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Alen Kristof | Københavns Universitet

BIOLOGISK INSTITUT 

THE MOLECULAR AND DEVELOPMENTAL BASIS OF BODYPLAN PATTERNING IN SIPUNCULA AND THE EVOLUTION OF SEGMENTATION 

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D E P A R T M E N T O F B I O L O G Y F A C U L T Y O F S C I E N C E U N I V E R S I T Y O F C O P E N H A G E N

PhD thesis Alen Kristof

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

Principal supervisor

Associate Prof. Dr. Andreas Wanninger

Co-supervisor

Prof. Dr. Pedro Martinez, University of Barcelona

April, 2011

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Reviewed by: Assistant Professor Anja Schulze Department of Marine Biology, 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 Themiste pyroides with a total length of 13 cm. Back: Confocal laserscanning micrograph of a Phascolosoma agassizii pelagosphera larva showing its musculature. Lateral view. Age of the specimen is 15 days and its total size approximately 300 µm in length.

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“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 Zoology, 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 species 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 Animal Models in Evolution

and Development at the University of Gothenburg, Sweden, Comparative Embryology

of Marine Invertebrates 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 life 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 animals.

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 family, 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 annelid 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 Themiste pyroides......................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 Trochozoa..................................................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 annelids......................................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 Peanut 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 order 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 trochophore larva. In addition,

throughout development newly formed ring muscles emerge along the entire anterior-

posterior axis by fission 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 metamorphosis 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 arthropods 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,

Ecdysozoa, and Lophotrochozoa) 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 Bilateria

(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 (polychaete), 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

China 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, Sipunculidae and Phascolosomatidea, comprising four orders and six families

(Aspidosiphonidae, Phascolosomatidae, Golfingiidae, Phascolionidae, 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 class Phascolosomatidae was recovered in the by

far most comprehensive analysis on sipunculan phylogeny of Schulze et al. (2005).

Congruently, all analyses strongly support Sipunculus nudus 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 cleavage, 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 Spiralia, 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 plankton 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 sea cucumbers (holothurian echinoderms), as derived annelids, or as

an in-group of Gephrea (sipunculans, echiurans, and priapulids), then as relatives of

phoronids, bryozoans, and brachiopods (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 Mollusca (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. Tentacles, 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

(Themistidae)

Serotonin,

FMRFamide

(chapter I)

F-actin

(chapter IV)

Dapi, EdU

(chapter IV)

Thysanocardia

nigra (Golfingidae)

Serotonin,

FMRFamide

(chapter I)

F-actin

(chapter IV)

Dapi, EdU

(chapter IV)

Phascolosoma

agassizii

(Phascolosomatidae)

Serotonin,

FMRFamide

(chapters I-III)

F-actin

(chapter IV)

-

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

CT 78 5’-GGAATTCATICKRTTYTGRAACCAIATTY

42°C

Engrailed (Seaver et al. 2006, Nederbragt et al. 2002)

EnEH2 5’-TGGCCTGCITGGGTNTAYTGYAC

en2-2 5’-TGRTTRTANARNCCYTGNGCCAT

42 °C

Eng1 5’-ATGGAATTCCNGCNTGGGTNTWYAC

Eng4 5’-TGGAAGCTTRTANARNCCYTSNGSCAT

42 °C

Myosin heavy chain type II (Ruiz-Trillo et al. 2002)

Mio7 (F) 5’-TGYATCAAYTWYACYAAYGAG

Mi6 (R) 5’-CCYTCMARYACACCRTTRCA

53 °C

Tropomyosin (Paps et al. 2009)

TropoF 5’-ATYRAGAAGAARATGNBKGVCATG

TropoR 5’-GTHYGRTCCARTTGNYCACT

53 °C

Intermediate filaments (Paps et al. 2009)

FilF1 5’-TACATCGAGAAGGTGCGTTTCCTGG

FilR1 5’-CCTCACCCTCCAGCAGCTTTCTGTA

48 °C

FilF3 5’-TACATCGAGAAGGTGCGTTTCCTGG

FilR3 5’-CYTCNCCYTCCAGCAGYTTYCTGTA

50 °C

β-Actin (Matsuo and Shimizu 2006)

19 Material and methods

β-Actin F1 5’-TGGGAYGAYATGGARAARAT

β-Actin R1 5’-GCCATYTCYTGYTCRAA

50 °C

α-Tubulin (Zheng et a. 1998)

AT16-F 5’-ATHGGHAAYGCNTGYTGGG

AT412-R 5’-RAAYTCDCCYTCYTCCATDCC

50 °C

β-Tubulin (Garant and MacRea 2009)

BT-F 5’-

GGNCARTCNGGNGCNGGNAAYAAYTGGGCN

BT-R 5’-NGGRAANGGNACCATRTTNACNGC

55 °C

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

(lab)

CGGATCCCTGGAACTGGAGAAAGAATTCCACTTTAACAAATATCTCACAAGAGCCAGGCGGATAGAAATAGCTGCTGCACTTGGACTAAATGAAACACACAGGTTAAGATCTGGTTTCAGAACAGCCGCATGAATTCC

136/10-10

hox3 GGATCCTTGGAACTGGAGAAGGAATTCCATTTTAACCGTTACTTGTGTCGGCCACGCCGTATTGAAATGGCAGCCATGCTCAACCTGACAGAAAGACAGATCAAGATCTGGTTTCAGAATAGCAGCATGAATTCC

135/10-14

hox5

(scr)

GGATCCCTGGAACTGGAGAAAGAATTTCATTACAACAGATACCTAACAAGACGAAGGAGAATAGAAATAGCACATGCTTTGGGACTCACGGAACGCCAAATTAAGATCTGGTTTCAGAATAGCAGCATGAATTCC

135/10-14

hox5

(ftz)

GGATCCTTGGAATTGGAAAAAGAATTTCACTTCAACCGTTACTTGTCCAGCAAACGACGTACAGAGATAGCCGAGTCACTGGCTCTGACAGATAGACAAGTTAAGATCTGGTTTCAAAACAGCCGCATGAATTCC

135/10-12

hox8

(abd-A)

CGGATCCTTGGAATTGGAAAAAGAATTTCAGTTTAATCATTATTTAACCCGAAAAAGGAGAATAGAGATAGCGCACGCTTTGTGTCTAACAGAAAGACAGATAAAGATCTGGTTTCAGAATCGCAGCATGAATTCC

136/10-15

even-

skipped

CGGATCCTTGGAGTTGGAAAAAGAATTTTACAGAGAGAACTATGTTAGCAGACCAAGGAGGTGTGAACTGGCTGCGGAACTCAACCTGCCAGAATCAACAATCAAGGTAAGAAGAAAATAGTGCGACCATTTATCACAGTTTCAGAGCCAACATTTTCAAAGTCGATTTTTCTAATTTGGAATAAAATTGCAATAAATTAATGTTTTATTTTTTACTTTCAGATCTGGTTTCAGAACAGCAGCATGAATTCC

252/10-9

lox2 CGGATCCTTGGAGCTGGAGAAAGAATTCAAATTCAACAGATACTTAACTCGCAGAAGACGAATAGAACTGTCTCATATGTTGTGCTTAACTGAGCGCCAAATCAAGATCTGGTTTCAGAACAGCAGCATGAATTCC

136/10-14

caudal

(cdx)

GGAATTCATGCTGCTGTTCTGAAACCAGATTTTGACCTGTCTCTCAGACAGACTGAGTGACTGTGCTAGTTCTGCCTTTCTTCTGATTGTGATATAACGACTGTAATGGAATTCTTTCTCCAGTTCCAGGGATCCG

136/10-12

xlox CGGATCCCCTGGAATTGGAAAAAGAATTTCATTTTAATAAATATATCTCCAGACCAAGGAGAATAGAATTAGCTGCCATGTTAAATCTAACAGAGAGACATATAAAGATCTGGTTTCAGAATAGCAGCATGAATTCC

