Reproductive Biology and Phylogeny of Annelida

Reproductive Biology and Phylogeny of Annelida Volume edited by GREG ROUSE South Australian Museum Nth Terrace, Adelaide. and Earth and Environmental Sciences University of Adelaide S.A. 5000 Australia FREDRIK PLEIJEL Department of Marine Ecology Tjärnö Marine Biological Laboratory Göteborg University Strömstad, Sweden and Muséum national d’Histoire naturelle Département Systématique et Evolution Paris Cedex 05, France

Volume 4 of Series: Reproductive Biology and Phylogeny Series edited by BARRIE G.M. JAMIESON School of Integrative Biology University of Queensland St. Lucia, Queensland Australia

Science Publishers Enfield (NH)

Jersey

Plymouth

SCIENCE PUBLISHERS An Imprint of Edenbridge Ltd., British Isles. Post Office Box 699 Enfield, New Hampshire 03748 United States of America Website: http://www.scipub.net [email protected] (marketing department) [email protected] (editorial department) [email protected] (for all other enquiries) ISBN (Set) 1-57808-271-4 ISBN (Vol. 4) 1-57808-313-3 © 2006, Copyright reserved Library of Congress Cataloging-in-Publication Data Reproductive biology and phylogeny of Annelida/volume edited by Greg Rouse, Fredrik Pleijel. p. cm -- (Reproductive biology and phylogeny; v. 4) Includes bibliographical references (p. ). ISBN 1-57808-313-3 -- ISBN 1-57808-271-4 (set) 1. Annelida--Reproduction. I. Rouse, Greg W. II. Pleijel, Fredrik, III.Series. QL391.A6R44 2006 571.8’126--dc22 2005055989

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Preface to the Series This series was founded by the present series editor, Barrie Jamieson, in consultation with Science Publishers, Inc., in 2001. The series bears the title ‘Reproductive Biology and Phylogeny’ and this title is followed in each volume with the name of the taxonomic group which is the subject of the volume. Each publication has one or more invited volume editors and a large number of authors of international repute. The level of the taxonomic group which is the subject of each volume varies according, largely, to the amount of information available on the group, the advice of proposed volume editors, and the interest expressed by the zoological community in the proposed work. The order of publication of taxonomic groups reflects these concerns, and the availability of authors for the various chapters, and it is not proposed to proceed serially through the animal kingdom in a presumed “ladder of life” sequence. Nevertheless, a second aspect of the series is coverage of the phylogeny and classification of the group, as a necessary framework for an understanding of reproductive biology. Evidence for relationships from molecular studies is an important aspect of the chapter on phylogeny and classification. Other chapters may or may not have phylogenetic themes, according to the interests of the authors. It is not claimed that a single volume can, in fact, cover the entire gamut of reproductive topics for a given group but it is believed that the series gives an unsurpassed coverage of reproduction and provides a general text rather than being a mere collection of research papers on the subject. Coverage in different volumes will vary in terms of topics, though it is clear from the first volumes that the standard of the contributions by the authors will be uniformly high. The stress will vary from group to group; for instance, modes of external fertilization or vocalization, important in one group, might be inapplicable in another. The first three volumes on Urodela, edited by Professor David Sever, Anura, edited by myself, and Chondrichthyes, edited by Professor William Hamlett, reflected the above exacting criteria and the interests of certain research teams. This, the fourth volume, has resulted from our good fortune in the acceptance by Drs. Greg Rouse and Fredrik Pleijel of invitations to edit a volume on Annelida. Both editors are outstanding authorities on polychaetes. In contributions to this volume they have demonstrated, again,

LE Reproductive Biology and Phylogeny of Annelida the highest standards of scholarship and research. Their enthusiasm, despite heavy research commitments, for this pivotal group in studies on invertebrates, has been apparent in their rigorous editing and in the six highly authoritative chapters to which one or both have been the sole or collaborative authors. In their choice of topics for the volume they have drawn on a most distinguished group of authors who need no introduction to those familiar with annelid studies. Three further volumes in preparation are on Gymnophiona (Jean-Marie Exbrayat), Cetacea (Debra Miller) and Aves (Barrie Jamieson). While volume editing is by invitation, biologists who consider that a given taxonomic group should be included in the series and may wish to undertake the task of editing a volume should not hesitate to make their views known to the series editor who provides editorial support. My thanks are due to the School of Integrative Biology, University of Queensland, for facilities, and especially to the Executive Dean, Professor Mick McManus, for his encouragement. I am grateful to the publishers for their friendly support and high standards in producing this series. Sincere thanks must be given to the volume editors and the authors, who have freely contributed their chapters, in very full schedules. The editors and publishers are gratified that the enthusiasm and expertise of these contributors has been reflected by the reception of the series by our readers.

27th October, 2005

Barrie Jamieson The School of Integrative Biology University of Queensland Brisbane

Preface to this Volume Annelida is a diverse group of animals, commonly referred to as segmented worms and currently comprising around 14 000 described species. Found in most marine and freshwater areas, annelids have also successfully occupied many subterranean habitats. They vary greatly in form, and as adults range in length from a fraction of a millimeter to well over 6 meters. The tremendous variety of reproductive modes found among annelids may be a significant factor in this diversity and broad distribution. Terrestrial and freshwater annelids tend to show parental care of some form, ranging from brooding embryos in tubes to placing them in cocoons. There are also examples of increased parental investment such as feeding of young and even viviparity. Among marine annelids, these reproductive modes are also found, along with the simplest reproductive method of simply expelling sperm and eggs into the water column. There are many small clades of annelids that show a range of reproductive modes and this provides an excellent opportunity to study the factors in evolving features such as parental care (or the lack of it). To date this has yet to be exploited to any great degree. In a previous volume that dealt with polychaete annelids (Rouse and Pleijel 2001), we did not attempt a synthesis on reproduction. When offered the opportunity to do so in the series Reproductive Biology and Phylogeny edited by Barrie Jamieson we were very happy to accept. We are grateful indeed to Barrie for his patience and willingness to help throughout the course of this project. The scope of the project was such that we felt an edited book with a variety of experts was the best approach. The goal of this volume is to document annelid reproduction in the context of their phylogenetic relationships. We present an introduction and overview to the current systematics of annelids and then proceed with a section that provides reviews to broad aspects of reproduction across Annelida. These chapters cover oogenesis, sperm, mating, early development, larval development and larval ecology. Then follows a series of chapters that cover some of the major clades (or purported clades) of annelids and address similar issues but in more detail for the particular groups. The final chapter covers some of the more problematic annelid groups in terms of their phylogenetic placement. Unfortunately for various reasons we were not able to organise the inclusion of chapters covering

LEEE Reproductive Biology and Phylogeny of Annelida annelid endocrine systems and reproductive physiology or more specific chapters of important annelid groups such as Eunicida or Terebelliformia. Nevertheless we hope that this volume proves to be a useful resource for those interested in the fascinating diversity of reproductive modes in annelids. October 2005

Greg Rouse Adelaide, Australia Fredrik Pleijel Strömstad, Sweden and Paris, France

LITERATURE CITED Rouse, G. W. and Pleijel, F. 2001. Polychaetes, Oxford University Press, London. 354 pp.

Contents Preface to the Series – Barrie G. M. Jamieson Preface to this Volume – Greg W. Rouse and Fredrik Pleijel

v vii

I General Reproduction and Phylogeny 1. Annelid Phylogeny and Systematics Greg W. Rouse and Fredrik Pleijel

3

2. Oogenesis Kevin J. Eckelbarger

23

3. Annelid Sperm and Spermiogenesis Greg W. Rouse

45

4. Sexual Strategies and Mating Systems Gabriella Sella

77

5. Early Annelid Development, A Molecular Perspective Steven Q. Irvine and Elaine C. Seaver

93

6. Annelid Larval Morphology Greg W. Rouse

141

7. Larval Ecology of the Annelida Pei-Yuan Qian and Hans-Uwe Dahms

179

II Selected Groups of Annelida 8. Non-leech Clitellata Barrie G. M. Jamieson with Contributions by Marco Ferraguti

235

9. Hirudinida Mark E. Siddall, Alexandra E. Bely and Elizabeth Borda

393

10. Phyllodocida Fredrik Pleijel and Greg W. Rouse

431

11. Cirratuliformia Magdalena N. Halt, Greg W. Rouse, Mary E. Petersen and Fredrik Pleijel

497

N Reproductive Biology and Phylogeny of Annelida 12. Sabellida Greg W. Rouse, Elena Kupriyanova and Eijiroh Nishi

521

13. Spionida James A Blake

565

14. Problematic Annelid Groups Günter Purschke

639

Index

669

I General Reproduction and Phylogeny

CHAPTER

1

Annelid Phylogeny and Systematics Greg W. Rouse1 and Fredrik Pleijel2

1.1 INTRODUCTION Annelida is a group commonly referred to as segmented worms, and they are found worldwide in most habitats except the most arid or aerial. Earthworms and leeches are the most familiar annelids, but the bulk of the diversity of Annelida lies with polychaetes. These are found in nearly every marine habitat, from intertidal algal mats to the deepest sediments. There are pelagic polychaetes that swim or drift, preying on other plankton, and a few groups occurring in fresh water and moist terrestrial surroundings. Until recently, Annelida was split into three major groups given class rank: Polychaeta (bristleworms), Oligochaeta (earthworms etc.) and Hirudinida (leeches), though this has now been revised and revision is ongoing (see section 1.3). The first annelids were formally named by Linnaeus (1758) and today we estimate that the current number of accepted species level taxa is around 14,000 (9,000 polychaetes, 650 leeches, 150 branchiobdellids and 4,000 oligochaetes), though several thousand more have been named and are considered invalid.

1.2 MONOPHYLY OF ANNELIDA There have been a number of recent reviews on the monophyly and membership of Annelida (Rouse and Fauchald 1995, 1998; Westheide et al. 1999; McHugh 2000). In this section the morphological and molecular support for the monophyly and delineation of Annelida will be outlined. To date there has not been a combined analysis of morphology and molecular evidence at a broad level with comprehensive taxon sampling. 1

South Australian Museum Nth Terrace, Adelaide. S.A. 5000 Australia & Earth and Environmental Sciences, University of Adelaide SA. 5005 Australia 2 Department of Marine Ecology, Tjärnö Marine Biological Laboratory, Göteborg University, SE452 96 Strömstad, Sweden, and Muséum national d’Histoire naturelle, Département Systématique et Evolution, CNRS UMR 7138, ‘Systématique, Adaptation, Evolution’, 43, rue Cuvier, 75231 Paris Cedex 05, France.

