotonergic nerve ring underlying the ciliated metatroch (Figs. 4A, B, Insert, 5A, B, 27 Results

E, F, I-L). It has to be noted that in T. pyroides and T. nigra the serotonergic metatrochal nerve ring is considerebly weaker than that in P. agassizii (Fig. 5A, E, I- L).

Figure 5. Confocal micrographs of the serotonergic nervous system in pelagosphera larvae of Themiste pyroides. All aspects in ventral view except I, K, J and L, which are in ventro lateral left view. Anterior faces upwards and scale bars represent 50 µm in A, E, I, K, J and L, while they equal 10µm in B, C, D, F, G and H. A, E, I, and K are color-coded confocal micrographs, where blue and purple colors are Results 28

positioned to the ventral, colors of green and yellow in between, whereas orange and red colors are to the dorsal side. Age of larvae is given in days post fertilisation (dpf). (A) Early pelagosphera larva (3 dpf) showing a weakly stained metatrochal neurite (mtn), ventral neurite bundles (vn) that are interconnected by commissures (arrows) always anteriorly positioned to a pair of perikarya (asterisks); cg marks the cerebral ganglion. (B) Dorsal portion of the same specimen as in A; magnification of the apical pole showing four cells (arrowheads) in the apical organ. Note that two cells are flask-shaped. (C) Ventral portion of the same specimen as in A; magnification of the apical pole showing numerous cells associated with the neuropile of the developing adult cerebral ganglion. (D) Magnification of the ventral trunk area of an early pelagosphera larva (3 dpf) with three pairs of metamerically arranged perikarya associated with the ventral neural bundles. (E) Slighly older pelagosphera (4 dpf) as in A with stronger developed ventral neural bundles that bear four pairs of perikarya and a more elaborate cerebral ganglion. (F) Dorsal portion of the same specimen as in E; magnification of the apical pole showing five cells in the apical organ. (G) Ventral portion of the same specimen as in E, magnification of the apical pole showing numerous cells and a neuropile in the cerebral ganglion. (H) Same specimen as in E. Magnification of the ventral trunk area showing the ventral nervous system with four pairs of perikarya and three commissures interconnecting the median with the two outer neural bundles. (I) Magnification of the anterior region of a pelagosphera larva (8 dpf) showing a complex central nervous system that comprises the cerebral ganglion with numerous cells, the apical organ with its cells and own neuropile (double arrows), peripheral cells (double arrowheads), metatrochal neurite, peripheral neurites (pn), and the ventral neural bundle. Note that the ventral neural bundle has retained its paired character only in the most anterior part. (J) Dorsal portion of the same specimen as in I showing the apical organ with its cells and neuropile. Note that two of the cells are flask-shaped. (K) Magnification of the anterior region of a late pelagosphera larva (14 dpf) with a prominent cerebral ganglion, apical organ with two flask-shaped cells, peripheral cells and neurites, a prominent metatrochal neurite ring, and ventral neural bundles. (L) Dorsal portion of the same specimen as in K showing the apical organ that comprises less cells than in the previous stage. Three out of five cells disappeared while only the two flask-shaped cells and a part of the neuropile remain visible.

In addition to the loss of the metameric arrangement in the ventral nervous system towards metamorphosis, less serotonergic and FMRFamidergic apical cells are detectable in all of the investigated species, while the adult cerebral ganglion becomes more elaborate (Figs. 5I-L, 6D).

29 Results

Figure 6. Confocal micrographs of FMRFamidergic nervous system in pelagosphera larvae of Thysanocardia nigra. All aspects in ventral view except B and C which are in ventrolateral left view. Anterior faces upwards and scale bars represent 50 µm except the insert in A where it equals 30 µm. Age of larvae is given in days post fertilisation (dpf). (A) Early pelagosphera larva (3 dpf) with two flask-shaped cells in the apical organ (arrowheads), the anlage of the adult cerebral ganglion (cg), and a pair of ventral neurites (vn). Insert shows a slightly older stage (3 dpf), with the first interconnecting commissure (arrow) and pair of perikarya (asterisks) associated with the ventral neural bundles. Note the presence of a median ventral neurite. (B) Pelagosphera larva (4 dpf) showing a more elaborate cerebral ganglion and ventral nervous system that now has a second commissure that interconnects the ventral neurites; pn marks peripheral neurite. (C) Slighlty older pelagosphera larva (5 dpf) with a well Results 30

developed cerebral ganglion below the apical cells and ventral nervous system. (D) Late pelagosphera larva (11 dpf) with numerous peripheral neurites in the trunk area as well as in the ventral nervous system. Note the second pair of perikaya along the ventral neurites. No FMRF-positive cells appear in the apical organ.

Myogenesis and cell proliferation during sipunculan development (chapter IV) Myogenesis in all investigated species in the present PhD thesis display a similar mode of muscle system development. In Phascolosoma agassizii, Themiste pyroides and Thysanocardia nigra, the rudiments of the four longitudinal retractor muscles appear at the same time as the first circular body wall muscles. Throughout subsequent development, novel circular body wall muscles appear along the entire anterior-posterior axis and seem to be formed by fission from existing myocytes. In addition, all three sipunculans show loosely arranged longitudinal body wall muscles, except in the vicinity of the prominent retractor muscles. In contrast to P. agassizii, T. pyroides and T. nigra have no terminal organ and, therefore, no respective retractor muscle. Furthermore, the anlagen of the buccal musculature appear much earlier in P. agassizii (trochophore larva) than in T. pyroides and T. nigra (late pelagosphera, shortly before the onset of metamorphosis). However, in none of these species does myogenesis follow an annelid-like metameric development. Cell proliferation was described in Themiste pyroides and Thysanocardia nigra during development from early embryos until early juvenile stages by visualisation of the nucleotide analogue 5-ethynyl-2´-deoxyuridine (EdU) that is incorporated during the synthesis phase (S-phase) of the cell cycle. There is a similar distribution of S-phase cells during the development of both investigated species. First, the proliferating cells are scattered throughout the ectoderm. At the beginning of anterior-posterior axis elongation, proliferating cells are distributed in two lateral bands around the eyes, around the mouth, and the ventral trunk region. Slightly later, in the early pelagosphera larvae most S-phase cells appear in the ventral nerve cord and the rudiments of the digestive system. Interestingly, in the pelagosphera stage the proliferation is only present in the posterior tip of the ventral trunk area and in the venral nerve cord. EdU-positive cells are arranged in units and by intensity loss it seems as if the direction of proliferation is from posterior to anterior. As the larvae grow, more S-phase cells appear in the digestive system, around the , and in the 31 Results area of the developing tentacles of post metamorphic stages. However, no S-phase cells can be detected in the posterior tip of the ventral trunk region anymore.

General discussion Immunocytochemistry and F-actin labelling in conjunction with confocal microscopy and 3D reconstruction software has proven to be an excellent tool to characterise the neuromuscular system as well as the expression pattern of certain neurotransmitters, thus allowing for comparative analyses of neural and muscular characters (Wanninger 2009, Richter et al. 2010). Using these methods, the main goal of the present PhD thesis was to characterise nervous and muscle system formation as well as the distribution of proliferating cells during sipunculan development. Additional studies on myo- and neurogenesis in other sipunculans supplement this comparative approach (Wanninger et al. 2005a, Schulze and Rice 2009). So far, eight species representing four families and three different developmental modes have been investigated by the above mentioned methods.

Early neurogenesis in Trochozoa In all investigated sipunculan species neurogenesis begins in the trochophore larvae at the apical pole, from which two neurites grow posteriorly, forming a scaffold for the future ventral nervous system (Wanninger et al. 2005a; chapters I and II herein). Moreover, the investigated species herein, Phascolosoma agassizii, Themiste pyroides, and Thysanocardia nigra, all show initially two and later up to four serotonergic and FMRFamidergic flask-shaped cells within the apical organ (chapters I-III), whereas Phascolion strombi lacks serotonergic apical cells (Wanninger et al. 2005a). The most common developmental pathway in sipunculans is via a lecithotrophic trochophore and a planktotrophic pelagosphera (Fig. 1), which might remain pelagic for several weeks or even months (Scheltema and Hall 1975, Rice 1981, 1985). Compared to other sipunculans, Phascolion strombi has a relatively short larval development (approx. 3 days) (Åkesson 1958, Wanninger et al. 2005a). Accordingly, the lack of serotonergic expression in the apical organ might be a secondary condition. However, the above described pattern of early neurogenesis found in sipunculans is remarkably similar to that reported for the early trochophore larvae of the Polygordius lacteus, Pomatoceros lamarkckii, Sabellaria General discussion 32 alveolata, and Spirorbis cf. spirorbis (Hay-Schmidt 1995, McDougall et al. 2006, Brinkmann and Wanninger 2008, 2009). In sipunculans, congruently with annelids, mollusks, and nemerteans, the adult cerebral ganglion starts to form at the base of the apical organ (Croll and Dickinson 2004, Wanninger et al. 2005a, Brinkmann and Wanninger 2008, 2009, 2010a, Chernyshev and Magarlamov 2010, Fischer et al. 2010, Kristof and Klussmann-Kolb 2010; chapters I and II herein).

Establishment and loss of segmentation during sipunculan neurogenesis Adult sipunculans exhibit a single ventral neurite bundle that lacks ganglia. However, in all investigated sipunculans to date, a paired serotonin- and FMRFamide-positive ventral neurite bundle with interconnecting commissures appears during ontogeny (Wanninger et al. 2005a; chapters I-III herein). Furthermore, four pairs of serotonergic perikarya associated with the ventral neurite bundles and three serotonergic and FMRFamidergic commissures appear subsequently in a strict anterior-posterior progression during sipunculan ontogeny, leading to a rope ladder- like ventral nervous system (chapters I-III). Interestingly, the FMRFamidergic ventral nervous system exhibits a median neurite bundle between the two ventral neurites and one (Phascolosoma agassizii) or two perikarya (Themiste pyroides, Thysanocardia nigra), which appear in the same manner as the serotonergic ones subsequently on the outer ventral neurites (chapters I and II). A similar condition with a median neurite bundle in the ventral nervous system has been described for a number of polychaete, myzostomid, echiuran, as well as hirudinean larvae and adults (Sawyer 1986, Müller and Westheide 2000, 2002, Eeckhaut et al. 2003, Orrhage and Müller 2005, McDougall et al. 2006, Hessling 2002, Hessling and Westheide 2002, Hessling 2003). Accordingly, the segmental mode of neurogenesis, which indicates the existence of a posterior growth zone, and a median neurite bundle clearly link sipunculans to annelids, thus strongly supporting a segmented ancestry of both taxa. Unique to sipunculans, the metameric arrangement of the ventral nervous system is lost towards metamorphosis (chapters I-III). The serotonergic ventral neurite bundles fuse, their commissures disappear, and the metameric arrangement of the perikarya is lost by the time a fifth pair appears (chapters I-III). The two FMRFamidergic ventral neurite bundles that form the ventral nervous system come to lie closer to each other prior to metamorphosis, and more neurites appear between them, resulting in a single, non-metameric ventral nerve cord in the adult stage 33 General discussion

(chapters I-III). Accordingly, this is the first report of the ontogenetic establishment and loss of a metamerically arranged organ system in the nervous system of a metazoan animal.

Comparative trochozoan neurogenesis An early neurogenesis that starts with a paired ventral neurite is common for all annelids (i.g., including myzostomids, sipunculans and echiurans sensu Mwinyi et al. 2009 and Struck et al. 2011) as well as many lophotrochozoans, suggesting two ventral neurites to be part of the body plan of the last common ancestor of Lophotrochozoa (Wanninger 2009, Chernyshev and Magarlamov 2010, Fischer 2010; chapters I, and II herein). The sipunculan species investigated herein all have a serotonergic and FMRFamidergic neurite that underlies the metatrochal ciliary bands, which constitute the primary locomotory organ of the larvae (chapters I-III). However, it has to be noted that the immunoreactivity against both neuronal markers of the metatrochal neurite in Themiste pyroides and Thysanocardia nigra is much weaker than in Phascolosoma agassizii, and even absent in Phascolion strombi (Wanninger et al. 2005a; chapters I-III herein). This is probably due to the different life history patterns, since the pelagosphera larvae of P. agassizii are the ones that feed and have the longest development in the plankton. This might explain why their locomotory organ is well developed and innervated. T. pyroides and T. nigra both have lecithotrophic, short-lived pelagosphera stages (10-14 days at 17-19 °C), while this is even more reduced in P. strombi (pelagosphera stage for approx. 12 to 24 hours at 12-16 °C). However, a neurite underlying a ciliated locomotory organ is also found in other trochozoan larvae including polychaetes (Hay-Schmidt 1995, Voronezhskaya et al. 2003, McDougall et al. 2006, Brinkmann and Wanninger 2008, 2009, Fischer et al. 2010), mollusks (Friedrich et al. 2002, Croll et al. 1997, Kempf et al. 1997), nemerteans (Lacalli and West 1985, Maslakova 2010), ectoprocts (Wanninger et al. 2005b, Gruhl 2009), phoronids (Hay-Schmidt 1990a, b), and various (Nielsen and Hay-Schmidt 2007), although the expression of neural markers in this neurite varies. Hence, this condition might be an adaptation of marine larvae to swimming and feeding during their planktonic dispersal phase. A possible apomorphy of Lophotrochozoa might be an apical organ containing serotonergic and/or FMRFamidergic flask-shaped cells (Wanninger 2009). Similar to General discussion 34 the sipunculan species investigated herein, the polychaete Phyllodoce maculate exhibits four flask-shaped serotonergic and FMRFamidergic cells within the apical organ (Voronezhskaya et al. 2003; chapters I-III herein). In addition, some mollusks (but not the comparatively basal Polyplacophora with 8-10 flask-shaped apical cells), ectoprocts, nemerteans, and brachiopods share this feature of up to four immunoreactive flask-shaped cells within the apical organ (Hinman et al. 2003, Croll and Dickinson 2004, LaForge and Page 2007, Voronezhskaya et al. 2008, Dyachuk and Odintsova 2009, Gruhl 2009, Wanninger 2009, Altenburger and Wanninger 2010, Chernyshev and Magarlamov 2010, Kristof and Klussmann-Kolb 2010), thus indicating that the last common lophotrochozoan ancestor probably had an apical organ comprising approximately four serotonergic flask-shaped apical cells. Taken together, sipunculan neurogenesis shares numerous features with annelid larvae. These include early neurogenesis with an apical organ from which two neurite bundles are formed in posterior direction, giving rise to the ventral nervous system as well as the metatrochal and oral nerve ring (chapters I-III). Moreover, the formation of the ventral nerve cord follows a segmental pattern, whereby perikarya and commissures associated with the ventral nerve cord are formed one after another in an anterior to posterior progression (chapters I-III). This supports recent molecular phylogenetic analyses and a segmental ancestry of Sipuncula (chapters I-III).