137/10-14

not CGGATCCTTGGAGTTGGAGAAAGAATTTCACTTCAACCGTTACTTGTCCAGCAAACGACGTACAGAGATAG

271/10-11

21 Material and methods

CCGAGTCACTGGCTCTGACAGATAGACAAGTTAAGATCTGGTTTCAGAACAGCAGCATGAATTCCGGATCCCTGGAATTGGAGAAGGAATTCGAAAGGCAACAATACATGGTTGGATCTGAAAGGTATTATCTGGCAGTATCGCTTAATTTATCCGAATCCCAAGTGAAGATCTGGTTTCAGAACCGCCGCATGAATTCC

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 anus, 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 polychaetes 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 deuterostomes

(Nielsen and Hay-Schmidt 2007), although the expression of neural markers in this

neurite varies. Hence, this condition might be an adaptation of marine invertebrate

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 nephridium

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, arthropod and chordate 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 vertebrates 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

ReportSegmental Modeof Neural Patterning in Sipuncula

Alen Kristof,1 Tim Wollesen,1 and Andreas Wanninger1,*1Department of BiologyResearch Group for Comparative ZoologyUniversity of CopenhagenCopenhagen DK-2100Denmark

Summary

Recent molecular phylogenetic analyses suggest a close

relationship between two worm-shaped phyla, the nonseg-mented Sipuncula (peanut worms) and the segmented Anne-

lida (e.g., earthworms and polychaetes) [1–5]. The strikingdifferences in their bodyplans are exemplified by the anne-

lids’ paired, ladder-like ventral nervous system, which con-tains segmentally arranged ganglia, and the sipunculans’

single ventral nerve cord (VNC), which is devoid of any seg-mental structures [6, 7]. Investigating central nervous sys-

tem (CNS) formation with serotonin and FMRFamide labelingin a representative sipunculan, Phascolosoma agassizii, we

found that neurogenesis initially follows a segmental patternsimilar to that of annelids. Starting out with paired FMRFami-

dergic and serotonergic axons, four pairs of associatedserotonergic perikarya and interconnecting commissures

form one after another in an anterior-posterior progression.In late-stage larvae, the two serotonergic axons of the VNCs

fuse, the commissures disappear, and one additional pair of

perikarya is formed. These cells (ten in total) migrate towardone another, eventually forming two clusters of five cells

each. These neural-remodeling processes result in the sin-gle nonmetameric CNS of the adult sipunculan. Our data

confirm the segmental ancestry of Sipuncula and renderPhascolosoma a textbook example for the Haeckelian hy-

pothesis of ontogenetic recapitulation of the evolutionaryhistory of a species [8].

Results and Discussion

Ontogenetic Establishment of the Segmented

Sipunculan CNSDespite their proposed close phylogenetic relationship [1–5],annelids and sipunculans differ significantly from each otherin that most annelids exhibit a typical segmented body withmetamerically arranged appendages, commissures intercon-necting two ventral nerve cords (VNCs), and paired sets ofganglia, whereas sipunculans lack any trace of metamerismand have only a single VNC. Using antibodies against theneurotransmitters serotonin and FMRFamide, we found thatneurogenesis in Phascolosoma starts in the anterior regionwith the larval apical organ and the subsequently developingpaired serotonergic and FMRFamidergic axons of the VNC.Slightly later, one pair of serotonergic perikarya associatedwith the serotonergic portion of the VNCs appears(Figure 1A). As development and growth along the anterior-

posterior axis of the animal proceeds, a second pair of ventralperikarya arises (Figure 1B). In the following stages, the entireneural architecture of Phascolosoma is elaborated, resulting ina complex anterior nervous system that shares numerous fea-tures with annelid larvae. These include a (larval) serotonergicprototroch 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 andimportant, however, is the successive formation of two addi-tional pairs of serotonergic perikarya associated with the sero-tonergic axons of the VNCs, together with three serotonergicand FMRFamidergic commissures that interconnect therespective axons of the VNC (Figure 1B, inset, and Figures1C–1F). This results in a segmental central nervous system(CNS) comprising four metamerically arranged pairs of seroto-nergic perikarya along the VNCs of the late-stage Phascolo-soma larva (Figure 2A and Figure 3). Interestingly, only onepair of FMRFamidergic perikarya is found to be associatedwith the FMRFamidergic VNC (Figure 1F).

Annelid segmentation is not typified simply by the repetitivearrangement of organs along the longitudinal body axis but,more specifically, by their successive formation mode ina strict anterior-posterior progression; this indicates the pres-ence of a posterior growth zone from which the segmentallyarranged organs are budded off [9–13]. With the increasingmolecular evidence for an annelid-sipunculan assemblage[1–5], the question of whether the last common ancestor(LCA) of both taxa was segmented or not is still open becausewithin Lophotrochozoa (animals with a ciliated larval stage intheir life cycle), a segmented nervous system had been knownonly in annelids, including echiurans (spoon worms, a group ofmarine worms believed to nest within the annelid phylum) [1–5,12, 14, 15]. Our findings fill this gap in knowledge by providingthe hitherto-lacking proof of a segmented ancestry of the si-punculan nervous system [16], thus strongly arguing in favorof a segmented annelid-echiuran-sipunculan LCA.

Ontogenetic and Evolutionary Loss of the Segmental

Sipunculan CNSThe fully developed segmented nervous system of the larva ofPhascolosoma agassizii comprises four pairs of serotonergicperikarya and three serotonergic and FMRFamidergic com-missures interconnecting the VNCs (Figures 1D, IF, 2A, and3D). As the larva approaches metamorphic competence, thetwo serotonergic axons of the VNCs fuse, the serotonergiccommissures disappear, and a fifth pair of serotonergic peri-karya is formed (Figure 2B). Together with subsequent cellularmigratory processes, this eventually results in a CNS consist-ing of a single serotonergic VNC, which only retains its pairedcharacter in the anterior-most region where it connects to thebrain, as well as two neural cell clusters comprising five peri-karya each (Figures 2C and 3E). Accordingly, the segmentalserotonergic neural architecture is lost prior to the onset ofmetamorphosis of the Phascolosoma larva. At this stage, thethree FMRFamidergic commissures are still discernable, al-though the two FMRFamidergic axons of the VNCs lie inmuch closer proximity to each other than in the previous larvalstages (Figure 1F).*Correspondence: awanninger@bio.ku.dk

Initially paired ventral serotonergic nerves that fuse duringdevelopment have recently been described for another sipun-culan, Phascolion strombi. Although this species did not showunambiguous signs of segmentation during its ontogeny (afact that is probably due to its shortened larval phase), thetransitional occurrence of three FMRFamidergic commissuresin late P. strombi larvae indicates cryptic remnants of a once-segmented CNS in this species as well [17].

The findings of earlier morphological and developmentalstudies proposing a taxon comprising Sipuncula and Mollusca[18] have recently been rejected by a number of molecular anal-yses that clearly argue for an annelid-echiuran-sipunculanclade [1–5, 16]. Accordingly, novel data on CNS formation inthese groups cast new light on the evolution of annelid seg-mentation. Although the architecture of the adult annelid CNSseems to be quite plastic, ranging from one to five VNCs [19],neurogenesis consistently starts with an anterior neuropil (thepredecessor of the brain) and one pair of serotonergic andFMRFamidergic axons in the VNCs, irrespective of the numberof VNCs that are present in the adult stage [20–23]. This is con-gruent with the findings on echiuran and sipunculan neurogen-esis and suggests that a paired ventral CNS was also present inthe LCA of these three groups. However, various kinds of inde-pendent fusion and duplication events have taken place duringevolution, accounting for the phenotypic plasticity of the adultCNS that is encountered in these worm-shaped animals today.