" Reproductive Biology and Phylogeny of Annelida

1.2.1 Morphology and Monophyly The monophyly of Annelida is not well supported by anatomical features proposed to date, and only three are worthy of discussion: segmentation, chaetae, and nuchal organs representing another possible apomorphy. Metamerism (segmentation). Annelids have three body regions. In most annelids, the majority of the body is comprised of repeated units called segments. Each segment may be limited by septa dividing it from neighboring segments, and has a fluid-filled cavity within referred to as a coelom. Structures such as the excretory, locomotory and respiratory organs are generally repeated in each segment (Rouse and Pleijel 2001). Segments are formed sequentially in annelids and are established during development from growth zones located at the posterior end of the body; so the youngest segment in the body of an annelid is always the most posterior. The only parts of the annelid body that are not segmental are the head and a terminal post-segmental region called the pygidium (see Chapter 5). The head is comprised of two units, the prostomium and the peristomium. The postsegmental pygidium includes the zone from which new segments are proliferated during growth. The proposed homology of segmentation seen in annelids with that seen in Arthropoda has been used to unite the two as Articulata, a grouping that dates back to Cuvier (1817). The homology of this segmentation has been questioned, most recently by Seaver (2003), and arthropods are now viewed by many as closer to taxa such as Nematoda (Giribet 2002, 2003). This suggests that the form of segmentation seen in annelids may in fact represent an apomorphy. Pogonophora and Vestimentifera had been regarded as outside Annelida and the obvious segmentation (and chaetae) that they show in the posterior region was either overlooked by earlier authors or treated as nonhomologous to the annelid segmentation (see Rouse 2001). With their placement now this segmentation is instead viewed as homologous to that in annelids. With regard to echiurans, which appear to be unsegmented, Hessling and Westheide (2002), showed that differentiation of the echiuran nervous system proceeds from anterior to posterior, indicating the occurrence of a posterior growth zone. Their results can be interpreted as an indication that the echiuran unsegmented condition represent a loss and that they are derived from segmented ancestors (Hessling and Westheide 2002). However, it is also possible that what they show actually represents the plesiomorphic form of segmentation, retained in echiurans but further elaborated in (other) annelids. Further detailed study particularly with molecular sequence data is required to resolve this issue (see section 1.2.3). Chaetae. A distinctive feature of annelids are structures called chaetae. Chaetae (also called setae) are bundles of chitinous, thin-walled cylinders held together by sclerotinized protein. They are produced by a microvillar border of certain invaginated epidermal cells and so can be defined as cuticular structures that develop within epidermal follicles. Chaetae show

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#

a huge amount of variation, from long thin filaments (capillary chaetae) to stout multi-pronged hooks. It has been argued that the kind of chaetae found in annelids had evolved several times or was plesiomorphic feature for a large group of Metazoa and had been lost a number of times (see Orrhage 1973). This was based on the disparate occurrence of basically identical chaetae in the phyla Annelida, Brachiopoda, Pogonophora, Echiura, as well as in cephalopod molluscs (Brocco et al. 1974). As stated above (and see section 1.3.1), both Pogonophora and Echiura are now regarded as annelid taxa (Bartolomaeus 1995, 1997/98; McHugh 1997; Rouse and Fauchald 1997; Kojima 1998; Rousset et al. 2004). The position of Brachiopoda has been controversial, but evidence now suggests they are fairly close to annelids (Lüter and Bartolomaeus 1997; Stechmann and Schlegel 1999; Lüter 2000a; de Rosa 2001), though the homology of their chaetae with those of annelids is unresolved (Lüter 2000b). However, with further study there is a possibility that brachiopods might nest among annelids so that chaetae will still represent an apomorphy for Annelida. Further study on the proposed homology of annelid chaetae with those of the cephalopod Kölliker’s organ is warranted. Nuchal organs. These are ciliated, paired, chemosensory structures, innervated from the posterior part of the brain. They are present in nearly all polychaetes, and Rouse and Fauchald (1997) suggested that they may represent an apomorphy for Polychaeta. This has been challenged by other authors, who suggest that nuchal organs may be an apomorphy for Annelida as a whole and have been lost in Clitellata (Purschke et al. 2000). This latter scenario now appears to be supported by recent phylogenetic studies (see section 1.3.2).

1.2.2 Molecular Sequence Data The earliest molecular studies focussing on the status and delineation of Annelida can be found in Winnepenninckx et al. (1995). They used 18S rRNA sequences to examine relationships among protostome worms such as Annelida, Echiura, Nemertea, Pogonophora and Vestimentifera. They only included two annelids (Lanice and Eisenia) in their study and these did not form a clade in the parsimony analysis. McHugh (1997) and Kojima (1998) then found Clitellata and Pogonophora clustered among various polychaetes using analyses of the sequence of a nuclear gene, elongation factor-1α. The former study also found that Echiura nested among polychaetes. Their taxon sampling was such that the possibility of a number of other protostome taxa also being included in Annelida was not assessed. Brown et al. (1999) then studied relationships within Annelida using DNA sequence data from three genes and a broader taxon sample from among annelids and other protostomes. They also found clitellates and pogonophores nested among annelids and also Sipuncula. Martin (2001) analyzed available sequences of 18S rRNA with the primary aim of assessing the placement of Clitellata. He could not recover a monophyletic Annelida without also including taxa such as Mollusca and Sipuncula.

$ Reproductive Biology and Phylogeny of Annelida Subsequent phylogenetic analyses using molecular sequence data with more comprehensive taxon sampling have yet to show a monophyletic Annelida also in its current formulation with Pogonophora (and Vestimentifera) and Echiura included. Subsequent studies that have large samples of protostomes also consistently show taxa from Mollusca, Sipuncula, Brachiopoda and Phoronida nested among annelid taxa (Struck et al. 2002; Bleidorn et al. 2003a, b; Jördens et al. 2004). To date, large-scale molecular sequence studies have not been very encouraging, but no doubt there will be much larger analyses forthcoming. Morphological studies are also essential and critical gaps in our knowledge about basic anatomy of many groups have been revealed (see chapters 8 and 9 and Rouse and Pleijel 2001).

1.3. ANNELIDA SUBGROUPS 1.3.1 Pogonophora, Vestimentifera and Echiura Echiura (spoon or anchor worms), were classified as a class of annelids in the late 19th century (as Echiuroidea by Sedgwick 1898). They were then excluded from Annelida by Newby (1940), after his study on embryology and development in Urechis. However, recent evidence now suggests they are in fact annelids (e.g. McHugh 1997; Hessling and Westheide 2002), though their placement within the group is unresolved. The former phyla Pogonophora and Vestimentifera have also recently become regarded as a single, clearly annelid, group (Bartolomaeus 1995, 1997/98; Nielsen 1995; Rouse and Fauchald 1995, 1997; McHugh 1997; Kojima 1998; Rousset et al. 2004), and are now referred to by many by their original name, Siboglinidae (see McHugh 1997; Rouse and Fauchald 1997). Clear evidence that Vestimentifera is nested inside Siboglinidae has been provided from morphological (Rouse 2001; Schulze 2003) or molecular evidence (Halanych et al. 2001), or both (Rousset et al. 2004).

1.3.2 Clitellata, Hirudinida and Oligochaetes In recent years it has become well recognized that Hirudinida is nested within Oligochaeta and that giving both these taxa the rank of class renders the latter group paraphyletic. This idea does have a long history (see Chapter 8) and it may be some time before Class Oligochaeta and Class Hirudinida are eliminated. Comprehensive phylogenetic studies using molecular sequence data and morphology provide strong support that Lumbriculida is the sister group to the ectoparastitic clade comprised Hirudinida (also carnivorous), Acanthobdellida and Branchiobdellida (Martin 2001; Siddall et al. 2001; Erséus and Källersjö 2004; see also Jamieson et al. 2002) and should be referred to either as Oligochaeta (Siddall et al. 2001), or Clitellata (Martin 2001; Erséus and Källersjö 2004). There are arguments for using either name with respect to a monophyletic taxon (see Chapters 8 and 9) but we have used the name Clitellata in this volume. We employ the term ‘oligochaetes’ as an informal name for the paraphyletic

Annelid Phylogeny and Systematics

%

group non-leech Clitellata. It will be up to the specialists of the group to resolve the issue in the longer term. While the monophyly of Clitellata is well supported and the placement of Hirudinida, Acanthobdellida and Branchiobdellida as a clade well inside that group, is now beyond question, there still remains the major question as to what is the sister group to Clitellata?

1.3.3 Polychaetes There is increasing evidence that along with Echiura and Pogonophora, Clitellata may well belong inside Polychaeta (McHugh 1997; Westheide 1997; Westheide et al. 1999), and recent molecular studies all show them as nested among polychaetes (Struck et al. 2002; Bleidorn et al. 2003a, 2003b; Jördens et al. 2004), although none to date have provided robust support for any sister group relationship with a particular polychaete group. No doubt such a relationship will be recovered soon and either the name Polychaeta or Annelida will be redundant. Although largely arbitrary, we prefer to retain the name Annelida as it agrees better with the traditional use and inclusiveness. The informal name ‘polychaetes’ will be used in this volume for the paraphyletic constellation non-clitellate annelids, similar to the term ‘oligochaetes’ as explained above. As in Rouse and Pleijel (2001) the formal taxon names Polychaeta and Oligochaeta are not used in this volume. Once upon a time there was Polychaeta, and that group was conveniently separated into the equally-sized groups Errantia and Sedentaria (e.g., Fauvel 1923, 1927; Hartman 1959a, 1959b; Day 1963). Since Dales (1962), Fauchald (1977), Rouse and Fauchald (1997) and others, things have not been so simple, and this situation is likely to continue till we have a better understanding of the relationships about the more basal relationships among the subgroups. One major problem in this context that requires solution is the position of the root for the annelid branch (see section 1.5).