Sipunculan growth zone(s) Annelid segment formation is defined by the presence of a posterior growth zone from which the segmentally arranged organs are progressively budded off, leading to the anterior-posterior progression of developing organ systems such as the nervous and muscle system (Iwanonff 1928, Anderson 1966, 1973, Weisblatt et al. 1988). Interestingly, the sipunculans Themiste pyroides and Thysanocardia nigra show a metameric pattern of mitotic cells in their development that originates from the ventral posterior trunk area just dorsal to the roots of the retractor muscles, and seem to progress from posterior to anterior (chapter IV). Congruently with sipunculan neurogenesis (chapters I-III), this metameric pattern disappears in metamorphic competent larvae (chapter IV). Furthermore, these findings are strikingly similar to the description of a growth zone in sipunculan larvae that only remains for a short period of time (Åkesson 1958). 35 General discussion

While Åkesson (1958) described the embryology and neurogenesis in Phascolion strombi, he also reported a growth zone that is situated anterior to the retractor roots. In addition, he showed that with the exception of P. strombi the growth zone in sipunculan larvae remains in this anterior position for only a part of their life cycle. However, in P. strombi the introvert retractors are attached at the very posterior end of the body. Moreover, during the juvenile stages the left degenerates and the ventral and dorsal retractors fuse, with the latter becoming more prominent, while the ventral retractor often degenerates (Åkesson 1958). P. strombi has a short larval development as well as fusion and degeneration processes in the first juvenile stages. Accordingly, similar to Åkesson’s description, a putative growth zone in T. pyroides and T. nigra is only present in their pelagosphera larvae prior to metatmorphosis (chapter IV). However, the question remains whether and how long such a putative growth zone may exist during ontogeny of P. strombi or Phascolosoma agassizii, since both display different developmental pathways than T. pyroides and T. nigra. Recently, ontogenetic studies showed that segmentation in annelids is more variable than previously thought (Seaver et al. 2005, Brinkmann and Wanninger 2008, 2010b). For instance, the segments in the polychaetes Capitella and Hydroides are formed from a ventrolateral region of the body, while a posterior growth zone is only evident in post-metamorphic stages (Seaver et al. 2005). This corroborates the data on another polychaete, Sabellaria alveolata, where no larval posterior growth zone appears (Brinkmann and Wanninger 2010b). Moreover, this variability of segment formation is also mirrored in the nervous and muscle system formation in Sabellaria alveolata, where certain parts develop synchronously and others subsequently in an anterior to posterior manner (Brinkmann and Wanninger 2008, 2010b). Clearly, more ontogenetic studies, preferebly on supposedly basal polychaetes such as Dinophilidae, Oweniidae, and Protodrilida, are needed to shed more light on segment formation in Annelida (including Sipuncula).

Myogenesis in sipunculans and annelids In all sipunculans investigated so far, myogenesis follows a similar pattern. Four introvert retractor muscles begin to develop together with a considerable number of circular muscles (Wanninger et al. 2005a, Schulze and Rice 2009; chapter IV herein). The planktotrophic species Phascolosoma agassizii and Nephasoma pellucidum General discussion 36 exhibit an additional retractor muscle for the terminal organ and the anlagen of the buccal musculature (Schulze and Rice 2009; chapter IV herein). This is probably due to the longer planktotrophic stage, since all other investigated species have either direct or indirect lecithotrophic development, hence develop the buccal organ considerably later (Wanninger et al. 2005a, Schulze and Rice 2009; chapter IV herein). However, the simultaneous establishment of circular body wall muscles along the entire anterior-posterior axis throughout sipunculan ontogeny seems to be independent from the life cycle pathway, suggesting that this mode of myogenesis may be ancestral to sipunculans (Wanninger et al. 2005a, Schulze and Rice 2009, Wanninger 2009; chapter IV herein). By contrast, the circular body wall muscles in annelids develop in a segmental manner from anterior to posterior, although circular muscles are not always present in polychaetes (Hill and Boyer 2001, Seaver et al. 2005, Tzetlin and Filippova 2005, Purschke and Müller 2006, Bergter et al. 2007, Hunnekuhl et al. 2009, Wanninger 2009, Fillipova et al. 2010). Furthermore, comparative analyses on polychaete larvae suggest that the body wall of the last common annelid ancestor probably had four longitudinal muscle strands that develop from anterior to posterior (Purschke and Müller 2006, Bergter et al. 2008, Brinkmann and Wanninger 2010b). From early development onwards, sipunculans exhibit four longitudinal retractor muscles (Wanninger et al. 2005a, Schulze and Rice 2009; chapter IV herein), which in post-metamorphic stages might fuse (Åkesson 1958). Moreover, longitudinal body wall muscles are always densely arranged in the areas of the retractor muscles and more loose towards the mid-body, thus suggesting that the retractor muscles evolved from fused longitudinal body wall muscles (chapter IV). Interestingly, one dorsal and one ventral pair of longitudinal body wall muscles appear first in polychaetes and oligochaetes, which develop then in anterior to posterior direction (McDougall et al. 2006, Bergter et al. 2007, 2008, Brinkmann and Wanninger 2010b, Fischer et al. 2010). Accordingly, sipunculan and annelid larvae exhibit both four separate longitudinal muscle strands that develop from anterior to posterior, and are considered having been present in their last common ancestor (e.g., Cutler 1994, Schulze and Rice 2009, Brinkmann and Wanninger 2010b). This again supports the recent phylogenetic analyses that consider the sipunculans as in-group annelids (e.g., Zrzavy et al. 2009, Struck et al. 2011).

37 Conclusions and future perspectives

Conclusions and future perspectives Despite the fact that circular body wall muscles do not develop in a segmental manner, the data presented herein document for the first time traits of segmentation in Sipuncula. For a short period of time during sipunculan ontogeny, a repeated pattern is generated from the posterior pole of the larvae in their nervous system, which resembles the annelid-like mode of segmentation. In agreement with recent molecular phylogenetic analyses, this strongly argues for a segmented sipunculan and annelid last common ancestor, which had four longitudinal body wall muscles that developed from anterior to posterior. However, the absence of circular body wall muscles in a number of adult polychaetes and their larvae casts doubt on the assumption that ring muscles were part of the annelid ground pattern, although multiple, independent loss of these muscles in the respective lineages are also possible. Accordingly, the different pathways of body segmentation in annelids (including sipunculans, echiurans, and myzostomids) might be due to adaptations to different modes of life, thus indicating that segmentation is more complex as well as labile to evolutionary changes than previously assumed. The fact that a segmented body is established and later secondarily lost during ontogeny renders sipunculans an ideal group to study the ontogeny and evolution of segmentation, in particular in comparison to the well studied annelid, and model organisms. This might help to shed light on the contradicting conclusions on morphological urbilaterian characteristics (i.e., a complex, segmented versus a simple, non-segmented species at the base of the bilaterian tree of life). In this context, one primary aim of future studies would be to deepen the knowledge on the “segmentation” process in Sipuncula from a molecular perspective. The way segments are specified in insects, especially Drosophila, is different from the way segments are controlled in or annelids (Chipman 2010). Therefore, knowing which genes are involved in sipunculan segmentation, which unlike the above mentioned model organisms looses its segmentation during ontogeny, can be a good objective for understanding the evolution of segmental specification and loss. Modern high-throughput sequencing technologies such as 454 reads or illumina are able to generate a large EST dataset of almost all genes expressed during the development of an animal. Accordingly, the analysis of the EST dataset and the gene expression of so-called “segmentation” genes (i.e., hairy, notch, delta, wingless, Conclusions and future perspectives 38 engrailed, even-skipped) will provide important insights into the molecular processes that underlie the establishment and loss of segmentation during sipunculan ontogeny. The developmental-morphological study herein provides an anatomical framework that enables detailed interpretation of gene expression patterns in sipunculans. As shown in the present thesis, the sipunculan Themiste pyroides is ideally suited as a model sipunculan for these kinds of studies in the future.

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47 Chapter II

Chapter II

Kristof A , Wollesen T & Wanninger A. 2008. Segmental mode of neural patterning in Sipuncula. Current Biology 18 : 1129-1132

Current Biology 18, 1129–1132, August 5, 2008 ª2008 Elsevier Ltd All rights reserved DOI 10.1016/j.cub.2008.06.066 Report Segmental Mode of Neural Patterning in Sipuncula

Alen Kristof,1 Tim Wollesen,1 and Andreas Wanninger1,* posterior axis of the animal proceeds, a second pair of ventral 1Department of Biology perikarya arises (Figure 1B). In the following stages, the entire Research Group for Comparative Zoology neural architecture of Phascolosoma is elaborated, resulting in University of Copenhagen a complex anterior nervous system that shares numerous fea- Copenhagen DK-2100 tures with annelid larvae. These include a (larval) serotonergic Denmark prototroch nerve ring, a (larval) apical organ comprising sev- eral serotonergic cells, the early anlage of the adult brain, and an oral nerve ring (Figures 1C and 1D). Most notable and Summary important, however, is the successive formation of two addi- tional pairs of serotonergic perikarya associated with the sero- Recent molecular phylogenetic analyses suggest a close tonergic axons of the VNCs, together with three serotonergic relationship between two worm-shaped phyla, the nonseg- and FMRFamidergic commissures that interconnect the mented Sipuncula (peanut worms) and the segmented Anne- respective axons of the VNC (Figure 1B, inset, and Figures lida (e.g., earthworms and polychaetes) [1–5]. The striking 1C–1F). This results in a segmental central nervous system differences in their bodyplans are exemplified by the anne- (CNS) comprising four metamerically arranged pairs of seroto- lids’ paired, ladder-like ventral nervous system, which con- nergic perikarya along the VNCs of the late-stage Phascolo- tains segmentally arranged ganglia, and the sipunculans’ soma larva (Figure 2A and Figure 3). Interestingly, only one single ventral nerve cord (VNC), which is devoid of any seg- pair of FMRFamidergic perikarya is found to be associated mental structures [6, 7]. Investigating central nervous sys- with the FMRFamidergic VNC (Figure 1F). tem (CNS) formation with serotonin and FMRFamide labeling Annelid segmentation is not typified simply by the repetitive in a representative sipunculan, Phascolosoma agassizii,we arrangement of organs along the longitudinal body axis but, found that neurogenesis initially follows a segmental pattern more specifically, by their successive formation mode in similar to that of annelids. Starting out with paired FMRFami- a strict anterior-posterior progression; this indicates the pres- dergic and serotonergic axons, four pairs of associated ence of a posterior growth zone from which the segmentally serotonergic perikarya and interconnecting commissures arranged organs are budded off [9–13]. With the increasing form one after another in an anterior-posterior progression. molecular evidence for an annelid-sipunculan assemblage In late-stage larvae, the two serotonergic axons of the VNCs [1–5], the question of whether the last common ancestor fuse, the commissures disappear, and one additional pair of (LCA) of both taxa was segmented or not is still open because perikarya is formed. These cells (ten in total) migrate toward within Lophotrochozoa (animals with a ciliated larval stage in one another, eventually forming two clusters of five cells their life cycle), a segmented nervous system had been known each. These neural-remodeling processes result in the sin- only in annelids, including echiurans (spoon worms, a group of gle nonmetameric CNS of the adult sipunculan. Our data marine worms believed to nest within the annelid phylum) [1–5, confirm the segmental ancestry of Sipuncula and render 12, 14, 15]. Our findings fill this gap in knowledge by providing Phascolosoma a textbook example for the Haeckelian hy- the hitherto-lacking proof of a segmented ancestry of the si- pothesis of ontogenetic recapitulation of the evolutionary punculan nervous system [16], thus strongly arguing in favor history of a species [8]. of a segmented annelid-echiuran-sipunculan LCA.