In many annelids, ganglia and commissures are commonlyformed after the establishment of the first rudiments of thepaired VNC. Development of these commissures and gangliais strictly directional and follows an anterior-posterior pro-gression, a fact that is typically assigned to the presence ofa posterior growth zone [9, 10, 24, 25], and that eventuallyleads to the typical ladder-like metameric annelid ventralCNS. Although rather obvious in ‘‘typical’’ (e.g., polychaete)annelids, indications for the segmental ancestry of theechiurans are much more subtle and were only revealed re-cently by studies of their neural development [12, 14, 15].As such, echiurans exhibit a double-stranded VNC in earlylarval stages similar to the annelids [12]. Although thesenerve cords fuse during subsequent development, the asso-ciated perikarya form in anterior-posterior succession untilthe juvenile neural bodyplan is established [12, 14, 15].Thus, although modified, the segmental origin of the echiuranCNS can still be recognized in the adult stage by the meta-meric arrangement of the perikarya associated with theVNC. Moreover, cell-proliferation markers indicate an anne-lid-like posterior growth zone in Bonellia viridis, thus furtherstrengthening the evidence for the segmented origin ofechiurans [15]. In sipunculans, reduction of neural segmenta-tion traits has progressed even further than in echiurans,resulting in a CNS that is devoid of any metameric structuresin the adult stage (Figures 2C and 3E).

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 fourcells in theapicalor-

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 fusionof the twoserotonergicaxonsof theven-

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

the ventral nerve cord results in only one remaining

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-

mentof the adult cerebralganglionandexpressing

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.

Current Biology Vol 18 No 151130

To our knowledge, this work for the first time documentsontogenetic establishment and loss of CNS segmentation inthe life cycle of a recent metazoan and demonstrates

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).

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).

the segmental origin of Sipuncula. Ac-cordingly, Phascolosoma confirms theHaeckelian notion that cryptic characterstates, which have been lost during evo-lution in the adult bodyplan, may havebeen conserved during ontogeny [8],

thus stressing the importance of developmental studies forthe reconstruction of ancestral bodyplan features of metazoanclades.

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

Segmental Neurogenesis in Sipuncula1131

of the manuscript, as well as members of the Wanninger lab for discussion.

The comments of three anonymous reviewers helped to improve the manu-

script. A.K. is the recipient of an EU fellowship within the MOLMORPH net-

work under the 6th Framework Programme, ‘‘Marie Curie Host Fellowships

for Early Stage Research Training (EST)’’ (contract number MEST-CT-2005 –

020542). T.W. is funded by a PhD fellowship from the University of Copen-

hagen. The research of A.W. is supported by grants from the Danish Re-

search Agency (21-04-0356 and 271-05-0174) and the Carlsberg Foundation

(2005-1-249). Author contributions are as follows: A.K. performed research,

T.W. acquired the study material, A.K. and A.W. analyzed data, and A.W.

designed research and wrote the paper.

Received: April 23, 2008

Revised: May 29, 2008

Accepted: June 18, 2008

Published online: July 24, 2008

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Current Biology Vol 18 No 151132

Chapter III 52

Chapter III

Wanninger A, Kristof A & Brinkmann N. 2009. Sipunculans and

segmentation. Communicative & Integrative Biology 2: 56-59

Sipunculans and segmentation

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

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

Comparative molecular, developmental and morphogenetic analyses show that the three major segmented animal groups—Lophotrochozoa, Ecdysozoa and Vertebrata—use a wide range of ontogenetic pathways to establish metameric body organization. Even in the life history of a single specimen, different mecha-nisms may act on the level of gene expression, cell proliferation, tissue differentiation and organ system formation in individual segments. Accordingly, in some polychaete annelids the first three pairs of segmental peripheral neurons arise synchronously, while the metameric commissures of the ventral nervous system form in anterior-posterior progression. Contrary to traditional belief, loss of segmentation may have occurred more often than commonly assumed, as exemplified in the sipunculans, which show remnants of segmentation in larval stages but are unsegmented as adults. The developmental plasticity and potential evolutionary lability of segmentation nourishes the controversy of a segmented bilaterian ancestor versus multiple independent evolution of segmentation in respective metazoan lineages.

Ontogeny and Functional Implications of Segmentation

The evolution of a segmented bodyplan is often considered a crucial metazoan innovation because it allows the subdivision and specialization of individual body regions along the anterior-posterior axis of an animal.1,2 This partitioning typically involves both the ectodermal and the endodermal germ layers and often results in metameric ectodermal appendages (parapodia) and segmen-tally arranged, paired mesodermal body cavities (coelomic sacs). Traditionally, a condition where all segments have the same type of parapodia and house identical sets of internal organs such as ganglia, nephridia, gonads and muscles has been regarded as basal for annelids and arthropods (homonomic segmentation).1,2 From this basal (plesiomorphic) condition, concentration of individual organ systems into segments of specific body regions combined with reductions of organs in other segments are thought to have occurred multiple times within various lineages, and eventually led to morphologically distinct segments along the anterior-posterior body axis (heteronomic segmentation).1,2

While often considered an important hint towards ancestral segmentation of a species, serial repetition of organs along the ante-rior-posterior axis alone is not decisive for a segmental evolutionary history (cf., e.g., the multiple ring muscles in the non-segmented platyhelminths). On the cellular and organ system level, segmen-tation can only be proven with the aid of developmental studies, because segmented animals typically exhibit a posterior growth zone from which all segments are progressively budded off.3-7 Accordingly, ontogenetically older segments—and thus also the organs associated with them—are found anterior to the younger segments, a fact that is illustrated by the gradual decrease of the degree of organ system differentiation from anterior to posterior (Fig. 1).8 This makes the pattern of organogenesis an ideal marker to test for the segmental ancestry of worm-shaped lophotrochozoan taxa.8-12

Coelomic compartmentalization of a cylindrical body has frequently been proposed to be of selective advantage due to the fact that these animals are able to regulate the hemolymphic pressure in each compartment (segment) individually. The interplay of coelomic pressure and the contractile ring and longitudinal muscles of the body wall enable direct and independent control over the diameter of the body in each individual segment, thus allowing for a diversity of complex movement patterns.2 However, while coelomic segmen-tation has often (but not always) been retained in large, burrowing annelids (e.g., earthworms), secondary loss is often observed in non-benthic free-living (e.g., leeches), interstitial (e.g., Protodrilus), or sessile forms (e.g., tube worms).

Loss of Segmentation

Despite the loss of segmentation in various annelid taxa, ontoge-netic remnants of their segmented ancestry are present in a number of annelids that do not show obvious segmental features in the adult body (e.g., leeches).3,4 Recent developmental studies have shown that this holds also true for representatives of the Sipuncula (peanut worms), unsegmented lophotrochozoans that are regarded either as derived ingroup annelids or as a direct annelid sister clade.13-17 Interestingly, however, segmental traits in the sipunculans are restricted to the nervous system but have been completely lost on the level of coelom organisation.18 Accordingly, larvae of Phascolosoma agassizii develop four pairs of perikarya that are associated with the paired ventral nerve cord and express the common neurotransmitter serotonin (Fig. 2A). These paired perikarya form, together with commissures that interconnect the ventral nerve cords, in a typical annelid-like anterior-posterior progression, thus demonstrating the segmental ancestry of Sipuncula. During subsequent larval

Mini-Review

Sipunculans and segmentationAndreas 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

*Correspondence to: Andreas Wanninger; University of Copenhagen; Department of Biology; Research Group for Comparative Zoology; Universitetsparken 15; Copenhagen DK-2100 Denmark; Email: awanninger@bio.ku.dk

Submitted: 11/26/08; Accepted: 12/01/08

Previously published online as a Communicative & Integrative Biology E-publication: http://www.landesbioscience.com/journals/cib/article/7505

Sipunculans and segmentation

www.landesbioscience.com Communicative & Integrative Biology 57

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 nerve cord, this species lacks the associ-ated perikarya and only has retained three transitional ventral commissures as the sole remnants of the ancestral segmented neural bodyplan.20

Cryptic segmentation in taxa with close annelid affinities seems to be more common than previously assumed. The echiurans (spoon worms), now considered as clustering within Annelida,13,16,17 do not show any segmental traits in their adult gross morphology. However, neurogenesis revealed the same ontogenetic mechanisms as found in annelids and sipunculans, namely that paired sets of perikarya are formed in a discrete anterior-posterior progression.9-11 In contrast to the sipunculans and similar to the condition found in “typical” annelids, this metameric organization of the nervous system persists in the adult echiurans. Accordingly, the annelid-echi-uran-sipunculan lineage shows a gradual decrease of preservation of nervous system segmentation, a notion that is further supported by patterns of myogenesis. Hereby, annelids exhibit the typical anterior-posterior progression of ring and dorsoventral muscle formation, while sipunculan myogenesis starts with synchronous formation of early ring muscle rudiments, followed by the emer-gence of additional ring muscles along the entire anterior-posterior axis by fission from already existing myocytes.8,20 Accordingly, Sipuncula represents a developmental mosaic of segmental and non-segmental bodyplan patterning mechanisms, whereby the ectoderm-derived nervous system has to some degree maintained its segmental ancestry while the mesodermal musculature is formed entirely non-metamerically.