1.4 SYSTEMATIZATION USED IN THIS VOLUME As is abundantly clear from the foregoing, the systematics of annelids is undergoing major revision. We have chosen to implement the most recent comprehensive systematic treatment (Rouse and Fauchald 1997) (see Table 1.1) but emphasize that many changes are to come in the future, some of which are foreshadowed in the taxon chapters in this book. It would appear that the intensive efforts of a number of workers is resulting in significant movement towards a stable classification for Clitellata (see chapters 8 and 9). This cannot be said for the remaining annelids. The history of polychaete classification is reviewed in Fauchald and Rouse (1997) and Rouse and Pleijel (2001) and is not covered here. Rouse and Fauchald (1997) used various cladistic analyses including ‘complete’ and a ‘restricted’ taxon samples and explored different techniques for character coding. There was

& Reproductive Biology and Phylogeny of Annelida partial incongruence between the different analyses and the choice of topology used to revise the taxonomy was largely arbitrary. Given that the overall topology of their complete analyses were incongruous with the restricted analyses, the placements and delineations of a number of these taxa should be further investigated. For example, the position of the clades (Arenicolidae, Capitellidae, Maldanidae, Acrocirridae, Cirratulidae, Flabelligeridae and Oweniidae) differs markedly between the complete and restricted analyses under one coding. Additionally, in a subsequent analysis, Rouse (1999) added a number of larval characters to the character set of Rouse and Fauchald (1997). This resulted in some slightly different tree topologies, particularly with regard to Chaetopteridae and Oweniidae. While these are not taken into consideration in our systematic treatment here, they deserve further attention. Clitellata (Fig. 1.1A). The name Clitellata, derived from clitellum (Latin) meaning saddle bag, was introduced by Michaelsen (1919, 1928) for aeolosomatids, leeches and oligochaetes. Aeolosomatids were removed from the Oligochaeta by Brinkhurst and Jamieson (1971) and are here referred to as Annelida incertae sedis, otherwise we apply the name in Michaelsen’s (1928) sense with Hirudinea nested inside a paraphyletic Oligochaeta. As discussed above Clitellata is unlikely to be the sister group to the remaining Annelida, although its precise placement among annelids is as yet uncertain. Apart from molecular data (Erséus and Källersjö 2004; Siddall et al. 2001) the monophyly of Clitellata is supported by the presence of the clitellum, the organization of the reproductive system, several ultrastructural features in the sperm (see Rouse and Fauchald 1995 and references within) and, possibly, by the loss of nuchal organs (Purschke et al. 2000) (see also chapters 8 and 9). Scolecida (Fig. 1.1B). The name Scolecida, derived from skolex (Greek) meaning worm, was reappraised by Rouse and Fauchald (1997). The name is derived from Scoleciformia, a name introduced by Benham (1896) for a similar group of taxa. Only two apomorphies support the clade Scolecida in Rouse and Fauchald (1997), the presence of parapodia with similar rami and the possession of two or more pairs of pygidial cirri, and these are homoplastic. In many ways, this group represents the simple-bodied forms of polychaetes and it is likely that further analysis will show that it is not monophyletic. In fact to date no molecular sequence analyses have recovered any assemblage like Scolecida. Palpata. Palpata was a new name coined by Rouse and Fauchald (1997) and such a group of polychaetes has never been formulated before. Virtually all non-Scolecida polychaetes, except a few incertae sedis taxa, were placed into Palpata. The name is based on an apomorphy for the group, the presence of palps. Palps can be divided into two structurally different groups, grooved ‘feeding’ palps and ventral, ‘sensory’ palps (Orrhage 1980). Feeding palps usually have ciliated paths, often located in a longitudinal groove, giving each palp a U-shaped cross-section. Ventral sensory palps are morphologically more uniform than grooved palps. In

Annelid Phylogeny and Systematics

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most cases they are tapering or digitiform and relatively short, compared to grooved palps. Given the fact that the presence of palps and a limited peristomium were the only synapomorphies supporting this taxon in Rouse and Fauchald (1997), the delineation of the Palpata requires further investigation. Palpata contains two major clades Aciculata and Canalipalpata. Aciculata. Aciculata was a new name erected by Rouse and Fauchald (1997) for one of most the strongly supported clades in their analyses. The name refers to one of the apomorphies for the group; the presence of a particular chaetal type called aciculae. Aciculae are stout chaetae that differ from others in that much or all of the chaeta remains internalized in the parapodium. Numerous other features such as the presence of ventral sensory palps, prostomial antennae, dorsal cirri, ventral cirri, one pair of pygidial cirri, and segmental organs in most segments were proposed to be apomorphic for Aciculata. Aciculata is divided here into three major clades Amphinomida, Eunicida and Phyllodocida, and also with a few taxa as incertae sedis. Amphinomida. A taxon, Amphinomae, equivalent to Amphinomida dates back to Savigny (1822). Amphinomida, today divided into Amphinomidae and Euphrosinidae, was first used as a taxon name by Dales (1962), but names such as Amphinomorpha have also been used. Although Amphinomida is well supported by a number of apomorphies, including calcareous chaetae and nuchal organs forming a caruncle, both their position within Aciculata and the interrelationships within the group are uncertain. Rouse and Fauchald (1997) in their classification included Amphinomidae and Euphrosinidae in Eunicida, whereas Rouse and Pleijel (2001) instead treated them as a separate taxon Amphinomida. There is no systematic chapter in this book especially covering Amphinomida. Eunicida (Fig. 1.1C). The name Eunicida was first used by Dales (1962), but encompassed a well-delineated group known prior to that as Eunicea, Euniformia, Eunicimorpha or simply Eunicidae. This grouping included polychaetes with a ventral muscularized pharynx with ventral mandibles and dorsal maxillae (and a few that had lost jaws) and is very likely a monophyletic assemblage. Another putative apomorphy for the clade would appear to be the peristomium forming two rings, although this feature is not present in all the subgroups. Rouse and Fauchald (1997) expanded the traditionally delineated Eunicida to include Amphinomidae and Euphrosinidae and this is not accepted here. Eunicida, following Rouse and Pleijel (2001), includes Dorvilleidae, Eunicidae, Hartmaniellidae, Histriobdellidae, Lumbrineridae, Oenonidae and Onuphidae. There is no systematic chapter in this book especially covering Eunicida. Phyllodocida (Fig. 1.1D). The name Phyllodocida was coined by Dales (1962) and was subsequently used by other workers, such as Fauchald (1977) and Pettibone (1982) for essentially the same group of polychaetes. Prior to Dales (1962), the group had never been formulated as a monophyletic taxon. Rouse and Fauchald (1997) found strong support for

 Reproductive Biology and Phylogeny of Annelida the monophyly of this group, indicated by the ventral position of sensory palps, the presence of anterior enlarged cirri, the loss of dorsolateral folds, the presence of an axial muscular proboscis, and the presence of compound chaetae with a single ligament. There is a strong body of opinion that Phyllodocida, as formulated here, may be paraphyletic (see Section 1.5). A major difference in the formulation of Phyllodocida by Rouse and Fauchald (1997), and that of previous workers, was that they placed Myzostomida (as Myzostomatidae) in the group and this is accepted here (but see Eeckhaut et al. 2000). Rouse and Fauchald (1997) did not further subdivide Phyllodocida, since their analyses resulted in dramatically different topologies for various Phyllodocida taxa. Based on the results of Pleijel and Dahlgren (1998), two names are used here for clades within Phyllodocida; Aphroditiformia (or scaleworms) and Nereidiformia. The remaining taxa in Phyllodocida are treated as ‘unplaced’. Aciculata unplaced. Rouse and Fauchald (1997) placed Aberranta (as Aberrantidae), Nerillidae (Fig. 1.1E) and Spinther (as Spintheridae) as parts of Aciculata, but did not include them in any other sub-taxon of the group. These three taxa have never been related to each other and are not to be regarded as closely related by being grouped together here. Of these taxa, only Spinther actually has aciculae, but all three almost certainly will be found to have sister groups with different members of Aciculata. Canalipalpata. Canalipalpata (referring to the presence of grooved palps) is a name that was first used by Rouse and Fauchald (1997). The Canalipalpata is a massive group of polychaetes that encompasses around half the number of described species of polychaetes. The only apomorphy for Canalipalpata is the presence of grooved palps and so the clade must be regarded as weakly supported. The groove along each palp is longitudinal and ciliated and these palps, in contrast to those of Aciculata, are feeding structures. Rouse and Fauchald (1997) identified three major clades within the Canalipalpata (Sabellida, Spionida and Terebellida) and a number of taxa that are regarded as incertae sedis for the group. Sabellida (Fig. 1.2B, C). The apomorphy for Sabellida, as formulated by Rouse and Fauchald (1997), was the fusion of the prostomium with the peristomium, hence it is weakly supported. This name has been used to contain Sabellariidae, Sabellidae and Serpulidae (Fitzhugh 1989) and these three were included by Rouse and Fauchald (1997), plus two other taxa. A dramatic shift compared with traditional systematics was that Rouse and Fauchald (1997) placed Siboglinidae (formerly outside polychaetes as Pogonophora and Vestimentifera) as part of Sabellida. They also included Oweniidae, a taxon that has a ‘chequered’ systematic history as a polychaete. It should be noted that in further analyses by Rouse (1999a, 2000a), Oweniidae did not group with the remaining Sabellida, and Chaetopteridae (included here in as part of Spionida) did. A recent combined analysis of molecular and morphological data (Rousset et al. 2004) also did not recover the Sabellida as formulated here. Clearly further study is required. In addition, similarities between Sabellariidae and