Results and Discussion Ontogenetic and Evolutionary Loss of the Segmental Sipunculan CNS Ontogenetic Establishment of the Segmented The fully developed segmented nervous system of the larva of Sipunculan CNS Phascolosoma agassizii comprises four pairs of serotonergic Despite their proposed close phylogenetic relationship [1–5], perikarya and three serotonergic and FMRFamidergic com- annelids and sipunculans differ significantly from each other missures interconnecting the VNCs (Figures 1D, IF, 2A, and in that most annelids exhibit a typical segmented body with 3D). As the larva approaches metamorphic competence, the metamerically arranged appendages, commissures intercon- two serotonergic axons of the VNCs fuse, the serotonergic necting two ventral nerve cords (VNCs), and paired sets of commissures disappear, and a fifth pair of serotonergic peri- ganglia, whereas sipunculans lack any trace of metamerism karya is formed (Figure 2B). Together with subsequent cellular and have only a single VNC. Using antibodies against the migratory processes, this eventually results in a CNS consist- neurotransmitters serotonin and FMRFamide, we found that ing of a single serotonergic VNC, which only retains its paired neurogenesis in Phascolosoma starts in the anterior region character in the anterior-most region where it connects to the with the larval apical organ and the subsequently developing brain, as well as two neural cell clusters comprising five peri- paired serotonergic and FMRFamidergic axons of the VNC. karya each (Figures 2C and 3E). Accordingly, the segmental Slightly later, one pair of serotonergic perikarya associated serotonergic neural architecture is lost prior to the onset of with the serotonergic portion of the VNCs appears metamorphosis of the Phascolosoma larva. At this stage, the (Figure 1A). As development and growth along the anterior- three FMRFamidergic commissures are still discernable, al- though the two FMRFamidergic axons of the VNCs lie in much closer proximity to each other than in the previous larval *Correspondence: [email protected] stages (Figure 1F). Current Biology Vol 18 No 15 1130

Figure 1. Ontogeny of the Serotonergic and FMRFamidergic Nervous System in the Sipuncu- lan Phascolosoma agassizii Anterior faces upward and scale bars represent 30 mm in all aspects. (A) Early larva expressing four cells in the apicalor- gan (indicated by arrowheads), one pair of seroto- nergic axons in the ventral nerve cord (Svnc) with one commissure (indicated by an arrow), and the first pair of perikarya (marked by asterisks), which is associated with the ventral nerve cord. (B) Slightly older larva with two pairs of perikarya, three commissures, and a weak signal in the future oral nerve ring (indicated by a double arrow). Note that fusion of the two serotonergic axons of the ven- tral nerve cord has already begun in the posterior part of the animal. The large open arrow indicates the fusion point of the serotonergic axons. The Svnc is fused in the entire region posterior to this point. The inset shows a close-up of a transitional stage with two commissures (indicated by arrows). (C) Larva with three pairs of ventral perikarya and elaborated apical organ (ao) with underlying rudi- ment of the future cerebral ganglion (cg) (‘‘brain’’). The continuing fusion of the serotonergic axons of theventralnervecordresultsinonlyoneremaining commissure. (D) Late-stage larva with four pairs of meta- merically arranged perikarya along the almost- completely-fused serotonergic axons of the ven- tral nerve cord. As in (C), only the first commissure in the anterior part of the larva has been retained. ‘‘ptr’’ indicates the serotonergic nerve ring under- lying the prototroch. (E) Larva with paired FMRFamidergic axons of the ventral nerve cord (Fvnc) emerging from the rudi- ment of the adult cerebral ganglion and expressing the first commissure. (F) Late larva with three FMRFamidergic commissures (indicated by arrows) and one pair of FMRFamidergic perikarya (marked by asterisks). Note that the two ventral nerve cords have begun to migrate toward each other. ‘‘pn’’ indicates the peripheral nervous system.

Initially paired ventral serotonergic nerves that fuse during In many annelids, ganglia and commissures are commonly development have recently been described for another sipun- formed after the establishment of the first rudiments of the culan, Phascolion strombi. Although this species did not show paired VNC. Development of these commissures and ganglia unambiguous signs of segmentation during its ontogeny (a is strictly directional and follows an anterior-posterior pro- fact that is probably due to its shortened larval phase), the gression, a fact that is typically assigned to the presence of transitional occurrence of three FMRFamidergic commissures a posterior growth zone [9, 10, 24, 25], and that eventually in late P. strombi larvae indicates cryptic remnants of a once- leads to the typical ladder-like metameric annelid ventral segmented CNS in this species as well [17]. CNS. Although rather obvious in ‘‘typical’’ (e.g., polychaete) The findings of earlier morphological and developmental annelids, indications for the segmental ancestry of the studies proposing a taxon comprising Sipuncula and Mollusca echiurans are much more subtle and were only revealed re- [18] have recently been rejected by a number of molecular anal- cently by studies of their neural development [12, 14, 15]. yses that clearly argue for an annelid-echiuran-sipunculan As such, echiurans exhibit a double-stranded VNC in early clade [1–5, 16]. Accordingly, novel data on CNS formation in larval stages similar to the annelids [12]. Although these these groups cast new light on the evolution of annelid seg- nerve cords fuse during subsequent development, the asso- mentation. Although the architecture of the adult annelid CNS ciated perikarya form in anterior-posterior succession until seems to be quite plastic, ranging from one to five VNCs [19], the juvenile neural bodyplan is established [12, 14, 15]. neurogenesis consistently starts with an anterior neuropil (the Thus, although modified, the segmental origin of the echiuran predecessor of the brain) and one pair of serotonergic and CNS can still be recognized in the adult stage by the meta- FMRFamidergic axons in the VNCs, irrespective of the number meric arrangement of the perikarya associated with the of VNCs that are present in the adult stage [20–23]. This is con- VNC. Moreover, cell-proliferation markers indicate an anne- gruent with the findings on echiuran and sipunculan neurogen- lid-like posterior growth zone in Bonellia viridis, thus further esis and suggests that a paired ventral CNS was also present in strengthening the evidence for the segmented origin of the LCA of these three groups. However, various kinds of inde- echiurans [15]. In sipunculans, reduction of neural segmenta- pendent fusion and duplication events have taken place during tion traits has progressed even further than in echiurans, evolution, accounting for the phenotypic plasticity of the adult resulting in a CNS that is devoid of any metameric structures CNS that is encountered in these worm-shaped animals today. in the adult stage (Figures 2C and 3E). Segmental Neurogenesis in Sipuncula 1131

Figure 2. Loss of the Metameric CNS Pattern during Ontogeny of the Serotonergic Nervous System in Late Phascolosoma Larvae Anterior faces upward and scale bars represent 15 mm in all aspects. The total size of the speci- mens is approximately 300 mm in length. (A) Late larva with four pairs of metameric peri- karya (marked by asterisks) and progressively fused serotonergic axons of the ventral nerve cord (Svnc). The large open arrow indicates the fusion point of the serotonergic axons. The Svnc is fused in the entire region posterior to this point. (B) Appearance of two additional cell bodies and start of disintegration of the paired metameric arrangement of the perikarya. (C) Disintegration of the metameric arrange- ment of the perikarya is completed, resulting in two clusters of five perikarya each (boxed areas).

the segmental origin of Sipuncula. Ac- cordingly, Phascolosoma confirms the Haeckelian notion that cryptic character states, which have been lost during evo- lution in the adult bodyplan, may have been conserved during ontogeny [8], To our knowledge, this work for the first time documents thus stressing the importance of developmental studies for ontogenetic establishment and loss of CNS segmentation in the reconstruction of ancestral bodyplan features of metazoan the life cycle of a recent metazoan and demonstrates clades.

Supplemental Data

Supplemental Data include Supplemental Experimental Procedures and two movies and can be found with this article online at http://www. current-biology.com/cgi/content/full/18/15/1129/DC1/.

Acknowledgments

We thank Bernard M. Degnan, Danny Eibye-Jacobsen, Gerhard Haszpru- nar, Claus Nielsen, and Christiane Todt for comments on an earlier version

Figure 3. Summary Figure of Establishment and Secondary Loss of the Segmental Serotonergic Neural Bodyplan in Phascolosoma as Depicted by 3D Reconstructions of Selected Confocal Microscopy Data Sets from Figures 1 and 2 Anterior faces upward and scale bars represent 15 mm in all aspects except for (A), in which it equals 7.5 mm. The total size of the specimens is approx- imately 150 mm in (A), (C), and the inset, 250 mm in (B), and 300 mm in (D) and (E). ‘‘a-p’’ indicates the anterior-posterior axis of the animals. (A) Early larva with paired serotonergic axons of the ventral nerve cord (Svnc), the first pair of perikarya (marked by asterisks), and the first commis- sure (indicated by an arrow). (B) Larva with two pairs of perikarya. The large open arrow indicates the fu- sion point of the serotonergic axons. The Svnc is fused in the entire region posterior to this point. The inset shows the transitional stage with two com- missures established. (C) Larva with three pairs of perikarya. Continued fusion of the serotonergic axons of the ventral nerve cord has begun. Note the prominent commissure in the anterior region. (D) Fully established metameric nervous system with four pairs of perikarya present. (E) The metameric character of the nervous system has been lost, and the ten perikarya are now arranged in two distinct clusters containing five cells each (boxed areas). Current Biology Vol 18 No 15 1132

of the manuscript, as well as members of the Wanninger lab for discussion. bands of Pomatoceros lamarckii (Annelida): Heterochrony in poly- The comments of three anonymous reviewers helped to improve the manu- chaetes. Front. Zool. 3, 16. script. A.K. is the recipient of an EU fellowship within the MOLMORPH net- 22. Denes, A.S., Je´ kely, G., Steinmetz, P.R.H., Raible, F., Snyman, H., work under the 6th Framework Programme, ‘‘Marie Curie Host Fellowships Prud’homme, B., Ferrier, D.E.K., Balavoine, G., and Arendt, D. (2007). for Early Stage Research Training (EST)’’ (contract number MEST-CT-2005 – Molecular architecture of annelid nerve cord supports common origin 020542). T.W. is funded by a PhD fellowship from the University of Copen- of nervous system centralization in Bilateria. Cell 129, 277–288. hagen. The research of A.W. is supported by grants from the Danish Re- 23. Brinkmann, N., and Wanninger, A. (2008). Larval neurogenesis in Sabel- search Agency (21-04-0356 and 271-05-0174) and the Carlsberg Foundation laria alveolata reveals plasticity in polychaete neural patterning. Evol. (2005-1-249). Author contributions are as follows: A.K. performed research, Dev., in press. T.W. acquired the study material, A.K. and A.W. analyzed data, and A.W. 24. Anderson, D. (1966). The comparative morphology of the Polychaeta. designed research and wrote the paper. Acta Zool. 47, 1–42. 25. Anderson, D. (1973). Embryology and Phylogeny in Annelids and Arthro- Received: April 23, 2008 pods (Oxford: Pergamon Press). Revised: May 29, 2008 Accepted: June 18, 2008 Published online: July 24, 2008

References

1. McHugh, D. (1997). Molecular evidence that echiurans and pogonopho- rans are derived annelids. Proc. Natl. Acad. Sci. USA 94, 8006–8009. 2. Boore, J.L., and Staton, J.L. (2002). The mitochondrial genome of the si- punculid supports its association with Annelida rather than Mollusca. Mol. Biol. Evol. 19, 127–137. 3. Halanych, K.M., Dahlgren, T.G., and McHugh, D. (2002). Unsegmented an- nelids? Possible origins of four lophotrochozoan worm taxa. Integr. Comp. Biol. 42, 678–684. 4. Struck, T.H., Schult, N., Kusen, T., Hickman, E., Bleidorn, C., McHugh, D., and Halanych, K.M. (2007). Annelid phylogeny and the status of Sipuncula and Echiura. BMC Evol. Biol. 7, 57. 5. Dunn, C.W., Hejnol, A., Matus, D.Q., Pang, K., Browne, W.E., Smith, S.A., Seaver, E., Rouse, G.W., Obst, M., Edgecombe, G.D., et al. (2008). Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452, 745–749. 6. Brusca, R.C., and Brusca, G.J. (2003). Invertebrates, Second Edition (Sunderland, MA: Sinauer Associates). 7. Ruppert, E.E., Fox, R.S., and Barnes, R.D. (2004). Invertebrate Zoology, Seventh Edition (Belmont, CA: Thomson, Brooks/Cole). 8. Haeckel, E. (1874). Anthropogenie (Leipzig: Engelmann). 9. Weisblat, D.A., Price, D.J., and Wedeen, C.J. (1988). Segmentation in leech development. Development. Suppl. 104, 161–168. 10. Shankland, M. (1991). Leech segmentation: Cell lineage and the forma- tion of complex body patterns. Dev. Biol. 144, 221–231. 11. Davis, G.K., and Patel, N.H. (1999). The origin and evolution of segmen- tation. Trends Cell Biol. 12, M68–M72. 12. Hessling, R., and Westheide, W. (2002). Are Echiura derived from a seg- mented ancestor? Immunohistochemical analysis of the nervous system in developmental stages of Bonellia viridis. J. Morphol. 252, 100–113. 13. De Rosa, R., Prud’homme, B., and Balavoine, G. (2005). Caudal and even-skipped in the annelid Platynereis dumerilii and the ancestry of posterior growth. Evol. Dev. 7, 574–587. 14. Hessling, R. (2002). Metameric organisation of the nervous system in developmental stages of Urechis caupo (Echiura) and its phylogenetic implications. Zoomorphology 121, 221–234. 15. Hessling, R. (2003). Novel aspects of the nervous system of Bonellia viridis (Echiura) revealed by the combination of immunohistochemistry, confocal laser-scanning microscopy and three-dimensional recon- struction. Hydrobiologia 496, 225–239. 16. Chipman, A.D. (2008). Annelids step forward. Evol. Dev. 10, 141–142. 17. Wanninger, A., Koop, D., Bromham, L., Noonan, E., and Degnan, B.M. (2005). Nervous and muscle system development in Phascolion strombus (Sipuncula). Dev. Genes Evol. 215, 509–518. 18. Scheltema, A.H. (1993). Aplacophora as progenetic aculiferans and the coelomate origin of mollusks as the sister taxon of Sipuncula. Biol. Bull. 184, 57–78. 19. Mu¨ ller, M.C.M. (2006). Polychaete nervous systems: Ground pattern and variations – cLS microscopy and the importance of novel characteristics in phylogenetic analysis. Integr. Comp. Biol. 46, 125–133. 20. Voronezhskaya, E.E., Tsitrin, E.B., and Nezlin, L.P. (2003). Neuronal de- velopment in larval polychaete Phyllodoce maculata (Phyllodocidae). J. Comp. Neurol. 455, 299–309. 21. McDougall, C., Chen, W.C., Shimeld, S.M., and Ferrier, D.E.K. (2006). The development of the larval nervous system, musculature and ciliary Chapter III 52