Plasticity of Segmentation

Despite the long standing definitions concerning the charac-teristics of a segmented bodyplan (see above), recent data have shown that the ontogenetic establishment of annelid segmentation may follow quite different developmental pathways. Traditionally, it had been proposed that the first three (larval) segments form more or less synchronously by schizocoely from the paired lateral mesodermal band, while the following (adult) segments develop from a pre-anal growth zone.21 Accordingly, one would assume that the organ systems associated with the first three segments also arise synchronously, while only the subsequent segmental organs follow the anterior-posterior differentiation gradient. However, this is only partly true for the polychaete Sabellaria alveolata. While the three larval segments indeed arise synchronously in this species, only the corresponding pairs of peripheral segmental neurons form synchro-nously, while the ventral commissures develop subsequently one after another (Fig. 1).22 While this may be interpreted as (secondary)

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 peripheral segmental neurons and the ventral commissures, respectively. Both aspects are ventral views with anterior facing upwards. Total length of the specimens is approximately 280 μm in (A) and 330 μm in (B). (A) Late larva with synchronously established peripheral segmental neurons (yellow). Ventral commissures and perikarya along the paired ventral nerve cord (vnc) are still lacking. The prototroch 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 of the adult neural bodyplan. (B) Larva prior to metamorphosis. The ventral commissures (asterisks) of the first five segments have been established progressively, together with the paired, metameric sets of perikarya (red dots) along the ventral nerve cords (vnc). The six pairs of peripheral segmental neurons (yellow) correspond to the segments II–VII, because development of segment I is retarded in this species, 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 segments VI and VII. pns – the nerves of the peripheral nervous system.

Sipunculans and segmentation

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

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, morphologically similar segments may follow different ontogenetic pathways even within the same individual, and distinct metamerically arranged subunits of the nervous system may form differently in individual segments of the same animal (e.g., the ventral commissures versus the peripheral segmental neurons in poly-chaetes; see above). Lastly, a mosaic of segmental and non-segmental modes of

organogenesis may occur within an individual, indicating the occur-rence of dissociation of organogenesis from the segmentation process in some species (e.g., muscle formation in sipunculans; see above).

Given the incongruencies and the plasticity of the processes involved in the ontogeny of segmentation in the various major bila-terian subgroups, no final statement as to whether or not Urbilateria was segmented can yet be made. In any case, assuming a segmented urbilaterian would imply a wide range of evolutionary modifications of the ancestral segmentation pathway on the molecular, cellular and morphogenetic level, as well as secondary loss of a segmented body in a number of lineages. Both, experimental developmental genetics employing RNAi experiments and comparative morphoge-netic analyses provide exciting tools that enable us to directly test for evolutionary hypotheses concerning shared molecular segmentation pathways on the one hand and for cryptic remnants of a segmented bodyplan in seemingly non-segmented recent phyla on the other. This should eventually lead to a sound reconstruction of the ancestry

chronological dissociation of larval segment formation and the development of the ventral commissures, it could alternatively be explained as the result of a heterochronic shift of the first three segments, which were originally derived from a posterior growth zone, into the larval stage of the animal. This notion is supported by reports that describe an—albeit rapid—progressive formation of these first three pairs of coelomic sacs in several polychaete taxa.23 Whatever alternative holds true, this example demonstrates that the developmental mechanisms that underlie annelid segmentation are much more complex than previously assumed. This is confirmed by recent cell proliferation pattern analyses, which suggest that the loca-tion of the growth zone might have shifted from a posterior-median position to both lateral sides in some species, indicating positional variability of the annelid growth zone.24 However, despite the high morphogenetic plasticity of segmentation, some molecular mecha-nisms appear similar even between distant phylogenetic entities such as arthropods and vertebrates.25,26

Figure 2. Expression and loss of the segmental pattern of the ventral CNS in the sipunculan Phascolosoma agassizii, as depicted by 3D reconstruction of serotonin immunoreactivity. Both aspects are ventral views with anterior facing upwards. Total length of the specimens is approximately 150 μm. (A) Late larva with four pairs of metameric perikarya (red; boxed area) associated with the ventral nerve cord (vnc). 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 mass (np) of the adult brain, the first cell bodies of the developing adult brain (light blue), two cells of the peripheral nervous system (orange), and the larval prototroch nerve ring (green). (B) Larva prior to metamorphosis in which the metameric arrangement of the ventral perikarya (red) has been lost in favor of two cell clusters (boxed areas) comprising five cells each. Note the increased number of cells belong-ing to the adult brain (light blue).

Sipunculans and segmentation

www.landesbioscience.com Communicative & Integrative Biology 59

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.

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

Cellular and Muscular GrowthPatterns During SipunculanDevelopmentALEN KRISTOF1, TIM WOLLESEN1, ANASTASSYA S. MAIOROVA2,AND ANDREAS WANNINGER1�1Department of Biology, Research Group for Comparative Zoology, University of Copenhagen,Copenhagen, Denmark

2A. V. Zhirmunsky Institute of Marine Biology, Vladivostok, Russia

Sipuncula is a lophotrochozoan taxon with annelid affinities, albeit lacking segmentation of theadult body. Here, we present data on cell proliferation and myogenesis during development ofthree sipunculan species, Phascolosoma agassizii, Thysanocardia nigra, and Themiste pyroides. Thefirst anlagen of the circular body wall muscles appear simultaneously and not subsequently as inthe annelids. At the same time, the rudiments of four longitudinal retractor muscles appear. Thissupports 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 retractormuscles, indicating that the latter evolved from modified longitudinal body wall muscles. For ashort time interval, the distribution of S-phase mitotic cells shows a metameric pattern in thedeveloping ventral nerve cord during the pelagosphera stage. This pattern disappears close tometamorphic competence. Our findings are congruent with data on sipunculan neurogenesis, aswell as with recent molecular analyses that place Sipuncula within Annelida, and thus stronglysupport a segmental ancestry of Sipuncula. J. Exp. Zool. (Mol. Dev. Evol.) 314B, 2011. & 2011Wiley-Liss, Inc.

How to cite this article: Kristof A, Wollesen T, Maiorova AS, Wanninger A. 2011. Cellular andmuscular growth patterns during sipunculan development. J. Exp. Zool. (Mol. Dev. Evol.)314B:[page range].

The phylogenetic position and evolutionary origin of the

sipunculans, a small and exclusively marine group of coelomate,

vermiform animals that show no obvious segmental organization

in the adult stage, has been controversial for decades. They have

been related to taxa as diverse as holothurians, echiurids,

priapulids, phoronids, mollusks, or annelids (e.g., Akesson, ’58;

Hyman, ’59; Rice ’85; Scheltema, ’93; Cutler, ’94). Recently, a

number of independent molecular phylogenetic analyses have

suggested a close relationship to Annelida (including Echiura) or

even a nested position within this phylum (Boore and Staton,

2002; Staton, 2003; Jennings and Halanych, 2005; Bleidorn

et al., 2006; Struck et al., 2007; Dunn et al., 2008; Hejnol et al.,

2009; Mwinyi et al., 2009; Shen et al., 2009; Sperling et al., 2009;

Zrzavy et al., 2009). The latter scenario has received significant

support by a recent study, whereby topology tests significantly

reject the sistergroup relationship of Sipuncula and Annelida

(Dordel et al., 2010). Moreover, ultrastructural similarities have

been found in the foregut of certain sipunculans and polychaetes

as well as in their collagenous cuticle (Tzetlin and Purschke,

2006). This notion is further supported by recent developmental

studies on sipunculans and echiurans that have revealed

segmental traits during neurogenesis (Hessling, 2002, 2003;

Hessling and Westheide, 2002; Kristof et al., 2008; Wanninger

et al., 2009). Given their proposed inclusion within Annelida is

correct, it seems plausible to assume secondary loss of a once

Published online in Wiley Online Library (wileyonlinelibrary.com).