Annelid Phylogeny and Systematics



Pectinariidae (in Terebellida) also deserve further study. Sabellariidae has previously been considered as part of Terebellida (e.g., Fauchald 1977). Terebellida (Cirratuliformia, Terebelliformia) (Fig. 1.2C, D). The name Terebellida was first used by Dales (1962) and included Ampharetidae, Pectinariidae and Terebellidae, all polychaetes having multiple grooved palps. Prior to this, the names Terebellomorpha or Terebelliformia had often been used for essentially the same grouping of taxa. Terebellida was expanded by Rouse and Fauchald (1997) to include a clade in which most have a single pair of palps (e.g., Acrocirridae, Cirratulidae, Flabelligeridae). Rouse and Fauchald (1997) identified several clear synapomorphies for this overall grouping, namely the presence of a first segment with no chaetae, a gular membrane and a heart body. The clade within Terebellida that has taxa with a single pair of palps (with exceptions such as some Cirratulidae with numerous palps and Ctenodrilinae and Fauveliopsidae with none) is here referred to as Cirratuliformia, a name that has been used previously for a somewhat similar grouping (e.g., Fauchald 1977). It contains Acrocirridae, Cirratulidae, Fauveliopsidae, Flabelligeridae, Poeobiidae (and Flota) and Sternaspis. The clade comprised of Alvinellidae, Ampharetidae, Pectinariidae and Terebellidae is referred to here as Terebelliformia, also a fairly ‘traditional’ formulation. Spionida (Fig. 1.2E). The name Spionida was first used by Dales (1962) to contain Spionidae and a number of similar groups, as well as taxa such as Paraonidae and Sabellariidae. These groups are not considered closely related today, and Spionida in Rouse and Pleijel (2001) contained Apistobranchus, Chaetopteridae, Magelonidae, Spionidae, Heterospio, Poecilochaetus, Trochochaeta, and Uncispionidae. In Chapter 13, Heterospio, Poecilochaetus, Trochochaetus and Uncispio are all treated as members of Spionidae. The positions of Chaetopteridae and Magelonidae deserve further investigation. The synapomorphies for Spionida listed by Rouse and Fauchald (1997) were the presence of a pair of peristomial grooved palps, nuchal organs forming posterior projections, and anterior excretory nephridia and posterior segmental organs for gamete release. Canalipalpata unplaced. Rouse and Fauchald (1997) placed Polygordiidae and Protodrilida (as Protodriloididae, Protodrilidae and Saccocirridae) as part of Canalipalpata, but did not place them within any other sub-taxon. In their complete cladistic analyses, these taxa formed a clade that was either associated with taxa that belong within Canalipalpata, Scolecida or were part of a basal polytomy of Polychaeta, so their decision was arbitrary. However, there is good evidence to support Protodrilida being placed in Canalipalpata somewhere (Purschke and Jouin 1988), perhaps near Spionida. On the other hand, placement of Polygordiidae in Canalipalpata must be regarded as suspect. Rouse and Fauchald (1997) made scoring errors in regard to Polygordiidae, the most important being that they cannot be regarded as having grooved palps. It was suggested by Rouse and Pleijel (2001) that investigation of a sister group relationship with, or within, Opheliidae may be worthwhile, and this is actually an old idea (e.g. Giard 1880).



Reproductive Biology and Phylogeny of Annelida

Annelida unplaced. Rouse and Fauchald (1997) included Aeolosomatidae, Potamodrilus (as Potamodrilidae), Parergodrilidae and Psammodrilidae in their complete taxon set analyses. In one analysis, these taxa either fell as part of a large basal polytomy of polychaetes or Aeolosomatidae, Potamodrilus and Parergodrilidae formed a clade with Ctenodrilidae, and Psammodrilidae grouped with Capitellidae, Arenicolidae and Maldanidae. In another analysis, Aeolosomatidae, Potamodrilus and Parergodrilidae either formed a grade with respect to Polygordiidae, Protodrilidae and Protodriloididae, or they were a basal clade of polychaetes. Psammodrilidae either were a basal polychaete group, or were sister group to a large clade that mainly comprised taxa with grooved palps. Thus, it was not possible to place these taxa with any confidence. Prior to the study by Rouse and Fauchald (1997), Aeolosomatidae and Potamodrilus were usually considered in relation to Clitellata, either as a member of the group (Bunke 1967), or more recently as the sister group to Clitellata (Bunke 1985). This was then rejected by Bunke (1986), who also could not relate them to any group of polychaetes, thus leaving the two taxa ‘isolated’. The most recent analysis was from a molecular persepective (Struck et al. 2002), and this study found no support for a relationship of Aeolosomatidae with Clitellata or with any other particular annelid group. Fauchald (1977) grouped Parergodrilidae with Ctenodrilidae, but did not justify this decision. Otherwise, the group has also been treated as an ‘isolated’ group of polychaetes. A recent molecular study by Jördens et al. (2004) indicated that Parergodrilidae may be closely related to orbiniids; the position of the terrestrial polychaete Hrabeiella, however, was unconclusive. Psammodrilidae has been treated as a singular group of polychaetes since they were first discovered by Swedmark (1952). Rouse and Fauchald (1997) suggested that a relationship of Psammodrilidae with Arenicolidae and Maldanidae should be assessed, as proposed by Meyer and Bartolomaeus (1996; 1997), but the present anatomical evidence is weak.

1.5 ROOTING THE ANNELID TREE In a review of the fossil record of annelids Rouse and Pleijel (2001) suggested that the oldest unequivocal fossil polychaetes, such as Canadia from the Cambrian period, belong within Phyllodocida. This view was challenged by Eibye-Jacobsen (2004), who argued that there are no synapmorphies that would argue for placing Canadia in Phyllodocida, but did agree they were annelids of some sort. No other fossil polychaetes from the Cambrian can be unequivocally assigned to extant annelid taxa either. There are several likely appearances from the Ordovician, including Serpulidae, Spionidae and the radiation of Eunicida (Rouse and Pleijel 2001). Ensuing appearances suggest that by the end of the Carboniferous most major polychaetes lineages had appeared. The exception appears to be Scolecida, with the earliest known fossils being the dubious Archarenicola

Annelid Phylogeny and Systematics

!

Table 1.1 Systematization of Annelida used in this volume.

Major taxa

Less inclusive taxa

Scolecida

Amphinomida Eunicida

Aciculata

Palpata

Phyllodocida

Aciculata incertae sedis Sabellida Spionida Canalipalpata

Terebellida, Terebelliformia Terebellida, Cirratuliformia

Canalipalpata incertae sedis Clitellata Annelida incertae sedis

Arenicolidae, Capitellidae, Maldanidae, Cossuridae, Opheliidae, Orbiniidae, Paraonidae, Parergodrilidae?, Questidae, Scalibregmatidae Amphinomidae, Euphrosinidae Dorvilleidae, Eunicidae, Hartmaniellidae, Histriobdellidae, Lumbrineridae, Oenonidae, Onuphidae Acoetidae, Aphroditidae, Chrysopetalidae, Eulepethidae, Glyceridae, Goniadidae, Hesionidae, Ichthyotomus, Iospilidae, Lacydonia, Lopadorhynchidae, Myzostomida, Nautiliniellidae, Nephtyidae, Nereididae, Paralacydonia, Pholoidae, Phyllodocidae, Pilargidae, Pisionidae, Polynoidae, Pontodora, Sigalionidae, Sphaerodoridae, Syllidae, Typhloscolecidae, Tomopteridae Aberranta, Nerillidae, Spinther Oweniidae, Sabellariidae, Sabellidae, Serpulidae, Siboglinidae Apistobranchus Chaetopteridae, Magelonidae, Spionidae Alvinellidae, Ampharetidae, Pectinariidae, Terebellidae, Trichobranchidae Acrocirridae, Cirratulidae, Fauveliopsidae, Flabelligeridae, Poeobius, Sternaspis Polygordiidae, Protodrilidae, Protodriloididae, Saccocirridae (see Chapters 8 and 9) Aeolosomatidae, Echiura, Potamodrilidae, Psammodrilidae

(Arenicolidae) from the Triassic, and one assignable to Paraonidae from the Cretaceous. With the rooting option (based on Rouse and Fauchald 1997) employed in Figure 1.3A, it appears that some of the earliest appearing fossil polychaetes belong to derived clades (e.g., Eunicida and possibly

" Reproductive Biology and Phylogeny of Annelida

Colour Figure

Fig. 1.1. A. Fletcherodrilus fasciatus (Megascolecidae, Clitellata), Australia. B. The orbiniid Scoloplos armiger from Sweden. C. Female of the dorvilleid Ophryotrocha sp. with egg sac from Iceland. D. The syllid Amblyosyllis sp. from Iceland. E. Female of the nerillid Nerilla antennata from Iceland. Figure 1A courtesy of Conrad Hoskin via B.G.M. Jamieson; others by G.W. Rouse.

Annelid Phylogeny and Systematics

#

Colour Figure

Fig. 1.2. A. Female Osedax frankpressi, a siboglinid from a whale fall in Monterey Canyon. B. Ventral view of the sabellid Pseudopotamilla reniformis from Iceland in its tube with crown extended. C. Female of the trichobranchid Terebellides sp. from Japan. D. The cirratulid Cirratulus sp. from Japan. E. Spawning female of the spionid Malacoceros fuliginosus from Iceland. All figures by G.W. Rouse.

$ Reproductive Biology and Phylogeny of Annelida

Fig. 1.3. A. Cladogram of relationships among the major groups of annelids. Based on Pleijel and Rouse (2001, 2003). The ‘disconnected’ taxa could be attached on many places in the tree, and the broken line to Spinther also indicate uncertainty. B. Same tree but unrooted.