Chapter III

Wanninger A, Kristof A & Brinkmann N. 2009. Sipunculans and segmentation. Communicative & Integrative Biology 2: 56-59

[Communicative & Integrative Biology 2:1, 56-59; January/February 2009]; ©2009Sipunculans Landes Bioscience and segmentation

Mini-Review Sipunculans and segmentation

Andreas Wanninger,* Alen Kristof and Nora Brinkmann

University of Copenhagen; Department of Biology; Research Group for Comparative Zoology; Copenhagen, Denmark Key words: Sipuncula, evolution, segmentation, seriality, annelid, Echiura, nervous system, development, phylogeny, bodyplan

Comparative molecular, developmental and morphogenetic While often considered an important hint towards ancestral analyses show that the three major segmented animal groups— segmentation of a species, serial repetition of organs along the ante- Lophotrochozoa, Ecdysozoa and Vertebrata—use a wide range of rior-posterior axis alone is not decisive for a segmental evolutionary ontogenetic pathways to establish metameric body organization. history (cf., e.g., the multiple ring muscles in the non-segmented Even in the life history of a single specimen, different mecha- platyhelminths). On the cellular and organ system level, segmen- nisms may act on the level of gene expression, cell proliferation, tation can only be proven with the aid of developmental studies, tissue differentiation and organ system formation in individual because segmented animals typically exhibit a posterior growth zone segments. Accordingly, in some polychaete annelids the first three from which all segments are progressively budded off.3-7 Accordingly, pairs of segmental peripheral neurons arise synchronously, while ontogenetically older segments—and thus also the organs associated the metameric commissures of the ventral nervous system form in with them—are found anterior to the younger segments, a fact that anterior-posterior progression. Contrary to traditional belief, loss is illustrated by the gradual decrease of the degree of organ system of segmentation may have occurred more often than commonly differentiation from anterior to posterior (Fig. 1).8 This makes the assumed, as exemplified in the sipunculans, which show remnants pattern of organogenesis an ideal marker to test for the segmental 8-12 of segmentation in larval stages but are unsegmented as adults. ancestry of worm-shaped lophotrochozoan taxa. The developmental plasticity and potential evolutionary lability of Coelomic compartmentalization of a cylindrical body has segmentation nourishes the controversy of a segmented bilaterian frequently been proposed to be of selective advantage due to the fact ancestor versus multiple independent evolution of segmentation in that these animals are able to regulate the hemolymphic pressure in respective metazoan lineages. each compartment (segment) individually. The interplay of coelomic pressure and the contractile ring and longitudinal muscles of the Ontogeny and Functional Implications of Segmentation body wall enable direct and independent control over the diameter of the body in each individual segment, thus allowing for a diversity The evolution of a segmented bodyplan is often considered a of complex movement patterns.2 However, while coelomic segmen- crucial metazoan innovation because it allows the subdivision and tation has often (but not always) been retained in large, burrowing specialization of individual body regions along the anterior-posterior annelids (e.g., earthworms), secondary loss is often observed in non- axis of an animal.1,2 This partitioning typically involves both the benthic free-living (e.g., leeches), interstitial (e.g., Protodrilus), or ectodermal and the endodermal germ layers and often results sessile forms (e.g., tube worms). in metameric ectodermal appendages (parapodia) and segmen- tally arranged, paired mesodermal body cavities (coelomic sacs). Loss of Segmentation Traditionally, a condition where all segments have the same type of parapodia and house identical sets of internal organs such as Despite the loss of segmentation in various annelid taxa, ontoge- ganglia, nephridia, gonads and muscles has been regarded as basal netic remnants of their segmented ancestry are present in a number 1,2 of annelids that do not show obvious segmental features in the adult for annelids and arthropods (homonomic segmentation). From 3,4 this basal (plesiomorphic) condition, concentration of individual body (e.g., leeches). Recent developmental studies have shown organ systems into segments of specific body regions combined that this holds also true for representatives of the Sipuncula (peanut worms), unsegmented lophotrochozoans that are regarded either with reductions of organs in other segments are thought to have 13-17 occurred multiple times within various lineages, and eventually led to as derived ingroup annelids or as a direct annelid sister clade. morphologically distinct segments along the anterior-posterior body Interestingly, however, segmental traits in the sipunculans are axis (heteronomic segmentation).1,2 restricted to the nervous system but have been completely lost on the level of organisation.18 Accordingly, larvae of Phascolosoma agassizii develop four pairs of perikarya that are associated with the *Correspondence to: Andreas Wanninger; University of Copenhagen; Department of Biology; Research Group for Comparative Zoology; Universitetsparken 15; paired ventral nerve cord and express the common neurotransmitter Copenhagen DK-2100 Denmark; Email: [email protected] serotonin (Fig. 2A). These paired perikarya form, together with commissures that interconnect the ventral nerve cords, in a typical Submitted: 11/26/08; Accepted: 12/01/08 annelid-like anterior-posterior progression, thus demonstrating Previously published online as a Communicative & Integrative Biology E-publication: the segmental ancestry of Sipuncula. During subsequent larval http://www.landesbioscience.com/journals/cib/article/7505

56 Communicative & Integrative Biology 2009; Vol. 2 Issue 1 Sipunculans and segmentation

­development, the ventral nerve cords fuse and the paired perikarya migrate towards each other, resulting in two cell clusters of five cells each, which are grouped around the now single ventral nerve cord (Fig. 2B). This results in loss of the metameric neural pattern and eventually in the establish- ment of the non-segmented ventral nervous system with only one single nerve cord.18 Noticeably, within Sipuncula the degree of preservation of neural segmentation appears to be dependent on the duration of the larval phase and is thus directly corre- lated with the basal versus the derived mode of sipunculan development. The segmented nervous system in Phascolosoma is expressed in the so-called pelagosphera larva. This larval stage is a defining (apomorphic) character for Sipuncula and is thus consid- ered part of the ancestral sipunculan life cycle.19 Neurogenesis in another sipun- culan species that lacks the pelagosphera stage, Phascolion strombi, shows that the remnants of the metameric nervous system have been reduced even further. As such, while also having a primarily paired ventral Figure 1. Schematic representation of neurogenesis in the polychaete annelid Sabellaria alveolata based on serotonin immunoreactivity, revealing differences in the mode of establishment of metamery in the nerve cord, this species lacks the associ- peripheral segmental neurons and the ventral commissures, respectively. Both aspects are ventral views ated perikarya and only has retained three with anterior facing upwards. Total length of the specimens is approximately 280 μm in (A) and 330 transitional ventral commissures as the sole μm in (B). (A) Late larva with synchronously established peripheral segmental neurons (yellow). Ventral remnants of the ancestral segmented neural commissures and perikarya along the paired ventral nerve cord (vnc) are still lacking. The prototroch bodyplan.20 nerve ring (pnr) and the nerve ring underlying the telotroch (ttn) constitute subsets of the larval nervous system, while the circumoesophageal commissures (cc) and the longitudinal trunk neurons (ltn) are parts Cryptic segmentation in taxa with of the adult neural bodyplan. (B) Larva prior to metamorphosis. The ventral commissures (asterisks) of close annelid affinities seems to be more the first five segments have been established progressively, together with the paired, metameric sets of common than previously assumed. The perikarya (red dots) along the ventral nerve cords (vnc). The six pairs of peripheral segmental neurons echiurans (spoon worms), now considered (yellow) correspond to the segments II–VII, because development of segment I is retarded in this species, as clustering within Annelida,13,16,17 do resulting in development of the paired peripheral segmental neuron of this segment at a later stage. Note that ontogeny of the peripheral segmental neurons precedes development of the ventral commissures in not show any segmental traits in their adult segments VI and VII. pns – the nerves of the peripheral nervous system. gross morphology. However, neurogenesis revealed the same ontogenetic mechanisms as found in annelids and sipunculans, namely that paired sets of Plasticity of Segmentation perikarya are formed in a discrete anterior-posterior progression.9-11 In contrast to the sipunculans and similar to the condition found Despite the long standing definitions concerning the charac- in “typical” annelids, this metameric organization of the nervous teristics of a segmented bodyplan (see above), recent data have system persists in the adult echiurans. Accordingly, the annelid-echi- shown that the ontogenetic establishment of annelid segmentation uran-sipunculan lineage shows a gradual decrease of preservation may follow quite different developmental pathways. Traditionally, of nervous system segmentation, a notion that is further supported it had been proposed that the first three (larval) segments form by patterns of myogenesis. Hereby, annelids exhibit the typical more or less synchronously by schizocoely from the paired lateral anterior-posterior progression of ring and dorsoventral muscle mesodermal band, while the following (adult) segments develop formation, while sipunculan myogenesis starts with synchronous from a pre-anal growth zone.21 Accordingly, one would assume that formation of early ring muscle rudiments, followed by the emer- the organ systems associated with the first three segments also arise gence of additional ring muscles along the entire anterior-posterior synchronously, while only the subsequent segmental organs follow axis by fission from already existing myocytes.8,20 Accordingly, the anterior-posterior differentiation gradient. However, this is only Sipuncula represents a developmental mosaic of segmental and partly true for the polychaete Sabellaria alveolata. While the three non-segmental bodyplan patterning mechanisms, whereby the larval segments indeed arise synchronously in this species, only the ectoderm-derived nervous system has to some degree maintained corresponding pairs of peripheral segmental neurons form synchro- its segmental ancestry while the mesodermal musculature is formed nously, while the ventral commissures develop subsequently one entirely non-metamerically. after another (Fig. 1).22 While this may be interpreted as (secondary)

www.landesbioscience.com Communicative & Integrative Biology 57 Sipunculans and segmentation

Ancestry of Segmentation Comparative analyses of the develop- mental mechanisms that form metameric organs in segmented lophotrochozoan worms demonstrate a high plasticity of ontogenetic patterns that lead to a segmented bodyplan and show that segmentation may be lost during evolu- tion. This raises the question as to what extent such an evolutionary loss has yet remained unrecognized in other phyla, thus reviving the discussion about a possible segmented ancestor of Lophotrochozoa and Bilateria as a whole. Such a scenario has been repeatedly proposed by the advo- cates of a conserved molecular pathway that is thought to underlie the ontogeny of metazoan segmentation.7,26 However, despite some similarities on the molecular level, there are also significant differences in the way typical “segmentation genes” are expressed, and cellular and tissue differen- tiation pathways that eventually give rise to individual segments vary between lopho- trochozoans, vertebrates and ecdysozoans.25 To complicate matters further, segment formation is highly variable even between phyla within the respective “superclades” Ecdysozoa and Annelida.25 Moreover, Figure 2. Expression and loss of the segmental pattern of the ventral CNS in the sipunculan Phascolosoma morphologically similar segments may agassizii, as depicted by 3D reconstruction of serotonin immunoreactivity. Both aspects are ventral views follow different ontogenetic pathways even with anterior facing upwards. Total length of the specimens is approximately 150 μm. (A) Late larva within the same individual, and distinct with four pairs of metameric perikarya (red; boxed area) associated with the ventral nerve cord (vnc). metamerically arranged subunits of the The latter is already fused along the entire anterior-posterior axis except in the anterior-most region. Additional neural elements include the cells of the larval apical organ (dark blue) overlying the neuropil nervous system may form differently in mass (np) of the adult brain, the first cell bodies of the developing adult brain (light blue), two cells of individual segments of the same animal the peripheral nervous system (orange), and the larval prototroch nerve ring (green). (B) Larva prior to (e.g., the ventral commissures versus the metamorphosis in which the metameric arrangement of the ventral perikarya (red) has been lost in favor peripheral segmental neurons in poly- of two cell clusters (boxed areas) comprising five cells each. Note the increased number of cells belong- ing to the adult brain (light blue). chaetes; see above). Lastly, a mosaic of segmental and non-segmental modes of chronological dissociation of larval segment formation and the organogenesis may occur within an individual, indicating the occur- development of the ventral commissures, it could alternatively be rence of dissociation of organogenesis from the segmentation process explained as the result of a heterochronic shift of the first three in some species (e.g., muscle formation in sipunculans; see above). segments, which were originally derived from a posterior growth Given the incongruencies and the plasticity of the processes zone, into the larval stage of the animal. This notion is supported involved in the ontogeny of segmentation in the various major bila- by reports that describe an—albeit rapid—progressive formation of terian subgroups, no final statement as to whether or not Urbilateria these first three pairs of coelomic sacs in several polychaete taxa.23 was segmented can yet be made. In any case, assuming a segmented Whatever alternative holds true, this example demonstrates that the urbilaterian would imply a wide range of evolutionary modifications developmental mechanisms that underlie annelid segmentation are of the ancestral segmentation pathway on the molecular, cellular much more complex than previously assumed. This is confirmed by and morphogenetic level, as well as secondary loss of a segmented recent cell proliferation pattern analyses, which suggest that the loca- body in a number of lineages. Both, experimental developmental tion of the growth zone might have shifted from a posterior-median genetics employing RNAi experiments and comparative morphoge- position to both lateral sides in some species, indicating positional netic analyses provide exciting tools that enable us to directly test for variability of the annelid growth zone.24 However, despite the high evolutionary hypotheses concerning shared molecular segmentation morphogenetic plasticity of segmentation, some molecular mecha- pathways on the one hand and for cryptic remnants of a segmented nisms appear similar even between distant phylogenetic entities such bodyplan in seemingly non-segmented recent phyla on the other. as arthropods and vertebrates.25,26 This should eventually lead to a sound reconstruction of the ancestry