DOI: 10.1002/jez.b.21394

Received 7 July 2010; Revised 4 October 2010; Accepted 1 December 2010

Grant Sponsor: European Research Council; Grant number: MEST-CT-2005-

020542; Grant Sponsors: Faculty of Science, University of Copenhagen;

FEBRAS; Grant numbers: 10-III-B-06-089; 09-III-A-06-190; Grant Sponsor:

RFFI; Grant numbers: 08-04-01001-a; 09-04-98584_r_vostok_a; 10-04-

10062_k.�Correspondence to: Andreas Wanninger, Department of Biology, Research

Group for Comparative Zoology, University of Copenhagen, Universitetsparken

15, DK-2100 Copenhagen, Denmark. E-mail: awanninger@bio.ku.dk

ABSTRACT

J. Exp. Zool.(Mol. Dev. Evol.)314B, 2011

& 2011 WILEY-LISS, INC.

RESEARCH ARTICLE

segmented body plan rather than an initial evolutionary step

toward segmentation in these animals. Interestingly, the lack of

certain annelid key features, such as segmentation, coelomic

cavities, nuchal organs, and chaetae, is also known for a variety

of interstitial, parasitic, and sessile annelid representatives, but to

a much lesser extent for large burrowing forms, such as

earthworms (reviewed in Bleidorn, 2007).

Segmentation is usually considered a concerted repetition of

organs or organ systems that form subsequently from a posterior

growth zone along the anterior–posterior axis of an animal

(Willmer, ’90). Studies on the cell proliferation patterns and the

ontogeny of organ systems typically associated with annelid

segments (e.g., subsequently emerging sets of paired perikarya

associated with the ventral nerve cords, body wall ring muscles,

nephridia) have proven to be ideal markers to assess whether or

not a taxon has derived from a segmented ancestor (Muller and

Westheide, 2000; Hessling, 2002, 2003; Hessling and Westheide,

2002; de Rosa et al., 2005; Seaver et al., 2005; Bergter et al.,

2007; Brinkmann and Wanninger, 2008; Kristof et al., 2008;

Wanninger, 2009). Furthermore, the growing number of

immunocytochemical studies on neuro- and myogenesis of a

number of lophotrochozoan taxa allows for a comparison of

nervous and muscle system patterning pathways in putatively

segmented and nonsegmented clades (e.g., Hay-Schmidt, 2000;

Croll and Dickinson, 2004; McDougall et al., 2006; Bergter et al.,

2007; Wanninger, 2008; Wanninger et al., 2008, 2009). Although

some variation in annelid segment formation has been reported

(Seaver et al., 2005; Brinkmann and Wanninger, 2010), the

segmentation process driven from a posterior growth zone is

considered to be the ancestral condition for Annelida (Anderson,

’66; de Rosa et al., 2005; Seaver et al., 2005; Wanninger et al.,

2009). Herein, we compare the tempo–spatial distribution of

proliferating cells in Thysanocardia nigra and Themiste pyroides

with growth patterns reported for annelids. We supplement this

work with data on myogenesis in Phascolosoma agassizii,

T. nigra, and T. pyroides, and thus provide insights into the

evolution of the myogenic bodyplans within the Lophotrochozoa.

MATERIAL AND METHODS

Animals

Adult P. agassizii were collected in the vicinity of the Friday

Harbor Laboratories (Washington) and were kept in the

laboratory until gametes were released. After fertilization, the

developing larvae were maintained in natural seawater at

ambient temperature (101C). Development was followed until

15 days post-fertilization (dpf) when animals had reached the late

pelagosphera stage. Adult specimens of T. nigra and T. pyroides

were obtained from Crenomytilus grayanus mussel beds. Mussel

aggregations were collected by scuba divers at depths of 4–8m

from the Vostok Bay, Sea of Japan (Russia). Adults were placed in

small tanks (15–30 specimens each) with ambient seawater

(20–221C) until spawning occurred. In addition, fertilization

experiments were performed. Adult specimens were cut open and

gametes were transferred into glass jars. The eggs were fertilized

with a few drops of a diluted sperm suspension. Embryos, larvae,

and juveniles were reared in Petri dishes and glass vessels at

17–191C. Elongation of the anterior–posterior axis started at

3 dpf in both species. Metamorphosis occurred at 10 dpf in

T. nigra and at 15 dpf in T. pyroides. Development was followed

in both species until the first juvenile stages, which already

showed the anlagen of the primary tentacles (i.e., at 18 dpf in

T. pyroides and 15dpf in T. nigra).

EdU Labeling, F-Actin Staining, Confocal Laserscanning Microscopy,and 3D Reconstruction

Proliferating cells were visualized by in vivo labeling with the

nucleotide analogue 5-ethynyl-20-deoxyuridine (EdU) that is

incorporated during the synthesis phase (S-phase) of the cell cycle.

Larvae were incubated in EdU (Invitrogen, Taastrup, Denmark),

diluted in filtered seawater in the following concentrations and

time intervals at 17–191C: 250mM for 1hr, 8mM for 6hr, and 5mM

for 24hr. After EdU treatment and before F-actin staining, larvae

were anesthetized by adding drops of a 3.5 or 7% MgCl2 solution to

the seawater and were subsequently fixed in 4% paraformaldehyde

in 0.1M phosphate-buffered saline (PBS) (pH 7.3) for 1.5 hr at room

temperature or overnight at 41C. This procedure was followed by

three washes (15min each) in 0.1M PBS (pH 7.3) with 0.1% sodium

azide (NaN3). Until further processing, samples were stored in 0.1M

PBS with 0.1% NaN3 at 41C.

After storage, larvae were rinsed in 0.1 M PBS (pH 7.3) for

more than 6 hr, followed by incubation in a blocking and

permeabilization solution (saponin-based permeabilization and

wash reagent with 1% BSA) overnight at 41C. Incorporated EdU

was detected with a Click-iT EdU Kit (Cat] C3005, Invitrogen,

Taastrup, Denmark). The larvae were incubated with the reaction

cocktail provided by the supplier (7,5mL Alexa Flour 488 azide,

30mL CuSO4, 1313mL EdU reaction buffer, and 150mL EdU buffer

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

specimens were rinsed three times for 15min in PBS. This was

followed by incubation with a goat anti-rabbit Alexa 594

secondary antibody (dilution 1:300; Invitrogen, Taastrup, Denmark)

in 0.1 M PBS for 24hr at 41C. Finally, the specimens were

washed three times for 15min each in the saponin-based wash

reagent with 1% BSA, incubated with the nucleic acid stain

40, 6-diamidino-2-phenylindole (DAPI; 1:10 dilution; Invitrogen,

Taastrup, Denmark), and mounted in Fluoromount G (Southern-

Biotech, Birmingham, Alabama) on glass slides.

For F-actin labeling, the stored larvae were washed three

times for 15min in 0.1 M PBS, permeabilized for 6 hr in 0.1 M

PBS containing 4% Triton X-100 at 41C, and incubated in a 1:40

dilution of Alexa Flour 488 phalloidin (Invitrogen, Taastrup,

KRISTOF ET AL.2

J. Exp. Zool. (Mol. Dev. Evol.)

Denmark) in 0.1 M PBS in the dark for 24hr at room temperature.