Annelid Phylogeny and Systematics

%

Phyllodocida). This could be interpreted in two ways: 1) the root placement in Figure 1.3A is wrong, and so Aciculata, comprised Amphinomida, Eunicida and Phyllodocida, may in fact represent a paraphyletic ‘stem’ group for the rest of polychaetes; 2) a number of major polychaete clades had already evolved in, or before, the ‘Cambrian explosion’, but fossils have not yet been found. A third possibility is that the overall tree topology may be profoundly incorrect. If we accept that the topology shown in Fig. 1.3A is correct, but do not root the tree, then a diagram as shown in Fig. 1.3B is the result. This is the most conservative representation of our current understanding of annelid relationships. Westheide (1997; see also Westheide et al. 1999) suggested that the basic (i.e., pleisomorphic) ‘body plan’ of Annelida comprised, among other features, the following: (1) an unregionated segmented body, (2) biramous parapodia with numerous chaetae, (3) dorsal chaetae with a protective function, (4) gonads in all segments, (5) metanephridia, (6) prostomium with paired palps and presumably three antennae, (7) nuchal organs, (8) simple ciliated foregut (dorsolateral folds), at least in the juvenile stages, (9) collagenous cuticle, (10) epibenthic mode of life. Item (6) is only found in parts of Aciculata and item (3), dorsal protective chaetae, is arguably only present in a few parts of Aciculata, namely Amphinomida, Chrysopetalidae and Aphroditidae (the latter two both Phyllodocida). The only possible conclusion then for rooting a cladogram of Annelida on this suggestion (see Fig. 1.3B) would be with taxa from Aciculata. This may result in a paraphyletic Phyllodocida, Eunicida or Amphinomida, depending on which taxon is used as the root. Our pessimistic conclusion is that we at present have no knowledge whatsoever about the root position of annelids, and that the most ‘honest’ representation is the one in Fig. 1.3B. This, however, also means that we at present cannot identify a single clade within the group, unless basing it on the assumption that the root is situated elsewhere on the annelid branch. But we are optimistic in believing that this state of affairs is about to change in the near future.

1.6 ACKNOWLEDGEMENTS Financial for FP support was obtained from Formas, dnr 2004-0085, and for GWR from the South Australian Museum.

1.7 LITERATURE CITED Bartolomaeus, T. 1995. Structure and formation of the uncini in Pectinaria koreni, Pectinaria auricoma (Terebellida) and Spirorbis spirorbis (Sabellida): implications for annelid phylogeny and the position of the Pogonophora. Zoomorphology 115: 161-177. Bartolomaeus, T. 1997/98. Chaetogenesis in polychaetous Annelida — Significance for annelid systematics and the position of the Pogonophora. Zoology-Analysis of Complex Systems 100: 348-364.

& Reproductive Biology and Phylogeny of Annelida Benham, W. B. 1896. Archiannelida, Polychaeta, Myzostomidaria. Pp. 241-334. In S. F. Harmer and A. E. Shipley (eds), The Cambridge Natural History, Macmillan, London. Bleidorn, C., Vogt, L. and Bartolomaeus, T. 2003a. A contribution to sedentary polychaete phylogeny using 18S rRNA sequence data. Journal of Zoological Systematics and Evolutionary Research 41: 186-195. Bleidorn, C., Vogt, L. and Bartolomaeus, T. 2003b. New insights into polychaete phylogeny (Annelida) inferred from 18S rDNA sequences. Molecular Phylogenetics & Evolution 29: 279-288. Brinkhurst, R.O. and Jamieson, B.G.M. 1971. Aquatic Oligochaeta of the World. Oliver and Boyd, Edinburgh. 860 pp. Brocco, S. L., O’Clair, R. and Cloney, R. A. 1974. Cephalopod integument: the ultrastructure of Kölliker’s organs and their relationship to setae. Cell and Tissue Research 151: 293-308. Brown, A. C., Rouse, G. W., Hutchings, P. and Colgan, D. 1999. Assessing the usefulness of histone H3, U2 snRNA and 28S rDNA in analyses of polychaete relationships. Australian Journal of Zoology: 499-516. Bunke, D. 1967. Zur Morphologie und Systematik der Aeolosomatidae Beddard 1895 und Potamodrilidae nov. fam. (Oligochaeta). Zoologische Jahrbücher, Abteilung für Anatomie und Ontogenie der Tiere 94: 187-368. Bunke, D. 1985. Ultrastructure of the spermatozoon and spermiogenesis in the interstitial annelid Potamodrilus fluviatilis. Journal of Morphology 185: 203-216. Bunke, D. 1986. Ultrastructural investigation on the spermatozoon and its genesis on Aeolosoma litorale with considerations on the phylogenetic implications for the Aeolosomatidae. Journal of Ultrastructure and Molecular Structure Research 95: 113-130. Cuvier, G. 1817. Le régne animal distribué d’après son organisation, pour servir de base à l’histoire naturelle des animaux et d’introduction à l’anatomie comparée, Deterville, Paris, 255 pp. Dales, R. P. 1962. The polychaete stomodeum and the interrelationships of the families of the Polychaeta. Proceedings of the Zoological Society of London 139: 289-328. Day, J. H. 1963. The polychaete fauna of South Africa. Bulletin of the British Museum (Natural History), Zoology Series 10: 381-445. de Rosa, R. 2001. Molecular data indicate the protostome affinity of brachiopods. Systematic Biology 50: 848-859. Eeckhaut, I., McHugh, D., Mardulyn, P., Tiedemann, R., Monteyne, D., Jangoux, M. and Milinkovitch, C. 2000. Myzostomida: a link between trochozoans and flatworms? Proceedings of the Royal Society, Series B 267: 1383-1392. Eibye-Jacobsen, D. 2004. A reevaluation of Wiwaxia and the polychaetes of the Burgess Shale. Lethaia 37: 317-335. Erséus, C. and Källersjö, M. 2004. 18S rDNA phylogeny of Clitellata (Annelida). Zoologica Scripta 33: 187-196. Fauchald, K. 1977. The polychaete worms. Definitions and keys to the orders, families and genera. Natural History Museum of Los Angeles County. Science Series 28: 1-188. Fauchald, K. and Rouse, G. W. 1997. Polychaete systematics: Past and present. Zoologica Scripta 26: 71-138. Fauvel, P. 1923. Polychètes Errantes. Faune de France, Paris 5: 1-488. Fauvel, P. 1927. Polychètes sédentaires. Addenda aux Errantes, Archiannelides, Myzostomaires. Faune de France, Paris 16: 494 pp.

Annelid Phylogeny and Systematics

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Fitzhugh, K. 1989. A systematic revision of the Sabellidae-CaobangiidaeSabellongidae complex (Annelida: Polychaeta). Bulletin of the American Museum of Natural History 192: 1-104. Giard, A. 1880. On the affinities of the genus Polygordius with the annelids of the family Opheliidae. Annals and Magazines of Natural History 6: 324-326. Giribet, G. 2002. Current advances in the phylogenetic reconstruction of metazoan evolution. A new paradigm for the Cambrian explosion? Molecular Phylogenetics and Evolution 24: 345-357. Giribet, G. 2003. Molecules, development and fossils in the study of metazoan evolution; Articulata versus Ecdysozoa revisited. Zoology 106: 303-326. Halanych, K. M., Feldman, R. A. and Vrijenhoek, R. C. 2001. Molecular evidence that Sclerolinum brattstromi is closely related to vestimentiferans, not to frenulate pogonophorans (Siboglinidae, Annelida). Biological Bulletin 201: 65-75. Hartman, O. 1959a. Catalogue of the polychaetous Annelids of the world. Part I. Allan Hancock Foundation Publications. Occasional Paper 23: 1-353. Hartman, O. 1959b. Catalogue of the polychaetous Annelids of the world. Part II. Allan Hancock Foundation Publications. Occasional Paper 23: 355-628. Hessling, R. and Westheide, W. 2002. Are Echiura derived from a segmented ancestor? Immunohistochemical analysis of the nervous system in developmental stages of Bonellia viridis. Journal of Morphology 252: 100-113. Jamieson, B. G. M., Tillier, S., Tillier, A., Justine, J.-L., Ling, E., James, S., McDonald, K. and Hugall, A. F. 2002. Phylogeny of the Megascolecidae and Crassiclitellata (Annelida, Oligochaeta): combined versus partitioned analysis using nuclear (28S) and mitochondrial (12S, 16S) rDNA. Zoosystema 24(4): 707-734. Jördens, J., Struck, T. and Purschke, G. 2004. Phylogenetic inference regarding Parergodrilidae and Hrabeiella periglandulata (‘Polychaeta’, Annelida) based on 18S rDNA, 28S rDNA and COI sequences. Journal of Zoological Systematics and Evolutionary Research 42: 270-280. Kojima, S. 1998. Paraphyletic status of Polychaeta suggested by phylogenetic analysis based on the amino acid sequences of Elongation Factor 1-alpha. Molecular Phylogenetics and Evolution 9: 255-261. Linnaeus, C. 1758. Systema Naturae, 10th ed., Stockholm. Lüter, C. 2000a. The origin of the coelom in Brachiopoda and its phylogenetic significance. Zoomorphology 120: 15-28. Lüter, C. 2000b. Ultrastructure of larval and adult setae of Brachiopoda. Zoologischer Anzeiger 239: 75-90. Lüter, C. and Bartolomaeus, T. 1997. The phylogenetic position of Brachiopoda— a comparison of morphological and molecular data. Zoologica Scripta 26: 245-253. Martin, P. 2001. On the origin of the Hirudinea and the demise of the Oligochaeta. Proceedings of the Royal Society of London - Series B: Biological Sciences 268: 1089-1098. McHugh, D. 1997. Molecular evidence that echiurans and pogonophorans are derived annelids. Proceedings of the National Academy of Sciences of the United States of America 94: 8006-8009. McHugh, D. 2000. Molecular phylogeny of the Annelida. Canadian Journal of Zoology 78: 1873-1884. Meyer, K. and Bartolomaeus, T. 1996. Ultrastructure and formation of the hooked setae in Owenia fusiformis delle Chiaje, 1842 - implications for annelid phylogeny. Canadian Journal of Zoology 74: 2143-2153. Meyer, R. and Bartolomaeus, T. 1997. Ultrastruktur und morphogenese der hakenborsten bei Psammodrilus balanoglossoides— bedeutung fur die stellung der Psammodrilida (Annelida). Microfauna Marina 11: 87-113.