58 Communicative & Integrative Biology 2009; Vol. 2 Issue 1 Sipunculans and segmentation

of one of the key innovations of metazoan evolution: the origin of segmented bodies. Acknowledgements Research in the lab of Andreas Wanninger is funded by the EU Early Stage Research Training Network MOLMORPH under the 6th Framework Programme (contract number MEST-CT-2005— 020542). Both Alen Kristof and Nora Brinkmann are recipients of a fellowship within the MOLMORPH programme. References 1. Brusca RC, Brusca GJ. Invertebrates. Sunderland: Sinauer Associates 2003. 2. Ruppert EE, Fox RS, Barnes RD. Invertebrate Zoology. Belmont: Thomson, Brooks/Cole 2004. 3. Weisblat DA, Price DJ, Wedeen CJ. Segmentation in leech development. Development 1988; 104:161-8. 4. Shankland M. Leech segmentation: cell lineage and the formation of complex body pat- terns. Dev Biol 1991; 144:221-31. 5. Davis GK, Patel NH. The origin and evolution of segmentation. Trends Biochem Sci 1999; 12:68-72. 6. Nielsen C. Animal Evolution. Interrelationships of the Living Phyla. Oxford: Oxford University Press 2001. 7. De Rosa R, Prud’homme B, Balavoine G. Caudal and even-skipped in the annelid Platynereis dumerilii and the ancestry of posterior growth. Evol Dev 2005; 7:574-87. 8. Wanninger A. Shaping the things to come: Ontogeny of lophotrochozoan neuromuscular systems and the Tetraneuralia concept. Biol Bull 2009; In press. 9. Hessling R. Metameric organization of the nervous system in developmental stages of Urechis caupo (Echiura) and its phylogenetic implications. Zoomorphology 2002; 121:221-34. 10. Hessling R. Novel aspects of the nervous system of Bonellia viridis (Echiura) revealed by the combination of immunohistochemistry, confocal laser-scanning microscopy and three- dimensional reconstruction. Hydrobiologia 2003; 496:225-39. 11. Hessling R, Westheide W. Are Echiura derived from a segmented ancestor? Immunohistochemical analysis of the nervous system in developmental stages of Bonellia viridis. J Morphol 2002; 252:100-13. 12. Wanninger A, Haszprunar G. Chiton myogenesis: Perspectives for the development and evolution of larval and adult muscle systems in molluscs. J Morphol 2002; 251:103-13. 13. McHugh D. Molecular evidence that echiurans and pogonophorans are derived annelids. Proc Natl Acad Sci USA 1997; 94:8006-9. 14. Boore JL, Staton JL. The mitochondrial genome of the sipunculid Phascolopsis gouldii sup- ports its association with Annelida rather than Mollusca. Mol Biol Evol 2002; 19:127-37. 15. Halanych KM, Dahlgren TG, McHugh D. Unsegmented annelids? Possible origins of four lophotrochozoan worm taxa. Int Comp Biol 2002; 42:678-84. 16. Struck TH, Schult N, Kusen T, Hickman E, Bleidorn C, McHugh D, Halanych KM. Annelid phylogeny and the status of Sipuncula and Echiura. BMC Evol Biol 2007; 7:57. 17. Dunn CW, Hejnol A, Matus DQ, Pang K, Browne WE, Smith SA, et al. Broad ­phylogenomic sampling improves resolution of the animal tree of life. Nature 2008; 452:745-9. 18. Kristof A, Wollesen T, Wanninger A. Segmental mode of neural patterning in Sipuncula. Curr Biol 2008; 18:1129-32. 19. Jaeckle WB, Rice ME. In: Atlas of Marine Invertebrate Larvae. Young CM, ed. San Diego: Academic Press 2002; 375-96. 20. Wanninger A, Koop D, Bromham L, Noonan E, Degnan BM. Nervous and muscle system development in Phascolion strombus (Sipuncula). Dev Genes Evol 2005; 215:509-18. 21. Iwanoff PP. Die Entwicklung der Larvalsegmente bei den Anneliden. Z Morph Ökol Tiere 1928; 10:161-62. 22. Brinkmann N, Wanninger A. Larval neurogenesis in Sabellaria alveolata reveals plasticity in polychaete neural patterning. Evol Dev 2008; 10:606-18. 23. Anderson DT. Embryology and phylogeny in annelids and arthropods. Oxford: Pergamon Press 1973. 24. Seaver EC, Thamm K, Hill SD. Growth patterns during segmentation in the two poly- chaete annelids, Capitella sp. I and Hydroides elegans: comparisons at distinct life history stages. Evol Dev 2005; 7:312-26. 25. Blair SS. Segmentation in animals. Curr Biol 2008; 18:991-5. 26. De Robertis EM. The molecular ancestry of segmentation mechanisms. Proc Natl Acad Sci USA 2008; 105:16411-2.

www.landesbioscience.com Communicative & Integrative Biology 59 57 Chapter IV

Chapter IV

Kristof A , Wollesen T, Maiorova AS & Wanninger A. 2011. Cellular and muscular growth patterns during sipunculan development. Journal of Experimental Zoology Part B: Molecular and Developmental Evolution 316B: 227-240

RESEARCH ARTICLE Cellular and Muscular Growth Patterns During Sipunculan Development ALEN KRISTOF1, TIM WOLLESEN1, ANASTASSYA S. MAIOROVA2, 1Ã AND ANDREAS WANNINGER 1Department of Biology, Research Group for Comparative Zoology, University of Copenhagen, Copenhagen, Denmark 2A. V. Zhirmunsky Institute of Marine Biology, Vladivostok, Russia

ABSTRACT Sipuncula is a lophotrochozoan taxon with annelid affinities, albeit lacking segmentation of the adult body. Here, we present data on cell proliferation and myogenesis during development of three sipunculan species, Phascolosoma agassizii, Thysanocardia nigra,andThemiste pyroides.The first anlagen of the circular body wall muscles appear simultaneously and not subsequently as in the annelids. At the same time, the rudiments of four longitudinal retractor muscles appear. This supports the notion that four introvert retractors were part of the ancestral sipunculan bodyplan. The longitudinal muscle fibers form a pattern of densely arranged fibers around the retractor muscles, indicating that the latter evolved from modified longitudinal body wall muscles. For a short time interval, the distribution of S-phase mitotic cells shows a metameric pattern in the developing ventral nerve cord during the pelagosphera stage. This pattern disappears close to metamorphic competence. Our findings are congruent with data on sipunculan neurogenesis, as well as with recent molecular analyses that place Sipuncula within Annelida, and thus strongly support a segmental ancestry of Sipuncula. J. Exp. Zool. (Mol. Dev. Evol.) 314B, 2011. & 2011 Wiley-Liss, Inc. How to cite this article: Kristof A, Wollesen T, Maiorova AS, Wanninger A. 2011. Cellular and J. Exp. Zool. (Mol. Dev. Evol.) muscular growth patterns during sipunculan development. J. Exp. Zool. (Mol. Dev. Evol.) 314B, 2011 314B:[page range].

The phylogenetic position and evolutionary origin of the as well as in their collagenous cuticle (Tzetlin and Purschke, sipunculans, a small and exclusively marine group of coelomate, 2006). This notion is further supported by recent developmental vermiform animals that show no obvious segmental organization studies on sipunculans and echiurans that have revealed in the adult stage, has been controversial for decades. They have segmental traits during neurogenesis (Hessling, 2002, 2003; been related to taxa as diverse as holothurians, echiurids, Hessling and Westheide, 2002; Kristof et al., 2008; Wanninger priapulids, phoronids, mollusks, or annelids (e.g., A˚ kesson, ’58; et al., 2009). Given their proposed inclusion within Annelida is Hyman, ’59; Rice ’85; Scheltema, ’93; Cutler, ’94). Recently, a correct, it seems plausible to assume secondary loss of a once number of independent molecular phylogenetic analyses have suggested a close relationship to Annelida (including Echiura) or Grant Sponsor: European Research Council; Grant number: MEST-CT-2005- even a nested position within this phylum (Boore and Staton, 020542; Grant Sponsors: Faculty of Science, University of Copenhagen; 2002; Staton, 2003; Jennings and Halanych, 2005; Bleidorn FEBRAS; Grant numbers: 10-III-B-06-089; 09-III-A-06-190; Grant Sponsor: et al., 2006; Struck et al., 2007; Dunn et al., 2008; Hejnol et al., RFFI; Grant numbers: 08-04-01001-a; 09-04-98584_r_vostok_a; 10-04- 10062_k. 2009; Mwinyi et al., 2009; Shen et al., 2009; Sperling et al., 2009; Ã Zrzavy et al., 2009). The latter scenario has received significant Correspondence to: Andreas Wanninger, Department of Biology, Research Group for Comparative Zoology, University of Copenhagen, Universitetsparken support by a recent study, whereby topology tests significantly 15, DK-2100 Copenhagen, Denmark. E-mail: [email protected] reject the sistergroup relationship of Sipuncula and Annelida Received 7 July 2010; Revised 4 October 2010; Accepted 1 December 2010 (Dordel et al., 2010). Moreover, ultrastructural similarities have Published online in Wiley Online Library (wileyonlinelibrary.com). been found in the foregut of certain sipunculans and polychaetes DOI: 10.1002/jez.b.21394

& 2011 WILEY-LISS, INC. 2 KRISTOF ET AL. segmented body plan rather than an initial evolutionary step (20–221C) until spawning occurred. In addition, fertilization toward segmentation in these animals. Interestingly, the lack of experiments were performed. Adult specimens were cut open and certain annelid key features, such as segmentation, coelomic gametes were transferred into glass jars. The eggs were fertilized cavities, nuchal organs, and chaetae, is also known for a variety with a few drops of a diluted sperm suspension. Embryos, larvae, of interstitial, parasitic, and sessile annelid representatives, but to and juveniles were reared in Petri dishes and glass vessels at a much lesser extent for large burrowing forms, such as 17–191C. Elongation of the anterior–posterior axis started at earthworms (reviewed in Bleidorn, 2007). 3 dpf in both species. Metamorphosis occurred at 10 dpf in Segmentation is usually considered a concerted repetition of T. nigra and at 15 dpf in T. pyroides. Development was followed organs or organ systems that form subsequently from a posterior in both species until the first juvenile stages, which already growth zone along the anterior–posterior axis of an animal showed the anlagen of the primary tentacles (i.e., at 18 dpf in (Willmer, ’90). Studies on the cell proliferation patterns and the T. pyroides and 15 dpf in T. nigra). ontogeny of organ systems typically associated with annelid segments (e.g., subsequently emerging sets of paired perikarya EdU Labeling, F-Actin Staining, Confocal Laserscanning Microscopy, associated with the ventral nerve cords, body wall ring muscles, and 3D Reconstruction nephridia) have proven to be ideal markers to assess whether or Proliferating cells were visualized by in vivo labeling with the not a taxon has derived from a segmented ancestor (Mu¨ller and nucleotide analogue 5-ethynyl-20-deoxyuridine (EdU) that is Westheide, 2000; Hessling, 2002, 2003; Hessling and Westheide, incorporated during the synthesis phase (S-phase) of the cell cycle. 2002; de Rosa et al., 2005; Seaver et al., 2005; Bergter et al., Larvae were incubated in EdU (Invitrogen, Taastrup, Denmark), 2007; Brinkmann and Wanninger, 2008; Kristof et al., 2008; diluted in filtered seawater in the following concentrations and Wanninger, 2009). Furthermore, the growing number of time intervals at 17–191C: 250 mMfor1hr,8mMfor6hr,and5mM immunocytochemical studies on neuro- and myogenesis of a for 24 hr. After EdU treatment and before F-actin staining, larvae number of lophotrochozoan taxa allows for a comparison of were anesthetized by adding drops of a 3.5 or 7% MgCl2 solution to nervous and muscle system patterning pathways in putatively the seawater and were subsequently fixed in 4% paraformaldehyde segmented and nonsegmented clades (e.g., Hay-Schmidt, 2000; in 0.1 M phosphate-buffered saline (PBS) (pH 7.3) for 1.5 hr at room Croll and Dickinson, 2004; McDougall et al., 2006; Bergter et al., temperature or overnight at 41C. This procedure was followed by 2007; Wanninger, 2008; Wanninger et al., 2008, 2009). Although three washes (15 min each) in 0.1 M PBS (pH 7.3) with 0.1% sodium some variation in annelid segment formation has been reported azide (NaN3). Until further processing, samples were stored in 0.1 M