Thereafter, the samples were washed three times for 15min and

mounted in Fluoromount G on glass slides. Some specimens were

double labeled for DAPI and F-actin.

Analysis and digital image acquisition was performed on a

Leica DM IRBE microscope equipped with a Leica TCS SP2

confocal unit (Leica Microsystems, Wetzlar, Germany). Optical

sections taken at intervals of 0.5–0.2mm were digitally merged

into maximum projection images. Images were further processed

with Photoshop 9.0.2 (Adobe Systems, San Jose, CA) to adjust

contrast and brightness. 3D reconstructions were created from

selected confocal stacks using isosurface algorithms of the

imaging software Imaris v. 4.1 (Bitplane, Zurich, Switzerland).

Digital line drawings were created with Corel Draw 11.0 (Corel

Corporation, Ottawa, Ontario, Canada).

RESULTS

Cell Proliferation During Larval Development of T. pyroides and T. nigra

Dividing cells were labeled with the thymidine analogue EdU

during development of T. pyroides and T. nigra. Combination of

EdU and DAPI labeling enabled identification of proliferating

cells and their location in the specimens (Fig. 1). Subsequent

clones of S-phase cells that have incorporated EdU show

progressive loss of staining intensity in the respective daughter

cells, which allows following the direction of proliferation

(Fig. 1B, D, E; cf. Chehrehasa et al., 2009).

In the trochophore stages, mitotic cells are scattered through-

out the ectoderm (not shown). At the beginning of elongation

(‘‘teardrop stage’’; age: 3 dpf), proliferating cells are densely

arranged around the eyes, mouth, metatroch, ventral trunk region,

and along the lateral and dorsal epidermis (Figs. 1A, 2A and B).

Moreover, the S-phase cells are symmetrically arranged in two

lateral stripes in the introvert (Figs. 1A and 2A). Slightly later

(3.5 dpf), proliferating cells appear in the area of the developing

digestive system (Figs. 1B, 2C and D). The highest number of

proliferating cells is found along the developing ventral nerve

cord (Figs. 1B, 2C and D), which at this stage has three pairs of

serotonergic perikarya associated with the serotonergic nerve

cords (Fig. 1C). Moreover, intensity loss toward the posterior pole

of the ventral trunk region indicates an anterior to posterior

direction of cell proliferation (Figs. 1B, 2C and D). As in the 1hr

incubated pelagosphera larvae aged 8dpf, few units of proliferat-

ing cells appear in the posterior tip of the ventral nerve cord. The

signal intensity of these cells decreases from posterior to anterior

(Figs. 1D, 2E and F). In addition, few weakly stained mitotic cells

appear in the developing digestive system and in the posterior

trunk region (Figs. 1D, 2E and F). As expected, 8 dpf old

pelagosphera larvae that had been exposed to EdU for 6 hr show

a higher number of proliferating cells in the ventral nerve cord,

the developing digestive system, around the mouth, and in two

bilateral stripes in the introvert (Figs. 1E, 2G and H). Moreover, the

number of S-phase cells has increased in the posterior trunk

region, and more units of proliferating cells appear in the

posterior part of the ventral nerve cord (Figs. 1E, 2G, H). Lateral to

the ventral nerve cord, bilateral patches of stained nuclei seem to

migrate into the developing digestive system (Figs. 1E, 2G and H).

As in the 1hr incubated specimens, the 6 hr incubated pelago-

sphera larvae show the same pattern of intensity decrease from

posterior to anterior (Figs. 1E, 2G and H). As the larvae grow, most

proliferating cells occur in the developing digestive system,

around the anus, and in the anterior part of the ventral trunk area

(Figs. 1F, 2F, G). We found no units of S-phase cells in the

posterior ventral trunk region (Figs. 1F, G, 2I, J). In metamorphic

competent larvae (18 dpf), the majority of proliferating cells is

detected in the anterior trunk, the introvert, and the digestive

system (Fig. 1H). Post-metamorphic stages show proliferating cells

mostly in the introvert and tentacle anlagen (not shown). Overall,

the distribution pattern of proliferating cells during ontogeny of

T. pyroides and T. nigra does not unambiguously reveal a

textbook-like posterior growth zone. However, a distinct tissue

formation zone appears temporarily in the posterior ventral trunk

region in the pelagosphera stage.

Myogenesis in P. agassizii

In the early trochophore larva of P. agassizii (i.e., at 6 dpf),

numerous circular body wall muscles appear simultaneously,

together with the anlagen of the longitudinal retractor muscles

(Figs. 3A inset and 4A). Slightly later, the longitudinal retractors

can be distinguished as one pair of ventral, one pair of dorsal,

and one single unpaired terminal organ retractor muscle

(Fig. 3B). In addition, the mouth opening is visible and situated

anterior to the anlage of the buccal musculature (Fig. 3B).

The ventral and dorsal retractor muscles and the terminal organ

retractor muscle are well developed in the late trochophore larva

before the onset of longitudinal growth at 7 dpf (Figs. 3C and 4B).

At the same time, numerous circular muscles of the future buccal

musculature emerge posteriorly to the mouth opening (Figs. 3C

and 4B). In addition, the entire length of the larval trunk is

covered by circular body wall muscles (Figs. 3C and 4B). Their

number remains constant during the early phases of elongation

(7–8dpf) (Fig. 3D and E). By contrast, additional longitudinal

body wall and introvert retractor muscle fibers start to form

(Fig. 3D and E). At the same time, the esophageal ring, intestinal

ring, and longitudinal muscle fibers, together with the anal ring

muscles of the digestive system, are established (Figs. 3D, E and

4C). As the pelagosphera larva grows (9–12 dpf), new circular

body wall muscles are synchronously added along the entire

anterior–posterior axis by fission from existing myocytes

(Figs. 3F and 4D). Accordingly, the formation of new circular

body wall muscle fibers does not occur in a directional

anterior–posterior process. The ventral longitudinal retractor

muscles contribute to the muscles that encircle the mouth

opening and surround the musculature of the buccal organ from

SIPUNCULAN GROWTH PATTERNS 3

J. Exp. Zool. (Mol. Dev. Evol.)

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 50mm in all aspects except for C, in which it equals 10mm. 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.

KRISTOF ET AL.4

J. Exp. Zool. (Mol. Dev. Evol.)

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. Schematic representation of cell proliferation patterns

during Themiste pyroides development. Anterior faces upwards and

scale bars represent 50mm in all aspects. Relative staining intensity

is indicated from high (h) to low (l). Ventral views in A, C, E, G, and I

and lateral right views in B, D, F, H, and J. Incubation with EdU was

1 hr in A, B, E, and F and 6 hr in C, D, G, H, I, and J. Age of larvae is

given in days post-fertilization (dpf). (A) Tear-drop stage larva

(3 dpf) with proliferating cells around the mouth (asterisk), along

the metatroch (mt), the ventral nerve cord (vnc), the trunk

epidermis, and the introvert. Note the bilateral band of proliferating

cells in the introvert, laterally to the eyes (e). at marks the apical

tuft, pt the prototroch, and ds the anlage of the digestive

system. (B) Lateral right view of the same specimen as in A,

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

specimens. The number of units as well as proliferating cells per

unit has increased (arrows) and their intensity decreases from

posterior to anterior (stippled arrow). The number of proliferating

cells has also increased in the posterior trunk area (arrowheads),

the introvert, and the developing digestive system. Note the cluster

of S-phase cells in the anterior tip of the ventral nerve cord.

(H) Lateral right view of the same specimen as in G. (I) Late stage

larva (17 dpf) showing most mitotic cells in the digestive system

and some proliferation in the ventral nerve cord, around the mouth,

and the anus (a). Note that the proliferating cells in the ventral

nerve cord are not arranged in units and that no mitotic cells are

found in the posterior trunk area. (J) Lateral right view of the same

specimen as in I.

SIPUNCULAN GROWTH PATTERNS 5

J. Exp. Zool. (Mol. Dev. Evol.)

Figure 3. Confocal micrographs of myogenesis from the trochophore (A–C) to the pelagosphera stage (D–E) of Phascolosoma agassizii.