 Reproductive Biology and Phylogeny of Annelida Michaelsen, W. 1919. Über die Beziehungen der Hirudineen zu den Oligochäten. Jahrbuch der Hamburgischen wissenschaftlichen Anstalten, Hamburg 36: 131153. Michaelsen, W. 1928. Dritte Klasse der Vermes Polymera (Annelida) Clitellata = Gürtelwürmer. Pp. 1-352. In W. Kükenthal and T. Krumbach (eds), Handbuch der Zoologie 2(2) Lief. 8. Newby, W. W. 1940. The embryology of the echiuroid worm Urechis caupo. Memoirs of the American Philosophical Society 16: 1-219. Nielsen, C. 1995. Animal Evolution, Oxford University Press, Oxford. Orrhage, L. 1973. Light and electron microscope studies of some brachiopod and pogonophoran setae. Zeitschrift für Morphologie und Ökologie der Tiere 74: 253270. Orrhage, L. 1980. On the structure and homologues of the anterior end of the polychaete families Sabellidae and Serpulidae. Zoomorphologie 96: 113-168. Pettibone, M. H. 1982. Annelida. Pp. 1-43. In S. P. Parker (ed.), Synopsis and Classification of Living Organisms, vol. 2., McGraw-Hill Book Co, New York. Pleijel, F. and Dahlgren, T. 1998. Position and delineation of Chrysopetalidae and Hesionidae (Annelida, Polychaeta, Phyllodocida). Cladistics 14: 129-150. Purschke, G., Hessling, R. and Westheide, W. 2000. The phylogenetic position of the Clitellata and the Echiura— on the problematic assessment of absent characters. Journal of Zoological Systematics & Evolutionary Research 38: 165-173. Purschke, G. and Jouin, C. 1988. Anatomy and ultrastructure of the ventral pharyngeal organs of Saccocirrus (Saccocirridae) and Protodriloides (Protodriloidae fam. n.) with remarks on the phylogenetic relationships within Protodrilida (Annelida: Polychaeta). Journal of Zoology 215: 405-432. Rouse, G. W. 1999. Trochophore concepts: ciliary bands and the evolution of larvae in spiralian Metazoa. Biological Journal of the Linnean Society 66: 411-464. Rouse, G. W. 2001. A cladistic analysis of Siboglinidae Caullery, 1914 (Polychaeta, Annelida): formerly the phyla Pogonophora and Vestimentifera. Zoological Journal of the Linnean Society 132: 55-80. Rouse, G. W. and Fauchald, K. 1995. The articulation of annelids. Zoologica Scripta 24: 269-301. Rouse, G. W. and Fauchald, K. 1997. Cladistics and polychaetes. Zoologica Scripta 26: 139-204. Rouse, G. W. and Fauchald, K. 1998. Recent views on the status, delineation and classification of the Annelida. American Zoologist 38: 953-964. Rouse, G. W. and Pleijel, F. 2001. Polychaetes, Oxford University Press, London, 354 pp. Rousset, V., Rouse, G. W., Siddall, M. E., Tillier, A. and Pleijel, F. 2004. The phylogenetic position of Siboglinidae (Annelida), inferred from 18S rRNA, 28S rRNA, and morphological data. Cladistics 20: 518-533. Savigny, J.-C. 1822. Systèmes des annelides, principalement de celles des côtes de l’Égypte et de la Syrie. Pp. 128. In M. J. L. Savigny (ed.), Description de l’Égypte, Histoire Naturelle, vol. 1(3), Paris. Schulze, A. 2003. Phylogeny of Vestimentifera (Siboglinidae, Annelida) inferred from morphology. Zoologica Scripta 32: 321–342. Seaver, E. C. 2003. Segmentation: mono- or polyphyletic? International Journal of Developmental Biology 47: 583-595. Sedgwick, A. 1898. A student’s textbook of zoology. 1., Swan Sonnenschein & Co. Ltd., London.

Annelid Phylogeny and Systematics



Siddall, M. E., Apakupakul, K., Burreson, E. M., Coates, K. A., Erseus, C., Gelder, S. R., Kallersjo, M. and Trapido-Rosenthal, H. 2001. Validating Livanow: Molecular data agree that leeches, branchiobdellidans, and Acanthobdella peledina form a monophyletic group of oligochaetes. Molecular Phylogenetics & Evolution 21: 346-351. Stechmann, A. and Schlegel, M. 1999. Analysis of the complete mitochondrial DNA sequence of the brachiopod Terebratulina retusa places Brachiopoda within the protostomes. Proceedings of the Royal Society of London, Series B 266: 2043-2052. Struck, T., Hessling, R. and Purschke, G. 2002. The phylogenetic position of the Aeolosomatidae and Parergodrilidae, two enigmatic oligochaete-like taxa of the ‘Polychaeta’, based on molecular data from 18S rDNA sequences. Journal of Zoological Systematics & Evolutionary Research 40: 155-163. Swedmark, B. 1952. Note préliminaire sur un polychète sédentaire aberrant, Psammodrilus balanoglossoides, n.gen., n.sp. Arkiv för Zoologi 4: 159-162. Westheide, W. 1997. The direction of evolution within the Polychaeta. Journal of Natural History 31: 1-15. Westheide, W., McHugh, D., Purschke, G. and Rouse, G. W. 1999. Systematization of the Annelida: different approaches. Hydrobiologia 402: 291-307. Winnepenninckx, B., Backeljau, T. and De Wachter, R. 1995. Phylogeny of protostome worms derived from 18S r RNA sequences. Molecular Biology and Evolution 12: 641-649.

CHAPTER

2

Oogenesis Kevin J. Eckelbarger

2.1 INTRODUCTION Annelid eggs have served as models in pioneering 19th and 20th century investigations of oocyte organelle function, fertilization, and early embryological development (reviewed in Eckelbarger 1988), including those of polychaetes (reviewed in Eckelbarger 1988) and clitellates (reviewed in Dohle 1999). Over the last 100 years, studies of invertebrate oogenesis have involved most of the major phyla with polychaetous annelids receiving disproportionate attention. This may be due to the fact that polychaetes are such common members of the marine benthos and that they show unusual reproductive plasticity (Hermans and Schroeder 1975; Wilson 1991; Giangrande 1997). Polychaete oogenesis has been extensively reviewed by Schroeder and Hermans (1975), Olive (1983), and Eckelbarger (1983, 1984, 1986, 1988, 1992, 2005). Jamieson (1981, 1988, 1992) reviewed oogenesis in oligochaetes, while Fernandez et al. (1992) reviewed the process in leeches. Lasserre (1975) summarized the reproductive biology of clitellates, including gonad morphology and some aspects of oogenesis.

2.2 OVARIAN MORPHOLOGY AND PATTERNS OF OOGENESIS 2.2.1 Introduction In order to comprehend the evolutionary forces that have molded annelid life history patterns, one must examine the role played by the ovary and the various mechanisms of vitellogenesis that have arisen through selection. The ovary and the associated vitellogenic mechanisms employed during oogenesis play a direct role in the rate of egg production, the frequency of breeding, and the size and energy content of the egg and related consequences for larval dispersal ability. Oogenesis is best understood in the context of the general reproductive biology of the species. While the evolution of sperm morphology is highly correlated with fertilization Darling Marine Center and School of Marine Sciences, The University of Maine, 193 Clark’s Cove Road, Walpole, Maine 04573, USA

" Reproductive Biology and Phylogeny of Annelida biology, oogenesis appears to be correlated with habitat specialization and general life history pattern (Eckelbarger 1994). In annelids, one sees considerable variation in ovarian complexity, pattern of oogenesis, and life history strategy, all of which are interrelated. Polychaetes. The majority of polychaetes are gonochoric although many examples of hermaphroditism have been documented (reviewed by Schroeder and Hermans 1975). The diversity of reproductive features exhibited within some families can only be described as extraordinary (see Wilson 1991), for example, Syllidae (Franke 1999), Spionidae (Blake and Arnofsky 1999), Maldanidae (Rouse 1992), and the Cirratulidae (Petersen 1999). Polychaete life histories encompass semelparity, annual iteroparity, and continuous iteroparity (Olive 1984), but their ovaries show wide structural variation so patterns of oogenesis cannot be generalized. The majority of polychaetes have well-defined ovaries that often persist throughout their ontogeny. Westheide and Purschke (in Westheide et al. 1999) proposed that the basic annelid body plan includes gonads in all segments. Some polychaete families reflect this model, having paired, retroperitoneal ovaries that may be repeated in a large number of segments, as seen, for example, in Aphroditidae, Capitellidae, Glyceridae, Syllidae, Tomopteridae, Nephtyidae, Onuphidae, Orbiniidae, Polynoidae, Serpulidae, and Sabellariidae. Equally common, however, are ovaries that are fused and/or restricted to a few segments, as observed in Pectinariidae, Arenicolidae, Terebellidae, Ampharetidae, Maldanidae, Opheliidae, Sabellidae, and Serpulidae (partially reviewed in Clark and Olive 1973). In Cirratulidae and Spionidae, there are examples of species having both restricted and multiple genital segments. Segmentally repeated ovaries are most common when complete segmentation is present. The absence or reduction of intersegmental septa usually results in the restriction in number of ovarian segments. Small-bodied species that produce only a few eggs at a time show a marked restriction in the number of genital segments. Intrafamilial variation is rarely observed in terms of ovarian structure and distribution, although in serpulids some species lack ovaries, while in others they are well-defined (Kupriyanova et al. 2001). The great majority of polychaetes have well-defined and relatively permanent ovaries, although none has been observed in the Nereididae (Olive 1983), Alciopidae (Eckelbarger and Rice 1988), Sphaerodoridae (Christie 1984), and Phyllodocidae (Olive 1975). At least six different types of polychaete ovaries have been documented (reviewed in Eckelbarger 1988). Ovaries are often located in the parapodia or ventral-lateral regions of the body where they may be closely associated with parapodial connective tissue, intersegmental septa, the ventral peritoneum, or more commonly, elements of the circulatory system, and especially with nephridial blood vessels. Clitellata. Oligochaetes are hermaphroditic, they usually possess permanent ovaries in restricted segments, they have complex reproductive systems, and they undergo mutual cross-fertilization followed by zygote

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encapsulation, resulting in direct development. Their breeding patterns are highly variable even within species so it is impossible to generalize (Brinkhurst and Jamieson 1971; see also Chapter 8 of this volume). Leeches are protandrous hermaphrodites with annual reproductive cycles, distinct ovaries in a fixed position, and they undergo cross fertilization via copulation or the use of spermatophores introduced via hypodermic impregnation (Mann 1962; Lasserre 1975).