(Seaver et al., 2005; Brinkmann and Wanninger, 2010), the PBSwith0.1%NaN3 at 41C. segmentation process driven from a posterior growth zone is After storage, larvae were rinsed in 0.1 M PBS (pH 7.3) for considered to be the ancestral condition for Annelida (Anderson, more than 6 hr, followed by incubation in a blocking and ’66; de Rosa et al., 2005; Seaver et al., 2005; Wanninger et al., permeabilization solution (saponin-based permeabilization and 2009). Herein, we compare the tempo–spatial distribution of wash reagent with 1% BSA) overnight at 41C. Incorporated EdU proliferating cells in Thysanocardia nigra and Themiste pyroides was detected with a Click-iT EdU Kit (Cat] C3005, Invitrogen, with growth patterns reported for annelids. We supplement this Taastrup, Denmark). The larvae were incubated with the reaction work with data on myogenesis in Phascolosoma agassizii, cocktail provided by the supplier (7,5 mL Alexa Flour 488 azide,

T. nigra,andT. pyroides, and thus provide insights into the 30 mLCuSO4,1313mL EdU reaction buffer, and 150 mL EdU buffer evolution of the myogenic bodyplans within the Lophotrochozoa. additive) for 24 hr at 41C. Some specimens were additionally incubated for 24 hr at 41C in a polyclonal rabbit anti-serotonin primary antibody (Calbiochem, Cambridge; dilution 1:200). The MATERIAL AND METHODS specimens were rinsed three times for 15 min in PBS. This was Animals followed by incubation with a goat anti-rabbit Alexa 594 Adult P. agassizii were collected in the vicinity of the Friday secondary antibody (dilution 1:300; Invitrogen, Taastrup, Denmark) Harbor Laboratories (Washington) and were kept in the in 0.1 M PBS for 24 hr at 41C. Finally, the specimens were laboratory until gametes were released. After fertilization, the washed three times for 15 min each in the saponin-based wash developing larvae were maintained in natural seawater at reagent with 1% BSA, incubated with the nucleic acid stain ambient temperature (101C). Development was followed until 40, 6-diamidino-2-phenylindole (DAPI; 1:10 dilution; Invitrogen, 15 days post-fertilization (dpf) when animals had reached the late Taastrup, Denmark), and mounted in Fluoromount G (Southern- pelagosphera stage. Adult specimens of T. nigra and T. pyroides Biotech, Birmingham, Alabama) on glass slides. were obtained from Crenomytilus grayanus mussel beds. Mussel For F-actin labeling, the stored larvae were washed three aggregations were collected by scuba divers at depths of 4–8 m times for 15 min in 0.1 M PBS, permeabilized for 6 hr in 0.1 M from the Vostok Bay, Sea of Japan (Russia). Adults were placed in PBS containing 4% Triton X-100 at 41C, and incubated in a 1:40 small tanks (15–30 specimens each) with ambient seawater dilution of Alexa Flour 488 phalloidin (Invitrogen, Taastrup,

J. Exp. Zool. (Mol. Dev. Evol.) SIPUNCULAN GROWTH PATTERNS 3

Denmark) in 0.1 M PBS in the dark for 24 hr at room temperature. number of S-phase cells has increased in the posterior trunk Thereafter, the samples were washed three times for 15 min and region, and more units of proliferating cells appear in the mounted in Fluoromount G on glass slides. Some specimens were posterior part of the ventral nerve cord (Figs. 1E, 2G, H). Lateral to double labeled for DAPI and F-actin. the ventral nerve cord, bilateral patches of stained nuclei seem to Analysis and digital image acquisition was performed on a migrate into the developing digestive system (Figs. 1E, 2G and H). Leica DM IRBE microscope equipped with a Leica TCS SP2 As in the 1 hr incubated specimens, the 6 hr incubated pelago- confocal unit (Leica Microsystems, Wetzlar, Germany). Optical sphera larvae show the same pattern of intensity decrease from sections taken at intervals of 0.5–0.2 mm were digitally merged posterior to anterior (Figs. 1E, 2G and H). As the larvae grow, most into maximum projection images. Images were further processed proliferating cells occur in the developing digestive system, with Photoshop 9.0.2 (Adobe Systems, San Jose, CA) to adjust around the anus, and in the anterior part of the ventral trunk area contrast and brightness. 3D reconstructions were created from (Figs. 1F, 2F, G). We found no units of S-phase cells in the selected confocal stacks using isosurface algorithms of the posterior ventral trunk region (Figs. 1F, G, 2I, J). In metamorphic imaging software Imaris v. 4.1 (Bitplane, Zu¨rich, Switzerland). competent larvae (18 dpf), the majority of proliferating cells is Digital line drawings were created with Corel Draw 11.0 (Corel detected in the anterior trunk, the introvert, and the digestive Corporation, Ottawa, Ontario, Canada). system (Fig. 1H). Post-metamorphic stages show proliferating cells mostly in the introvert and anlagen (not shown). Overall, the distribution pattern of proliferating cells during ontogeny of RESULTS T. pyroides and T. nigra does not unambiguously reveal a Cell Proliferation During Larval Development of T. pyroides and T. nigra textbook-like posterior growth zone. However, a distinct tissue Dividing cells were labeled with the thymidine analogue EdU formation zone appears temporarily in the posterior ventral trunk during development of T. pyroides and T. nigra. Combination of region in the pelagosphera stage. EdU and DAPI labeling enabled identification of proliferating cells and their location in the specimens (Fig. 1). Subsequent Myogenesis in P. agassizii clones of S-phase cells that have incorporated EdU show In the early trochophore larva of P. agassizii (i.e., at 6 dpf), progressive loss of staining intensity in the respective daughter numerous circular body wall muscles appear simultaneously, cells, which allows following the direction of proliferation together with the anlagen of the longitudinal retractor muscles (Fig. 1B, D, E; cf. Chehrehasa et al., 2009). (Figs. 3A inset and 4A). Slightly later, the longitudinal retractors In the trochophore stages, mitotic cells are scattered through- can be distinguished as one pair of ventral, one pair of dorsal, out the ectoderm (not shown). At the beginning of elongation and one single unpaired terminal organ retractor muscle (‘‘teardrop stage’’; age: 3 dpf), proliferating cells are densely (Fig. 3B). In addition, the mouth opening is visible and situated arranged around the eyes, mouth, metatroch, ventral trunk region, anterior to the anlage of the buccal musculature (Fig. 3B). and along the lateral and dorsal (Figs. 1A, 2A and B). The ventral and dorsal retractor muscles and the terminal organ Moreover, the S-phase cells are symmetrically arranged in two retractor muscle are well developed in the late trochophore larva lateral stripes in the introvert (Figs. 1A and 2A). Slightly later before the onset of longitudinal growth at 7 dpf (Figs. 3C and 4B). (3.5 dpf), proliferating cells appear in the area of the developing At the same time, numerous circular muscles of the future buccal digestive system (Figs. 1B, 2C and D). The highest number of musculature emerge posteriorly to the mouth opening (Figs. 3C proliferating cells is found along the developing ventral nerve and 4B). In addition, the entire length of the larval trunk is cord (Figs. 1B, 2C and D), which at this stage has three pairs of covered by circular body wall muscles (Figs. 3C and 4B). Their serotonergic perikarya associated with the serotonergic nerve number remains constant during the early phases of elongation cords (Fig. 1C). Moreover, intensity loss toward the posterior pole (7–8 dpf) (Fig. 3D and E). By contrast, additional longitudinal of the ventral trunk region indicates an anterior to posterior body wall and introvert retractor muscle fibers start to form direction of cell proliferation (Figs. 1B, 2C and D). As in the 1 hr (Fig. 3D and E). At the same time, the esophageal ring, intestinal incubated pelagosphera larvae aged 8 dpf, few units of proliferat- ring, and longitudinal muscle fibers, together with the anal ring ing cells appear in the posterior tip of the ventral nerve cord. The muscles of the digestive system, are established (Figs. 3D, E and signal intensity of these cells decreases from posterior to anterior 4C). As the pelagosphera larva grows (9–12 dpf), new circular (Figs. 1D, 2E and F). In addition, few weakly stained mitotic cells body wall muscles are synchronously added along the entire appear in the developing digestive system and in the posterior anterior–posterior axis by fission from existing myocytes trunk region (Figs. 1D, 2E and F). As expected, 8 dpf old (Figs. 3F and 4D). Accordingly, the formation of new circular pelagosphera larvae that had been exposed to EdU for 6 hr show body wall muscle fibers does not occur in a directional a higher number of proliferating cells in the ventral nerve cord, anterior–posterior process. The ventral longitudinal retractor the developing digestive system, around the mouth, and in two muscles contribute to the muscles that encircle the mouth bilateral stripes in the introvert (Figs. 1E, 2G and H). Moreover, the opening and surround the musculature of the buccal organ from

J. Exp. Zool. (Mol. Dev. Evol.) 4 KRISTOF ET AL.

Figure 1. Confocal micrographs of cell nuclei (blue), proliferating cells (red), and the serotonergic nervous system (green) during ontogeny of Themiste pyroides. Incubation with EdU was 1 hr in A and D and 6 hr in B, E, F, G, and H. All aspects are ventral views except B, C and G, which are lateral right views. Anterior faces upwards and scale bars equal 50 mm in all aspects except for C, in which it equals 10 mm. The stippled arrow in D and E shows the posterior to anterior decrease of the staining intensity of proliferating cells. Age of larvae is given in days post- fertilization (dpf). (A) Larva at the beginning of elongation (3 dpf) showing densely arranged proliferating cells throughout the ectoderm, around the eyes (e), the mouth (asterisk), the metatroch (mt), the ventral trunk region, and along the lateral and dorsal epidermis. (B)Early pelagosphera larva (3.5 dpf) showing the majority of proliferating cells along the ventral trunk area, the mouth, and the developing digestive system (ds). Along the ventral nerve cord, the intensity of proliferating cells decreases from anterior to posterior; svnc marks the serotonergic portion of the ventral nerve cord (svnc). (C) Same specimen as in B showing metamerically arranged perikarya (double arrowheads) associated with the serotonergic ventral nerve cords. (D) EdU-stained (1 hr) pelagosphera larva (8 dpf) with numerous proliferating cells in the area of the ventral nerve cord. The highest density of S-phase cells is in the posterior tip of the ventral nerve cord (arrows). Two weakly stained S-phase cells appear in the posterior end of the trunk (arrowheads). (E) Same stage as in D, incubated with EdU for 6 hr, showing the highest density and strongest signal of proliferating cells (arrows) at the posterior end of the ventral nerve cord (vnc). Note the stained cells at the posterior end of the trunk (arrowheads), in the developing digestive system, and around the mouth. (F) Late pelagosphera larva (15 dpf) with numerous proliferating cells in the developing gut and the ventral nerve cord. Note that the posterior part of the ventral nerve cord shows only few S-phase cells, whereas no proliferation appears in the posterior trunk cells (arrowheads). (G) Late stage larva (17 dpf) showing a decrease in mitotic activity in the ventral nerve cord, including the posterior trunk area (arrowhead). The majority of the proliferating cells are found in the digestive system. (H) Metamorphic competent larva (18 dpf) showing proliferation in the digestive system, around the mouth, and in the anterior part of the ventral trunk area. Note that the cells in the posterior trunk area (arrowheads) are not mitotic.