Anterior faces upwards and scale bars represent 30mm in all aspects except for the inset in F, in which it equals 10mm. 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 esophagus (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).

KRISTOF ET AL.6

J. Exp. Zool. (Mol. Dev. Evol.)

Figure 4. Schematic representation of myogenesis in Phascolosoma agassizii. Anterior faces upwards and the total size of the specimen is

approximately 150mm in A and B, 190mm in C, 270mm in D, and 480mm 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) Same specimen as

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.

SIPUNCULAN GROWTH PATTERNS 7

J. Exp. Zool. (Mol. Dev. Evol.)

in the late pelagosphera larva (Figs. 4E, 5A and D). No oblique

muscles were found in the body wall during myogenesis of

P. agassizii.

Myogenesis in T. pyroides and T. nigra

Because myogenesis in T. pyroides and T. nigra is strikingly

similar, a detailed description is provided for T. pyroides only.

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 30mm in A and 10 mm

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) Same specimen

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) Same specimen as in A,

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.

KRISTOF ET AL.8

J. Exp. Zool. (Mol. Dev. Evol.)

Development from fertilization to the crawling juvenile is 11dpf

at 17–191C in T. nigra and 18dpf in T. pyroides. In contrast to

P. agassizii, T. pyroides and T. nigra have a lecithotrophic

development. Interestingly, an earlier report mentions direct

development in T. pyroides populations from the Pacific North-

east (Rice, ’67), whereas indirect development is known for this

species from the Sea of Japan, i.e., the West Pacific (Adrianov

et al., 2008; Adrianov and Maiorova, 2010).

In T. pyroides, the first muscles appear simultaneously in 2 dpf

old larvae as numerous circular body wall muscles, together with

the anlagen of two pairs of longitudinal retractor muscles (Fig. 6A

and F). In the tear-drop stage, where anterior–posterior axis

elongation begins (2.5 dpf), the two pairs of longitudinal retractor

muscles can be distinguished (Fig. 6B). In addition, the ventral

retractor muscles encircle the mouth opening and more circular

body wall muscles appear (Fig. 6B). Slightly later (at approxi-

mately 3 dpf), new circular body wall muscles form by fission

from existing myocytes along the entire anterior–posterior axis

(Fig. 6C and G). In addition, the longitudinal retractor muscles are

well developed and numerous longitudinal body wall muscles

can be distinguished as bands near the retractor muscles (Fig. 6C

and G). During elongation of the anterior–posterior axis in the

pelagosphera larva (3.5 dpf), the retractor muscles become denser

and new circular and longitudinal body wall muscles form

(Fig. 6D). In late pelagosphera larvae (8 dpf), the retractor muscles

become prominent and the body wall musculature consists of

homogeneously arranged circular muscles along the entire

anterior–posterior axis, as well as numerous longitudinal muscles

around the retractors and few in the mid-body region (Fig. 6E

and H). The body wall musculature of early juveniles (18 dpf)

comprises ring muscles and underlying longitudinal muscles,

which are densely arranged around the strongly developed

paired ventral and dorsal retractor muscles (not shown). As in

P. agassizii, no anterior to posterior development of newly

formed circular body wall muscles and no oblique body wall

muscle fibers were found during myogenesis of either T. pyroides

or T. nigra.

DISCUSSION

Proliferation Zones and Segmentation

Traditional descriptions of annelid segment formation have

mainly dealt with clitellates (leeches and oligocheates). In these

groups, segments are formed in anterior–posterior progression

from a posterior growth zone that contains both mesodermal

teloblasts and ectoblasts (e.g., Weisblat et al., ’88; Shankland, ’91;

de Rosa et al., 2005; Seaver et al., 2005; Brinkmann and

Wanninger, 2010). Divisions of the mesoteloblasts result in

mesodermal bands that extend anteriorly and then form

segmentally iterated muscle units. In the ectoderm, a ring of

ectoteloblasts localized in the growth zone generates the

segmental ectoderm (Anderson, ’66, ’73). In polychaetes,

segments are added sequentially from proliferating cells located

in one posterior or two lateral growth zone(s), and these cells

might be homologous to the clitellate teloblasts (Shankland and

Seaver, 2000; Seaver et al., 2005). Interestingly, a distinct

posterior region of increased cell proliferation comparable to an

annelid-like growth zone seems to be present in certain

echiurans, a derived polychaete taxon (Hessling, 2003).

In T. pyroides and T. nigra, proliferating cells were constantly

present in tissues of developing organ systems, such as the

digestive system, the ventral nerve cord, and the post-meta-

morphic tentacles. First, S-phase cells were scattered throughout

the ectoderm, then they appeared in the endodermal tissue of the

developing gut before they formed metameric clusters (units)

along the posterior ventral nerve cord during the pelagosphera

stage. The intensity of these EdU-labeled cells decreased in their

respective daughter cells in anterior direction, which also

indicates the direction of proliferation (Chehrehasa et al., 2009).

However, toward metamorphosis and in post-metamorphic

stages, these posterior clusters of mitotic cells disappear.

Interestingly, in regenerating segments of Platynereis dumer-

ilii juveniles, BrdU experiments showed many stained cells on the

ventral side and around the ventral nerve cord. Subsequently,

these cells migrate laterally as the segments become older

(intensity loss in respective daughter cells; de Rosa et al., 2005).

However, BrdU pulse (15 min incubation with BrdU) and chase

experiments (fixation after 24hr and 48hr, respectively) in

regenerating segments of P. dumerilii juveniles revealed a

distinct band of proliferating cells between the pygidium and

the newly formed segment (de Rosa et al., 2005). A recent study

on the polychaetes Capitella and Hydroides showed that larval

segments in these distantly related polychaetes are formed from a

field of dividing cells from the ventrolateral region of the body,

and that only post-metamorphic segments arise from the

posterior growth zone (Seaver et al., 2005). Likewise, neither an

obvious posterior band of proliferating cells were detected in the

larval segments of metatrochophores nor in later larval stages of

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

highly variable mode of establishment of the various parts of the

segmental nervous and muscle system in some polychaete

annelids (Brinkmann and Wanninger, 2008, 2010).

Our findings on T. pyroides and T. nigra show different

distribution patterns of proliferating cells during development.

Only after the metameric arrangement of the nervous system has

been established, proliferation increases in the ventral posterior

trunk area and seems to progress from posterior to anterior.

Accordingly, the distribution of mitotic cells in the sipunculan

pelagosphera larva—but not in the earlier stages—corroborates the

data on neurogenesis, which follows a segmental pattern

(Wanninger et al., 2005, 2009; Kristof et al., 2008). Interestingly,

a similar situation has recently been described for the polychaetes

SIPUNCULAN GROWTH PATTERNS 9

J. Exp. Zool. (Mol. Dev. Evol.)

Figure 6. Myogenesis from the early trochophore (A) to the pelagosphera stage (D-E) of Themiste pyroides. Anterior faces upwards and

scale bars represent 50mm 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.

KRISTOF ET AL.10

J. Exp. Zool. (Mol. Dev. Evol.)

Hydroides and Capitella, whereby the posterior growth zone was

established only in later larval stages after the first segments had

already been formed (Seaver et al., 2005).

In T. pyroides and T. nigra, this metameric pattern of

ectodermal S-phase cells is lost toward metamorphosis, as is

the case for the segmental arrangement of the nervous system in

P. agassizii (Kristof et al., 2008) and T. pyroides (unpublished

data). Future studies including expression patterns of genes

known to be involved in the establishment of segmentation in

annelids and arthropods, such as caudal, engrailed, brachyury,

and even-skipped, should shed further light on the evolution of

segmentation loss in Sipuncula.