2.2.2 Patterns of Oogenesis The size and structural complexity of annelid ovaries varies according to the pattern of oogenesis employed (reviewed in Eckelbarger 1988) (see Fig. 2.1). In polychaetes, two patterns have been defined, including extraovarian and intraovarian oogenesis (Eckelbarger 1983). Extraovarian oogenesis occurs when small, previtellogenic oocytes are released from the ovary where they complete vitellogenesis in the fluid-filled coelom. In these species, the ovaries are generally small, proliferative organs that are structurally simple and transient in nature (Fig. 2.1A, B). In many instances, it is common for these species to store abundant nutrient reserves in the somatic tissues of the female prior to vitellogenesis (Eckelbarger 1983). Two types of extraovarian oogenesis have been identified in polychaetes. In its simplest form, small, previtellogenic oocytes are released from the ovary and enter the coelom where they undergo solitary differentiation as freefloating cells (e.g. Sabellidae, Glyceridae, Serpulidae, Oweniidae) (Fig. 2.1D, E). In other families (e.g. Alciopidae, Nereididae, Phyllodocidae, Pholoidae, Sphaerodoridae, Terebellidae, Cirratulidae, Ampharetidae, Pectinariidae), clusters of previtellogenic oocytes are released into the coelom while enveloped in follicle cells (Fig. 2.1C). In Nereididae (Fischer 1975) and Alciopidae (Eckelbarger and Rice 1988), oocytes within these clusters are also connected by intercellular bridges. The oocyte clusters eventually lose their follicular envelope and the oocytes separate and undergo vitellogenesis while floating solitarily in the coelomic fluid. However, in Sphaerodoridae (Christie 1984) and Pholoidae (Heffernan and Keegan 1988), the oocyte clusters remain intact throughout most of vitellogenesis. In a few species, nurse cells are connected to developing oocytes by cytoplasmic intercellular bridges, but these examples are relatively rare (Fig 2.1H). Nurse cell-oocyte associations have been described in some interstitial species (reviewed by Eckelbarger 1992), in a number of syllids (Cognetti-Varriale 1965; Heacox and Schroeder 1981), in the tomopterid Tomopteris helgolandica (Åkesson 1962), in several species within the Onuphidae (Anderson and Huebner 1968; Paxton 1979; Hsieh 1984; Eckelbarger 1988, 1992), in the dorvelleids Ophryotrocha spp. (Korschelt 1893; Ruthmann 1964; Emanuelsson 1969; Pfannenstiel 1978), and in the maldanid Micromaldane nutricula (Rouse 1992). Intraovarian oogenesis occurs when oocytes are retained by the ovary until most or all of oogenesis (and vitellogenesis) is completed (Fig. 2.1F, G). In these species, ovaries are usually larger, more structurally complex, and

$ Reproductive Biology and Phylogeny of Annelida

Fig. 2.1 A. Transverse section of Nicolea zostericola showing the ventral ovary above the nerve cord and between two mucous glands. CO, coelom. B. Early germ cells in the ovary of Euratella sp. projecting into the coelom. C. Oocytes floating in the coelom of Rynchonerella angelini, including packets of oogonia surrounded by follicle cells (arrow), clusters of larger, previtellogenic oocytes, and solitary vitellogenic oocytes. D. Free-floating, solitary oocytes in various stages of oogenesis in the coelom of Terebella rubra. EVO, early vitellogenic oocyte. E. Solitary vitellogenic oocytes in the same stage of oogenesis in the coelom of Pseudoeurythoe sp.. F. The ovary of Methanoaricia Fig. 2.1 Contd. ...

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persistent throughout the sexual phase of the female. There is no consistency among intraovarian species with respect to ovarian morphology, although most ovaries are spherical in form and oocytes always develop in intimate contact with follicle cells. The polychaete families that have been studied are nearly equally divided between those that undergo extraovarian or intraovarian oogenesis. Clitellata. Clitellates show notable differences from polychaetes in having ovaries restricted to a few segments and in being entirely hermaphroditic (Jamieson 1992). The plesiomorphic condition for clitellate ovaries consists of paired structures restricted to one or two segments (Jamieson 1992; Fernández et al. 1992), and all species undergo intraovarian oogenesis (Jamieson 1988, 1992; Fernández et al. 1992). The ovaries of oligochaetes are small, sac-like organs posterior to the testes that project into the segmental coelom and are covered by a thin layer of peritoneum (Anderson 1971; Jamieson 1981). Oocytes are not released into the ovisac (a diverticulum of the coelom) until they have completed vitellogenesis and are metaphase primary oocytes (Jamieson 1992). The ovary is associated with blood vessels in a manner similar to some polychaetes (Type 6, Eckelbarger 1988), but the oocytes do not directly contact the blood vessel lumen. Dumont (1969) described packets of eight synchronously developing oocytes connected by intercellular bridges in Enchytraeus albidus. In Eisenia fetida, groups of oogonia or the premeiotic primary oocytes are linked by intercellular bridges to a central cytophore and each develops from one oogonium (Jamieson 1988, 1992). Cytoplasmic bridges have been reported in oocyte clusters of the polychaete Cirriformia sp. (Eckelbarger 1988), but this requires confirmation since it is the only known example in polychaetes. The ovaries of leeches are paired, rounded or elongated organs enclosed within fluid-filled coelomic cavities and they may grow to extend through several post-clitellar segments (Fernández et al. 1992) (Fig. 2.2). They are comprised of an outer region or ovisac, and one or more solid cords of cells (ovary cord) that include the germinal epithelium. The ovaries lie in a hemocoel and the ovary lumen is also a hemocoelic cavity. Division of each germinal cell within the ovary cord produces an oogonium and a follicle cell. The oogonium divides to produce a cluster of clonal, isogenic cells, or polyplast, that remains enveloped by descendants of the follicle cell. The oogonial polyplasts are comprised of pear-shaped cells that communicate with a central, anuclear cytophore. Further differentiation Fig. 2.1 Contd. ...

dendrobranchiata, showing oocytes attached to blood vessels arising from the body wall. G. Transverse section through Capitella jonesi showing paired ovaries suspended in the coelom below the gut and attached to blood vessels. H. Single oocyte with attached nurse cells in the coelom of Tomopteris pacifica. Abbreviations: BC, blood cell; BV, blood vessel; BW, body wall; CO, coelom; EVO, early vitellogenic oocyte; GT, gut; GV, germinal vesicle; MG, mucous glands; NC, nurse cell; OC, oocyte; OV, ovary; PVO, previtellogenic oocytes; VO, vitellogenic oocytes. Original.

& Reproductive Biology and Phylogeny of Annelida

Fig. 2.2 Piscicola geometra. Oocyte development within a follicle. A. Synaptonemal complex in the nucleus of a young oocyte. B. Intermediate cell with microvilli (arrows) surrounding the oocyte-nurse cell complex beneath. C. Ovarian follicle showing nurse cells connected by intercellular bridges to a single oocyte. Abbreviations: NC, nurse cells; OC, oocyte; SC, synaptonemal complex. From Fischer, A. and Weigelt, K.-R. 1975. Sonderdruck aus Verhandlungsbericht der Deutschen Zoologischen Gesellschaft 67: 319-323, Fig. 1.

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results in the formation of a larger, single oocyte while the remaining cells become nurse cells and only the oocyte undergoes meiosis. Mature polychaete eggs range in size from 20–40 µm in Polyophthalmus pictus (Fage and Legendre 1927) to 1.17 mm in diameter in Paronuphis antarctica (Hartman 1967). The size of clitellate eggs show similar variability ranging from only 30 µm in diameter in the megadrilid Eisenia fetida (Lechenault 1968) to 1 mm diameter in Glossiphonia spp. (Anderson 1973).

2.3 OOGENESIS: PREVITELLOGENESIS The cytological events occurring during this phase of oogenesis are very similar in both polychaetes and clitellates and mirror those described in most metazoans (Huebner and Anderson 1976; Wourms 1987; Eckelbarger 1994). Oocytes have a disproportionately large nucleus with one or more nucleoli, and ooplasm containing little more than a few, small mitochondria (Fig. 2.3). Oocytes in the zygotene/pachytene stage of the meiotic prophase lack nucleoli and have morphologically distinct synaptonemal complexes representing synapsed chromosomes. Following this premeiotic phase, the nucleolus and diffuse chromatin reappear and the nucleus greatly enlarges to form a spherical germinal vesicle. The nucleus undergoes numerous morphological changes during this phase, including the appearance of a single prominent nucleolus during the diplotene stage, and its later division into several satellite nucleoli that assume positions close to the inner nuclear envelope. Nucleocytoplasmic granules migrate through the nuclear pores into the perinuclear ooplasm where they form fibrogranular clusters called nuage (Fig. 2.4). Nuage is often closely associated with pleomorphic mitochondria and can assume a wide variety of distinct morphological forms, particularly in polychaetes (Eckelbarger 1988). During the late previtellogenic phase, Golgi complexes proliferate throughout the ooplasm and rough endoplasmic reticulum (RER) appears in various forms including parallel and whirled arrays. Depending on the species, microvilli usually proliferate along the surface of the oolemma during this stage in polychaetes (Eckelbarger 1984), oligochaetes (Jamieson 1992), and leeches (Fischer and Weigelt 1975; Fernández et al. 1992). This process has been more extensively studied in polychaetes where the microvilli show wide morphological variability and complexity (Eckelbarger 1992, 2005).

2.4 OOGENESIS: VITELLOGENESIS The vitellogenic phase of oogenesis usually occurs during the diplotene stage of first meiotic prophase and results in a rapid increase in cell volume due to the production and assembly of nutrients, including glycogen, lipid droplets, and chemically complex, membrane-bounded yolk bodies or platelets (reviewed in Eckelbarger 1986). Numerous studies have been published on polychaete vitellogenesis, while only a few are available for

! Reproductive Biology and Phylogeny of Annelida

Fig. 2.3 A. Previtellogenic oocytes in the ovary of Capitella sp. III surrounded by follicle cells. B. Microvilli projecting from the oocyte surface in Capitella sp. III. C. Previtellogenic oocyte of Capitella jonesi surrounded by follicle cells. Abbreviations: FC, follicle cells; MV, microvilli; Nu, nucleolus; OC, vitellogenic oocyte; PVO, previtellogenic oocytes. Original.