J. Exp. Zool. (Mol. Dev. Evol.) SIPUNCULAN GROWTH PATTERNS 5

either side (Figs. 3F, 4C and D). Similarly, the dorsal pair of longitudinal retractor muscle bands passes the prominent esophageal ring muscle fibers, which are situated dorsally to the buccal organ (Figs. 3F, 4C and D). In addition, the two branches of the terminal organ retractor muscle, which insert at the body wall dorsal to the anal ring muscles, can be distinguished (Figs. 3F, 4C and D). Close to metamorphosis (15 dpf), the myoanatomy of P. agassizii is composed of four solid longitudinal introvert retractors, which persist into adult- hood, a prominent longitudinal terminal organ retractor muscle with two distinct roots, and a strongly developed buccal and esophageal musculature (Figs. 4E and 5). In addition, the dorsal retractors intersect in the anterior introvert region (Figs. 4E and 5B). The body wall consists of numerous longitudinal muscle fibers which are loosely arranged in the mid-body region and in bands near the retractors, together with homogeneously arranged circular body wall muscles along the entire anterior–posterior axis (Figs. 4F, 5A, C and D). Moreover, new helicoid muscles, which probably connect the gut to the body wall, appear close to the insertion areas of the longitudinal retractor muscles

Figure 2. (Continued ) illustrating that the majority of the proliferating cells is found in the area of the ventral nerve cord, the mouth, and the metatroch. (C) Early pelagosphera larva (3.5 dpf), showing numerous S-phase cells in the developing digestive system, around the mouth, the introvert, and the ventral nerve cord. Note that the staining intensity in the ventral nerve cord decreases in anterior to posterior direction, and that the proliferating cells are clustered in units. (D) Lateral right view of the same specimen as in C. (E) Pelagosphera larva (8 dpf) showing the majority of the proliferating cells in the posterior portion of the ventral nerve cord arranged in several units (arrows). The intensity of the proliferating cells along the ventral nerve cord decreases from posterior to anterior (stippled arrow). Note the few weakly stained cells in the introvert and the posterior trunk area (arrowheads). (F) Lateral right view of the same specimen as in E. (G) Same stage as in D, with an increase of cell proliferation after 6 hr of incubation with EdU compared with the 1 hr incubated Figure 2. Schematic representation of cell proliferation patterns specimens. The number of units as well as proliferating cells per during Themiste pyroides development. Anterior faces upwards and unit has increased (arrows) and their intensity decreases from scale bars represent 50 mm in all aspects. Relative staining intensity posterior to anterior (stippled arrow). The number of proliferating is indicated from high (h) to low (l). Ventral views in A, C, E, G, and I cells has also increased in the posterior trunk area (arrowheads), and lateral right views in B, D, F, H, and J. Incubation with EdU was the introvert, and the developing digestive system. Note the cluster 1hrinA,B,E,andFand6hrinC,D,G,H,I,andJ.Ageoflarvaeis of S-phase cells in the anterior tip of the ventral nerve cord. given in days post-fertilization (dpf). (A) Tear-drop stage larva (H) Lateral right view of the same specimen as in G. (I)Latestage (3 dpf) with proliferating cells around the mouth (asterisk), along larva (17 dpf) showing most mitotic cells in the digestive system the metatroch (mt), the ventral nerve cord (vnc), the trunk and some proliferation in the ventral nerve cord, around the mouth, epidermis, and the introvert. Note the bilateral band of proliferating and the anus (a). Note that the proliferating cells in the ventral cells in the introvert, laterally to the eyes (e). at marks the apical nerve cord are not arranged in units and that no mitotic cells are tuft, pt the prototroch, and ds the anlage of the digestive found in the posterior trunk area. (J) Lateral right view of the same system. (B) Lateral right view of the same specimen as in A, specimen as in I.

J. Exp. Zool. (Mol. Dev. Evol.) 6 KRISTOF ET AL.

Figure 3. Confocal micrographs of myogenesis from the trochophore (A–C) to the pelagosphera stage (D–E)ofPhascolosoma agassizii. Anterior faces upwards and scale bars represent 30 mm in all aspects except for the inset in F, in which it equals 10 mm. A and C are in ventral view, whereas B, D, E, and F are in lateral left view. Age of larvae is given in days post-fertilization (dpf). (A) Early trochophore larva (6 dpf) showing the synchronous appearance of the first circular body wall muscles (cm: see inset) and the anlage of the ventral (vrm) and dorsal (drm) longitudinal retractor muscles as well as the terminal organ retractor muscle (trm). (B) Trochophore (6 dpf) with well established retractor muscles and the anlage of the buccal musculature (bm); asterisk marks the mouth opening. (C) Late trochophore (7 dpf) with well developed buccal musculature. Note that the entire trunk region is covered by circular body wall muscles. (D) Early pelagosphera larva (7 dpf) at the beginning of anterior–posterior elongation, showing the same number of circular muscles as in C, as well as the anal ring muscles (am). Note the establishment of additional longitudinal retractor muscle fibers (arrowheads). lm marks the longitudinal body wall muscles. (E) Larva (8 dpf), slightly more elongated than in D, but still retaining the number of circular muscles. The musculature of the (esm) and the intestine (inm) has been established. (F) Elongated pelagosphera larva (12 dpf) with new ring muscles being added. Strongly developed ventral and dorsal longitudinal retractor muscles, a terminal organ retractor muscle with two roots close to the anal ring muscles, and numerous well developed longitudinal body wall muscles (arrow) are present. The inset is a magnification of the boxed area in F. Note that new circular muscles are formed by fission (open triangles) from already existing myocytes (double arrow).

J. Exp. Zool. (Mol. Dev. Evol.) SIPUNCULAN GROWTH PATTERNS 7

Figure 4. Schematic representation of myogenesis in Phascolosoma agassizii. Anterior faces upwards and the total size of the specimen is approximately 150 mminAandB,190mminC,270mminD,and480mm in E and F. a-p indicates the anterior–posterior axis of the animals. Location of the apical tuft and the prototroch is indicated (purple). Eyes are omitted for clarity. Age of larvae is given in days post-fertilization (dpf). (A) Ventral view of a trochophore larva (6 dpf) with the anlagen of the paired ventral (yellow) and dorsal (turquoise) longitudinal retractor muscles, terminal organ retractor muscle (dark blue), buccal musculature (bm), and synchronously developing early anlagen of the circular body wall musculature (red). (B) Ventral view of a slightly later stage (6 dpf) with well developed buccal musculature and retractor muscles as well as circular body wall muscles along the entire trunk. Note the anlage of the digestive tract with a straight-through hindgut (hg). (C) Lateral left view of a longitudinal section through an early pelagosphera larva (8 dpf), showing the well developed U-shaped digestive tract with the dorsal anus (light green) and the retracted terminal organ (to). Note that the buccal musculature and the mouth opening (green) are enclosed by the ventral retractor muscles. The esophagus with its strong musculature (esm) lies between the dorsal retractor muscles, whereas the upper part of the intestine (in) lies between the two arms of the terminal organ retractor muscle. (D) Lateral left view of a pelagosphera larva (12 dpf) with newly formed circular muscle fibers. Note that new circular muscle fibers are formed along the entire anterior–posterior axis by fission from existing myocytes (double arrows). (E) Lateral left view of a metamorphic competent larva (15 dpf), showing strong and solid longitudinal retractor muscles. During anterior–posterior elongation of the body, the intestine establishes its typical U-shape with newly formed gut-fixing muscles (double arrowheads). Circular muscles omitted for clarity. (F)Samespecimenas in E highlighting the two-layered body wall musculature with outer circular and inner longitudinal muscle fibers (black). Note that the longitudinal body wall muscle fibers are densely arranged close to the laterally positioned retractor muscles and loosely toward the midbody region.

J. Exp. Zool. (Mol. Dev. Evol.) 8 KRISTOF ET AL.

Figure 5. Confocal micrographs of the myoanatomy of a metamorphic competent Phascolosoma agassizii larva aged 15 days post-fertilization (dpf). All aspects are in lateral left view except for B which is a dorsal view. Anterior faces upwards and scale bars represent 30 mminAand10mm in B, C, and D. (A) Larva with the prominent paired ventral (vrm), dorsal (drm), and the unpaired terminal organ retractor muscle (trm), longitudinal (lm) and circular body wall (cm) as well as buccal musculature (bm), helicoid gut-fixing muscles (double arrowheads), and the attachment sites of the longitudinal body wall muscles (asterisks). Left and right boxed areas are enlarged in C and D, respectively. (B) Dorsal view of the partly retracted introvert of the larva showing the intersection of the dorsal retractor muscles (open arrowheads) above the buccal musculature. (C)Samespecimen as in A, with focal depth being restricted to the trunk region, showing the attachment site of a ventral retractor muscle and the longitudinal body wall muscle fibers (arrows), as well as the attachment sites of the retractor muscle fibers at the body wall (asterisks). (D)SamespecimenasinA, with focal depth being restricted to the dorsal trunk region, showing the helicoid gut-fixing muscles (double arrowheads), which underlie the dorsal retractor muscle fibers, numerous longitudinal body wall muscle fibers (arrows), and the upended anal opening. am marks the anal ring muscles. in the late pelagosphera larva (Figs. 4E, 5A and D). No oblique Myogenesis in T. pyroides and T. nigra muscles were found in the body wall during myogenesis of Because myogenesis in T. pyroides and T. nigra is strikingly P. agassizii. similar, a detailed description is provided for T. pyroides only.

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Development from fertilization to the crawling juvenile is 11 dpf segments are added sequentially from proliferating cells located at 17–191CinT. nigra and 18 dpf in T. pyroides. In contrast to in one posterior or two lateral growth zone(s), and these cells P. agassizii, T. pyroides and T. nigra have a lecithotrophic might be homologous to the clitellate teloblasts (Shankland and development. Interestingly, an earlier report mentions direct Seaver, 2000; Seaver et al., 2005). Interestingly, a distinct development in T. pyroides populations from the Pacific North- posterior region of increased cell proliferation comparable to an east (Rice, ’67), whereas indirect development is known for this annelid-like growth zone seems to be present in certain species from the Sea of Japan, i.e., the West Pacific (Adrianov echiurans, a derived polychaete taxon (Hessling, 2003). et al., 2008; Adrianov and Maiorova, 2010). In T. pyroides and T. nigra, proliferating cells were constantly In T. pyroides, the first muscles appear simultaneously in 2 dpf present in tissues of developing organ systems, such as the old larvae as numerous circular body wall muscles, together with digestive system, the ventral nerve cord, and the post-meta- the anlagen of two pairs of longitudinal retractor muscles (Fig. 6A morphic tentacles. First, S-phase cells were scattered throughout and F). In the tear-drop stage, where anterior–posterior axis the ectoderm, then they appeared in the endodermal tissue of the elongation begins (2.5 dpf), the two pairs of longitudinal retractor developing gut before they formed metameric clusters (units) muscles can be distinguished (Fig. 6B). In addition, the ventral along the posterior ventral nerve cord during the pelagosphera retractor muscles encircle the mouth opening and more circular stage. The intensity of these EdU-labeled cells decreased in their body wall muscles appear (Fig. 6B). Slightly later (at approxi- respective daughter cells in anterior direction, which also mately 3 dpf), new circular body wall muscles form by fission indicates the direction of proliferation (Chehrehasa et al., 2009). from existing myocytes along the entire anterior–posterior axis However, toward metamorphosis and in post-metamorphic (Fig. 6C and G). In addition, the longitudinal retractor muscles are stages, these posterior clusters of mitotic cells disappear. well developed and numerous longitudinal body wall muscles Interestingly, in regenerating segments of Platynereis dumer- can be distinguished as bands near the retractor muscles (Fig. 6C ilii juveniles, BrdU experiments showed many stained cells on the and G). During elongation of the anterior–posterior axis in the ventral side and around the ventral nerve cord. Subsequently, pelagosphera larva (3.5 dpf), the retractor muscles become denser these cells migrate laterally as the segments become older and new circular and longitudinal body wall muscles form (intensity loss in respective daughter cells; de Rosa et al., 2005). (Fig. 6D). In late pelagosphera larvae (8 dpf), the retractor muscles However, BrdU pulse (15 min incubation with BrdU) and chase become prominent and the body wall musculature consists of experiments (fixation after 24 hr and 48 hr, respectively) in homogeneously arranged circular muscles along the entire regenerating segments of P. dumerilii juveniles revealed a anterior–posterior axis, as well as numerous longitudinal muscles distinct band of proliferating cells between the pygidium and around the retractors and few in the mid-body region (Fig. 6E the newly formed segment (de Rosa et al., 2005). A recent study and H). The body wall musculature of early juveniles (18 dpf) on the polychaetes Capitella and Hydroides showed that larval comprises ring muscles and underlying longitudinal muscles, segments in these distantly related polychaetes are formed from a which are densely arranged around the strongly developed field of dividing cells from the ventrolateral region of the body, paired ventral and dorsal retractor muscles (not shown). As in and that only post-metamorphic segments arise from the P. agassizii, no anterior to posterior development of newly posterior growth zone (Seaver et al., 2005). Likewise, neither an formed circular body wall muscles and no oblique body wall obvious posterior band of proliferating cells were detected in the muscle fibers were found during myogenesis of either T. pyroides larval segments of metatrochophores nor in later larval stages of or T. nigra. Sabellaria alveolata (Brinkmann and Wanninger, 2010). These data illustrate the ontogenetic plasticity of segment formation in annelids on a cellular level, a fact that is also reflected in the DISCUSSION highly variable mode of establishment of the various parts of the Proliferation Zones and Segmentation segmental nervous and muscle system in some polychaete Traditional descriptions of annelid segment formation have annelids (Brinkmann and Wanninger, 2008, 2010). mainly dealt with clitellates (leeches and oligocheates). In these Our findings on T. pyroides and T. nigra show different groups, segments are formed in anterior–posterior progression distribution patterns of proliferating cells during development. from a posterior growth zone that contains both mesodermal Only after the metameric arrangement of the nervous system has teloblasts and ectoblasts (e.g., Weisblat et al., ’88; Shankland, ’91; been established, proliferation increases in the ventral posterior de Rosa et al., 2005; Seaver et al., 2005; Brinkmann and trunk area and seems to progress from posterior to anterior. Wanninger, 2010). Divisions of the mesoteloblasts result in Accordingly, the distribution of mitotic cells in the sipunculan mesodermal bands that extend anteriorly and then form pelagosphera larva—but not in the earlier stages—corroborates the segmentally iterated muscle units. In the ectoderm, a ring of data on neurogenesis, which follows a segmental pattern ectoteloblasts localized in the growth zone generates the (Wanninger et al., 2005, 2009; Kristof et al., 2008). Interestingly, segmental ectoderm (Anderson, ’66, ’73). In polychaetes, a similar situation has recently been described for the polychaetes