Comparative Aspects of Myogenesis in Sipunculans and Annelids

The anlagen of the longitudinal retractor muscles and a

considerable number of circular muscles appear simultaneously

in the early trochophore larvae of P. agassizii, T. nigra, and

T. pyroides. Slightly later, the entire trunk is covered by circular

muscle fibers. Their number remains constant and increases only

during the later larval stages. Thereby, newly formed circular

muscle fibers of the body wall appear to be formed by fission

from already existing myocytes along the entire length of the

body axis. In Phascolion strombi, which has a shortened larval

phase and an entirely lecithotrophic mode of development, a

different pattern of muscle formation is found. Here, new circular

body wall muscle fibers are added between the already existing

circular muscles along the entire anterior–posterior axis, whereby

splitting of existing myocytes was not observed (Wanninger

et al., 2005). Accordingly, the process of simultaneous ring

muscle differentiation along the entire anterior–posterior axis

seems to be independent of a planktrotrophic or a lecithotrophic

life style, and thus probably ancestral to sipunculans (Wanninger,

2009). This is in striking contrast to the situation found in the

segmented annelids, where ring muscles, if present, are typically

formed in an anterior–posterior progression, probably owing to

the existence of the posterior growth zone (e.g., Hill and Boyer,

2001; Seaver et al., 2005; Bergter et al., 2007; Hunnekuhl et al.,

2009; Wanninger, 2009).

In all sipunculans investigated so far, the longitudinal body

wall musculature develops later than the circular muscles, and

because metamorphosis occurs without major dramatic gross

morphological changes, these larval muscles form the scaffold of

the adult musculature (Jaeckle and Rice, 2002; Wanninger et al.,

2005; Schulze and Rice, 2009). A body wall comprising an outer

layer of circular and an inner layer of longitudinal muscle fibers

as expressed in a number of oligochaetes is often considered as

basal for Annelida (Dales, ’63; Lanzavecchia et al., ’88; Gardiner,

’92), although ring muscles are absent in various polychaete

clades (Purschke and Muller, 2006). In contrast to the uniform

muscle layers found in clitellates, polychaetes exhibit a high

diversity of complex muscle patterns of distinct longitudinal

muscle bands and sometimes absent, incomplete, or only weakly

developed circular muscles (Tzetlin and Filippova, 2005;

Purschke and Muller, 2006). This may be correlated with the

lifestyle of many polychaete taxa that use their parapodia for

walking and swimming, thus probably positively selecting for a

pronounced longitudinal musculature rather than a prominent

ring musculature. However, irrespective of their developmental

modes, one ventral and one dorsal pair of longitudinal muscles

form the first rudiments of the longitudinal musculature in

polychaetes and oligochaetes, whereas additional muscles, if

present, arise considerably later (McDougall et al., 2006; Bergter

et al., 2007, 2008; Brinkmann and Wanninger, 2010). In contrast

to circular body wall muscles, which are absent in the majority of

polychaetes, it seems likely that these two pairs of primary

longitudinal body wall muscles are part of the annelid muscular

groundpattern.

Several recent phylogenetic analyses suggest that the last

common annelid ancestor had a polychaete-like morphology

with parapodia and an aquatic, epibenthic lifestyle (Rousset et al.,

2007; Struck et al., 2007; Zrzavy et al., 2009). Because body wall

ring muscles are common in most spiralians, it must be assumed

that these are part of the spiralian groundplan. Accordingly, two

scenarios seem possible: (1) circular body wall muscles were lost

at the base of Annelida (including the echiurans and sipunculans)

and independently regained in some polychaetes, oligochaetes,

echiurans, and sipunculans or (2) circular body wall muscles were

present in the last common ancestor of Annelida and lost several

times independently within several subtaxa of the phylum.

Interestingly, a study by Bergter et al. (2007) showed that in the

hirudinean Erpobdella octoculata the circular muscles do not

show a segmental appearance from anterior to posterior, but are

formed simultaneously along the anterior–posterior body axis, as

is the case in the sipunculans. Because Hirudinea (and maybe also

Sipuncula; see Struck et al., 2007; Dordel et al., 2010) is

considered a derived annelid taxon that does not show coelomic

segmentation in the adults, the nonsegmental mode of myogen-

esis in these two clades is most likely a secondary condition that

evolved independently in these taxa (Erseus and Kallersjo, 2004;

Erseus, 2005; Jamieson and Ferraguti, 2006; Siddall et al., 2006;

Bergter et al., 2007).

P. agassizii, T. nigra, and T. pyroides exhibit two pairs of

introvert retractor muscles, thus corroborating earlier and recent

data on sipunculan pelagosphera larvae and early juveniles

(Akesson, ’58; Hall and Scheltema, ’75; Wanninger et al., 2005;

Schulze and Rice, 2009). These findings support the view of a

hypothetical ancestral sipunculan with four separate introvert

retractor muscles (Cutler and Gibbs, ’85; Cutler, ’94; Schulze and

Rice, 2009). In adult sipunculans, however, the number and

arrangement of the introvert retractors varies between four (e.g.,

Phascolosoma) and one (e.g., Phascolion), and is therefore

considered of taxonomic relevance (Gibbs, ’77; Cutler, ’94).

The number of longitudinal body wall fibers, which underlie

the circular body wall muscles, increases markedly during the

SIPUNCULAN GROWTH PATTERNS 11

J. Exp. Zool. (Mol. Dev. Evol.)

pelagosphera stages of P. agassizii, T. nigra, and T. pyroides, and

these muscles form a pattern of densely arranged fibers in the

area of the retractor muscles while they are looser toward the

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

longitudinal body wall muscles. The only sipunculans with

intermediate oblique body wall muscle fibers are considerably

large representatives of the Sipunculidae (Cutler, ’94). However,

such muscles have neither been found in larvae or juveniles of

P. strombi (Wanninger et al., 2005) nor in late-stage larvae of

P. agassizii or juveniles of T. nigra and T. pyroides (this study).

This may be owing to secondary loss of such muscles, because a

body wall musculature containing multilayered grids, including

oblique muscle fibers, is present in numerous vermiform soft-

bodied lophotrochozoans, such as annelids, platyhelminthes,

nemertines, or basal mollusks (Solenogastres, Caudofoveata,

larval Polyplacophora), and was thus likely a feature of the

spiralian stem species (Westheide and Rieger, ’96; Haszprunar

and Wanninger, 2000; Ladurner and Rieger, 2000; Wanninger

and Haszprunar, 2002; Filippova et al., 2005; Sørensen, 2005).

A similar arrangement in the muscular bodyplan of Acoela,

supposedly the most basal bilaterian offshoot, increases the

probability that such a multilayered body wall musculature was

also present in the last common bilaterian ancestor (Hooge and

Tyler, 2006; Semmler et al., 2008).

CONCLUSIONSDevelopment of Phascolosoma expresses a mosaic of segmental

and nonsegmental features. Although the ontogeny of the serially

repeated circular body wall muscles does not follow an

anterior–posterior pattern, neurogenesis and the distribution of

proliferating cells exhibit transitional stages that resemble a

segmental (i.e., annelid-like) mode of formation. This strongly

supports a sipunculan–annelid clade, as suggested by recent

molecular phylogenetic analyses, and argues in favour of a

segmented sipunculan stem species. The obvious loss of

segmentation in adult sipunculans indicates that segmented

bodyplans may be more prone to evolutionary loss than

previously thought and raises the question as to what extent this

may have also occurred in other ‘‘nonsegmented’’ metazoan taxa.

ACKNOWLEDGMENTSThe authors acknowledge the support, hospitality, and help of

Andrey V. Adrianov and the staff of the Vostok Marine Biological

Station of the A. V. Zhirmunsky Institute of Marine Biology

(Vladivostok, Russia). AK has been a recipient of an EU fellowship

within the MOLMORPH network under the 6th Framework

Program ‘‘Marie Curie Host Fellowships for Early Stage Research

Training,’’ which is coordinated by AW. AK and TW are funded

by a Ph.D. fellowship from the University of Copenhagen. ASM is

supported by FEBRAS and RFFI.

AUTHORS’ CERTIFICATIONThe authors declare that they agreed to be listed as such, and that

they have read and approved the final version of the manuscript.

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