J. Exp. Zool. (Mol. Dev. Evol.) 10 KRISTOF ET AL.

Figure 6. Myogenesis from the early trochophore (A) to the pelagosphera stage (D-E)ofThemiste pyroides. Anterior faces upwards and scale bars represent 50 mm in all aspects. A-E are confocal micrographs whereas F-H are schematic reconstructions. Color code in F, G, and H is as follows: circular body wall muscles in red, paired ventral and dorsal retractor muscles in yellow and turquoise, respectively, and longitudinal body wall muscles in black. Ventral views in all aspects except E, which is a lateral right view. Age of larvae is given in days post-fertilization (dpf). (A) Early trochophore larva (2 dpf) showing the anlage of the paired ventral (vrm) and dorsal (drm) longitudinal retractor muscles as well as numerous circular body wall muscles (cm); asterisk marks the mouth opening. (B) Slightly later stage (2.5 dpf) at the onset of elongation of the anterior–posterior axis. Elaborated retractor muscles and synchronously developing circular body wall muscles are present. Note that the ventral retractor muscles encircle the mouth opening. (C) Early pelagosphera larva (approximately 3 dpf), showing body wall musculature with outer circular and inner longitudinal muscles (lm). Note that the entire trunk is covered by circular body wall muscles and that the retractor muscles become more prominent. (D) Pelagosphera larva (3.5 dpf) with strong retractor muscles as well as circular and longitudinal body wall muscles. Note that longitudinal body wall muscles are more numerous near the retractors than in the mid-body region. (E) Late pelagosphera larva (8 dpf) with trunk covered by homogeneously arranged outer circular and inner longitudinal body wall muscles that are arranged in bands near the well established retractor muscles. (F) Schematic reconstruction of A. Note the early anlage of the circular body wall muscles and the paired ventral and dorsal retractor muscles. at marks the apical tuft, cb the ciliary bands, e the eye, eg the egg envelope, and m the mouth opening. (G)Schematic reconstruction of C. Note that new circular muscle fibers are formed along the entire anterior–posterior axis by fission from existing myocytes (double arrows), and that longitudinal body wall muscles are numerous close to the retractor muscles. pt marks the prototroch. (H)Schematic reconstruction of (E). Note the strong and solid retractor muscles of the pelagosphera larva.

J. Exp. Zool. (Mol. Dev. Evol.) SIPUNCULAN GROWTH PATTERNS 11

Hydroides and Capitella, whereby the posterior growth zone was developed circular muscles (Tzetlin and Filippova, 2005; established only in later larval stages after the first segments had Purschke and Mu¨ller, 2006). This may be correlated with the already been formed (Seaver et al., 2005). lifestyle of many polychaete taxa that use their parapodia for In T. pyroides and T. nigra, this metameric pattern of walking and swimming, thus probably positively selecting for a ectodermal S-phase cells is lost toward metamorphosis, as is pronounced longitudinal musculature rather than a prominent the case for the segmental arrangement of the nervous system in ring musculature. However, irrespective of their developmental P. agassizii (Kristof et al., 2008) and T. pyroides (unpublished modes, one ventral and one dorsal pair of longitudinal muscles data). Future studies including expression patterns of genes form the first rudiments of the longitudinal musculature in known to be involved in the establishment of segmentation in polychaetes and oligochaetes, whereas additional muscles, if annelids and arthropods, such as caudal, engrailed, brachyury, present, arise considerably later (McDougall et al., 2006; Bergter and even-skipped, should shed further light on the evolution of et al., 2007, 2008; Brinkmann and Wanninger, 2010). In contrast segmentation loss in Sipuncula. to circular body wall muscles, which are absent in the majority of polychaetes, it seems likely that these two pairs of primary Comparative Aspects of Myogenesis in Sipunculans and Annelids longitudinal body wall muscles are part of the annelid muscular The anlagen of the longitudinal retractor muscles and a groundpattern. considerable number of circular muscles appear simultaneously Several recent phylogenetic analyses suggest that the last in the early trochophore larvae of P. agassizii, T. nigra,and common annelid ancestor had a polychaete-like morphology T. pyroides. Slightly later, the entire trunk is covered by circular with parapodia and an aquatic, epibenthic lifestyle (Rousset et al., muscle fibers. Their number remains constant and increases only 2007; Struck et al., 2007; Zrzavy et al., 2009). Because body wall during the later larval stages. Thereby, newly formed circular ring muscles are common in most spiralians, it must be assumed muscle fibers of the body wall appear to be formed by fission that these are part of the spiralian groundplan. Accordingly, two from already existing myocytes along the entire length of the scenarios seem possible: (1) circular body wall muscles were lost body axis. In Phascolion strombi, which has a shortened larval at the base of Annelida (including the echiurans and sipunculans) phase and an entirely lecithotrophic mode of development, a and independently regained in some polychaetes, oligochaetes, different pattern of muscle formation is found. Here, new circular echiurans, and sipunculans or (2) circular body wall muscles were body wall muscle fibers are added between the already existing present in the last common ancestor of Annelida and lost several circular muscles along the entire anterior–posterior axis, whereby times independently within several subtaxa of the phylum. splitting of existing myocytes was not observed (Wanninger Interestingly, a study by Bergter et al. (2007) showed that in the et al., 2005). Accordingly, the process of simultaneous ring hirudinean Erpobdella octoculata the circular muscles do not muscle differentiation along the entire anterior–posterior axis show a segmental appearance from anterior to posterior, but are seems to be independent of a planktrotrophic or a lecithotrophic formed simultaneously along the anterior–posterior body axis, as life style, and thus probably ancestral to sipunculans (Wanninger, is the case in the sipunculans. Because Hirudinea (and maybe also 2009). This is in striking contrast to the situation found in the Sipuncula; see Struck et al., 2007; Dordel et al., 2010) is segmented annelids, where ring muscles, if present, are typically considered a derived annelid taxon that does not show coelomic formed in an anterior–posterior progression, probably owing to segmentation in the adults, the nonsegmental mode of myogen- the existence of the posterior growth zone (e.g., Hill and Boyer, esis in these two clades is most likely a secondary condition that 2001; Seaver et al., 2005; Bergter et al., 2007; Hunnekuhl et al., evolved independently in these taxa (Erseus and Ka¨llersjo¨,2004; 2009; Wanninger, 2009). Erseus, 2005; Jamieson and Ferraguti, 2006; Siddall et al., 2006; In all sipunculans investigated so far, the longitudinal body Bergter et al., 2007). wall musculature develops later than the circular muscles, and P. agassizii, T. nigra,andT. pyroides exhibit two pairs of because metamorphosis occurs without major dramatic gross introvert retractor muscles, thus corroborating earlier and recent morphological changes, these larval muscles form the scaffold of data on sipunculan pelagosphera larvae and early juveniles the adult musculature (Jaeckle and Rice, 2002; Wanninger et al., (A˚ kesson, ’58; Hall and Scheltema, ’75; Wanninger et al., 2005; 2005; Schulze and Rice, 2009). A body wall comprising an outer Schulze and Rice, 2009). These findings support the view of a layer of circular and an inner layer of longitudinal muscle fibers hypothetical ancestral sipunculan with four separate introvert as expressed in a number of oligochaetes is often considered as retractor muscles (Cutler and Gibbs, ’85; Cutler, ’94; Schulze and basal for Annelida (Dales, ’63; Lanzavecchia et al., ’88; Gardiner, Rice, 2009). In adult sipunculans, however, the number and ’92), although ring muscles are absent in various polychaete arrangement of the introvert retractors varies between four (e.g., clades (Purschke and Mu¨ller, 2006). In contrast to the uniform Phascolosoma) and one (e.g., Phascolion), and is therefore muscle layers found in clitellates, polychaetes exhibit a high considered of taxonomic relevance (Gibbs, ’77; Cutler, ’94). diversity of complex muscle patterns of distinct longitudinal The number of longitudinal body wall fibers, which underlie muscle bands and sometimes absent, incomplete, or only weakly the circular body wall muscles, increases markedly during the

J. Exp. Zool. (Mol. Dev. Evol.) 12 KRISTOF ET AL. pelagosphera stages of P. agassizii, T. nigra,andT. pyroides,and AUTHORS’ CERTIFICATION these muscles form a pattern of densely arranged fibers in the The authors declare that they agreed to be listed as such, and that area of the retractor muscles while they are looser toward the they have read and approved the final version of the manuscript. mid-body region. Because all sipunculans investigated so far exhibit this pattern of longitudinal body wall muscles, it seems likely that the introvert retractor muscles evolved from fused LITERATURE CITED longitudinal body wall muscles. The only sipunculans with Adrianov AV, Maiorova AS. 2010. Reproduction and development of intermediate oblique body wall muscle fibers are considerably common species of peanut worms (Sipuncula) from the Sea of large representatives of the Sipunculidae (Cutler, ’94). However, Japan. Russ J Mar Biol 36:1–15. such muscles have neither been found in larvae or juveniles of Adrianov AV, Maiorova AS, Malakhov VV. 2008. Embryonic and larval P. strombi (Wanninger et al., 2005) nor in late-stage larvae of development of the peanut worm Themiste pyroides (Sipuncula: P. agassizii or juveniles of T. nigra and T. pyroides (this study). Sipunculoidea) from the Sea of Japan. Invert Reprod Dev 52:143–151. This may be owing to secondary loss of such muscles, because a A˚kesson B. 1958. A study of the nervous system of the body wall musculature containing multilayered grids, including Sipunculoideae, with some remarks on the development of the oblique muscle fibers, is present in numerous vermiform soft- two species Phascolion strombi Montagu and Golfingia minuta bodied lophotrochozoans, such as annelids, platyhelminthes, Keferstein. Underso¨kningar Over O¨ resund 38:1–249. nemertines, or basal mollusks (Solenogastres, Caudofoveata, Anderson DT. 1966. The comparative embryology of the Polychaeta. larval Polyplacophora), and was thus likely a feature of the Acta Zool 47:1–42. spiralian stem species (Westheide and Rieger, ’96; Haszprunar Anderson DT. 1973. Embryology and phylogeny in annelids and and Wanninger, 2000; Ladurner and Rieger, 2000; Wanninger arthropods. Oxford: Pergamon Press. and Haszprunar, 2002; Filippova et al., 2005; Sørensen, 2005). Bergter A, Hunnekuhl VS, Schniederjans M, Paululat A. 2007. A similar arrangement in the muscular bodyplan of Acoela, Evolutionary aspects of patterning formation during clitellate supposedly the most basal bilaterian offshoot, increases the muscle development. Evol Dev 9:602–617. probability that such a multilayered body wall musculature was Bergter A, Brubacher JL, Paululat A. 2008. Muscle formation during also present in the last common bilaterian ancestor (Hooge and embryogenesis of the polychaete Ophryothrocha diadema (Dorvil- Tyler, 2006; Semmler et al., 2008). leidae)—new insights into annelid muscle patterns. Front Zool 5:1. Bleidorn C. 2007. The role of character loss in phylogenetic CONCLUSIONS reconstruction as exemplified for the Annelida. J Zool Syst Evol Development of Phascolosoma expresses a mosaic of segmental Res 45:285–387. and nonsegmental features. Although the ontogeny of the serially Bleidorn C, Podsiadlowski L, Bartolomaeus T. 2006. The complete repeated circular body wall muscles does not follow an mitochondrial genome of the orbiniid polychaete Orbinia latreillii anterior–posterior pattern, neurogenesis and the distribution of (Annelida, Orbiniidae)—a novel gene order for Annelida and proliferating cells exhibit transitional stages that resemble a implications for annelid phylogeny. Gene 370:96–103. segmental (i.e., annelid-like) mode of formation. This strongly Boore JL, Staton JL. 2002. The mitochondrial genome of the sipunculid supports a sipunculan–annelid clade, as suggested by recent Phascolopsis gouldii supports its association with Annelida rather molecular phylogenetic analyses, and argues in favour of a than with Mollusca. Mol Biol Evol 19:127–137. segmented sipunculan stem species. The obvious loss of Brinkmann N, Wanninger A. 2008. Larval neurogenesis in Sabellaria segmentation in adult sipunculans indicates that segmented alveolata reveals plasticity in polychaete neural patterning. Evol bodyplans may be more prone to evolutionary loss than Dev 10:606–618. previously thought and raises the question as to what extent this Brinkmann N, Wanninger A. 2010. Integrative analysis of polychaete may have also occurred in other ‘‘nonsegmented’’ metazoan taxa. ontogeny: cell proliferation patterns and myogenesis in trocho- phore larvae of Sabellaria alveolata. Evol Dev 12:5–15. ACKNOWLEDGMENTS Chehrehasa F, Meedeniya ACB, Dwyer P, Abrahamsen G, Mackay-Sim A. The authors acknowledge the support, hospitality, and help of 2009. EdU, a new thymidine analogue for labelling proliferating cells Andrey V. Adrianov and the staff of the Vostok Marine Biological in the nervous system. J Neurosci Meth 177:122–130. Station of the A. V. Zhirmunsky Institute of Marine Biology Croll RP, Dickinson AJG. 2004. Form and function of the larval nervous (Vladivostok, Russia). AK has been a recipient of an EU fellowship system in molluscs. Inv Reprod Dev 46:173–187. within the MOLMORPH network under the 6th Framework Cutler EB. 1994. The Sipuncula. Their systematics, biology and Program ‘‘Marie Curie Host Fellowships for Early Stage Research evolution. Ithaca, New York: Cornell University Press. Training,’’ which is coordinated by AW. AK and TW are funded Cutler EB, Gibbs PE. 1985. 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