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Fig. 2.4 A. Perinuclear region of early vitellogenic oocyte of Capitella jonesi showing fibergranular material (arrows) passing through the nuclear pores. B. Higher magnification of perinuclear region of oocyte of Capitella jonesi showing perinuclear (nuage) material (*) in association with mitochondria. C. Nuage (*) and clustered mitochondria in oocyte of Phyllodoce fragilis. D. Nucleolus (*) in previtellogenic oocyte of Polydora ligni. E. Nuage (*) in the perinuclear ooplasm in an oocyte of Diopatra cuprea. Abbreviations: M, mitochondria; N, nucleus;Nu, nucleolus; Y, yolk. Original.

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Reproductive Biology and Phylogeny of Annelida

clitellates. Based on the diverse studies of polychaetes, most annelids are likely to employ a combination of two proteosynthetic processes during vitellogenesis, similar to those described throughout the Metazoa (reviewed by Wourms, 1987; Eckelbarger 1994). Autosynthesis involves the manufacture of yolk via oocyte organelles (e.g. Golgi complex, RER), following the incorporation of low molecular weight extraoocytic precursors, while heterosynthesis involves the incorporation of large molecular weight, female-specific yolk proteins (vitellogenin) from extraovarian sources for assembly in the ooplasm. Vitellogenic processes have been extensively investigated on the biochemical level in nereidid polychaetes, providing the most detailed information available on yolk biosynthesis in annelids (reviewed in Eckelbarger 1988; Fischer and Hoeger 1993; Fischer et al. 1996). Polychaetes. Vitellogenic oocytes typically contain numerous Golgi complexes and extensive RER, both of which proliferate during early vitellogenesis (Fig. 2.5). Both organelles appear to cooperate in the synthesis of yolk bodies that assume a wide variety of morphological forms (Fig. 2.6). These proteosynthetic organelles appear to be ubiquitous in the majority of polychaete oocytes (pers. obs.), although their presence alone cannot definitively establish the vitellogenic mechanism. Extensive studies of nereidid vitellogenesis have demonstrated that the mere presence of Golgi complexes and RER does not exclude the possibility of heterosynthetic processes (reviewed in Eckelbarger 1992). However, morphological studies of the oocytes of Dinophilus ciliatus (Grün 1972) and Harmothoe imbricata (Garwood 1981) suggest that yolk production occurs exclusively within the RER cisternae. Sichel (1966) hypothesized that yolk was derived largely through the transformation of mitochondria in Mercierella enigmatica oocytes. In a few studies, ultrastructural evidence suggests that yolk synthesis occurs from the incorporation of large molecular weight yolk proteins via receptor-mediated endocytosis along the surface of the oocyte. In Phragmatopoma lapidosa (Eckelbarger 1979), Streblospio benedicti (Eckelbarger 1980), Spio setosa (Eckelbarger unpublished), and Leitoscoloplos fragilis (Eckelbarger unpublished), direct uptake of yolk proteins form the circulatory system occurs in oocytes attached to blood vessel walls (Fig. 2.7). Vitellogenesis in S. benedicti is unique in metazoans in that hemoglobin is inexplicably incorporated into developing yolk bodies from the circulatory system during vitellogenesis (Eckelbarger 1980). Polychaete oocytes are associated with follicle cells at some point during oogenesis and, in species undergoing intraovarian oogenesis, they maintain intimate contact until they are released from the ovary (Fig. 2.8). Many roles have been proposed for follicle cells (Eckelbarger 1992) but they are based largely on morphological studies. A significant biosynthetic function has been hypothesized for follicle cells in Kefersteinia cirrata (now known as Psamathe fusca Johnston, 1836) (Olive and Pillai 1983), Phragmatopoma lapidosa (Eckelbarger 1979), Streblospio benedicti (Eckelbarger 1980), and Capitella jonesi (Eckelbarger and Grassle 1982), where they

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Fig. 2.5 A. Rough endoplasmic reticulum in the perinuclear ooplasm of Streblospio benedicti. N, nucleus. B. Parallel arrays of RER in the ooplasm of Capitella jonesi. C. Rough endoplasmic reticulum cisternum and Golgi complex in perivitelline region of Streblospio benedicti oocyte. D. Golgi complexes in oocyte of Vanadis formosa. E. Golgi complexes closely associated with newly formed yolk body in oocyte of Capitella jonesi. Abbreviations: EE, egg envelop; G, Golgi complex; M, mitochondria; N, nucleus; Nu, nucleolus; RER, Rough endoplasmic reticulum; Y, yolk body. Original.

!" Reproductive Biology and Phylogeny of Annelida

Fig. 2.6 A. Cortical region of vitellogenic oocyte of Capitella jonesi showing yolk bodies and associated Golgi complex. Arrows indicate endocytotic pits along oolemma. B. Yolk bodies in oocyte of Diopatra cuprea. Yolk bodies from the oocytes of Fabricia sabella, C, Myzostoma sp., D, Phyllodoce fragilis, E, Phragmatopoma lapidosa, F, Polydora ligni, G, and Amphisamytha galapagensis, H. Abbreviations: G, Golgi complex; M, mitochondria; MV, microvilli; Y, yolk bodies. Original.

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Fig. 2.7 A. Intraovarian, early vitellogeneic oocytes attached to the genital blood vessel in Lamellibrachia sp. B. Cortical region of oocyte of Capitella sp. I showing endocytotic pits (arrows). C. Perivasal region of vitellogenic oocyte of Streblospio benedicti showing endocytotic pits (arrows) and endosomes (*). D. Perivasal region of vitellogenic oocyte of Streblospio benedicti showing endocytotic pits (arrow) and endosomes (*) migrating into ooplasm. Abbreviations: BV, blood vessel; OC, vitellogeneic oocytes; P, peritoneal cell; Y, yolk body. Original.

!$ Reproductive Biology and Phylogeny of Annelida

Fig. 2.8 A. Vitellogenic oocyte of Capitella jonesi with overlying layer of follicle cells. B. Follicle cell with prominent nucleus covering the surface of a vitellogenic oocyte of Capitella jonesi. C. Follicle cell from the ovary of Capitella sp. III showing Golgi complexes and RER. D. Follicle cell from ovary of Capitella sp. I containing parallel arrays of RER. E. Ovarian follicle cells (FC) containing whirled arrays of RER. Abbreviations: CO, coelom; FC, follicle cells; EE, egg envelop; G, Golgi complexes; M, mitochondrion; N, nucleus; Nu, nucleolus; OC, vitellogenic oocyte; Y, yolk body. Original.

Oogenesis

!%

undergo hypertrophy before and during vitellogenesis and exhibit significant biosynthetic activity. In those few species having nurse cells associated with oocytes, there have been no documented examples of direct nurse cell involvement in vitellogenesis so their role in oogenesis is presently unknown. Clitellata. Oligochaete vitellogenesis involves autosynthesis as well as heterosynthetic uptake of yolk precursors via endocytosis or incorporation of yolk from chloragocytes (Dumont 1969; Bondi and Facchini 1972; Jamieson 1981). Autosynthetic yolk formation has been described in Eisenia foetida involving the formation of lipid globules through the transformation of mitochondria, and via an ooplasmic lamellar or membrane system (Lechenault 1968). In Enchytraeus sp., early vitellogenic, third-stage oogonia have an abundance of vesicles of possible Golgi origin that coalesce to form nascent yolk bodies (Jamieson 1992), presumably via autosynthetic pathways. The oocytes of Branchiobdella pentodonta contain abundant Golgi complexes and RER resulting in the formation of prominent yolk platelets with a cortical crystalline substructure (Bondi and Facchini 1972). Heterosynthetic yolk formation is suggested by coated pits in stage III oogonia and incorporation of horseradish peroxidase tracer in the matrix of yolk spheres (Dumont 1969). Oligochaetes lack nurse cells but follicle cells are common and appear to play a supportive rather than a nutritive roll due to their lack of proteosynthetic activity (Jamieson, 1981, 1988). In leeches, a single oocyte within each polyplast communicates with nurse cells via intercellular bridges to a central cytophore in early and midclones (Fernández et al. 1992). Cytoplasmic communication between nurse cells and oocytes allows the transfer of large molecules and organelles from the former to the latter, suggesting a supportive role for nurse cells during vitellogenesis. The follicle cell envelope surrounding the oocyte-nurse cell complex appears to be involved in nutrient transport to the oocyte surface from the lumen of the ovisac. Ultrastructural studies have demonstrated endocytotic uptake of proteins by the oocyte, apparently leading to the deposition of yolk in the cortical region of the ooplasm where abundant Golgi complexes and RER are observed. Thus a combination of autosynthetic and heterosynthetic processes probably participate in vitellogenesis.

2.5 PHYLOGENETIC IMPLICATIONS OF OOGENESIS Sperm morphology and elements of spermatogenesis have provided fruitful data for phylogenetic assessments of clitellates (Jamieson et al. 1987; Ferraguti and Erséus 1999) and polychaetes (Jamieson and Rouse 1989; Rouse 1999). Rouse and Fitzhugh (1994) and Blake and Arnofsky (1999) demonstrated the value of incorporating other reproductive and developmental data in phylogenetic analyses with their studies of the Sabellidae and Spionidae, respectively. As noted by Nielsen (1998), oogenesis has attracted little attention in metazoan phylogenetic studies.

!& Reproductive Biology and Phylogeny of Annelida Theoretically, comparative investigations of oogenesis in annelids (or any group) have the potential of assisting us in assessing phylogenetic relationships as similar studies have done in vertebrate phylogeny (Jones 1978). However, unlike vertebrates, invertebrates show far greater diversity in ovarian morphology and patterns of oogenesis, and our knowledge of most groups is very fragmentary in comparison (reviewed in Eckelbarger 1994). Ovarian morphology and patterns of oogenesis in annelids are undoubtedly strongly correlated with life history strategies, and there are doubts about their usefulness in phylogenetic analyses due to sparse data (Eckelbarger 1986, 1988; Jamieson 1988; Rouse 1992). In stark contrast to sperm, the female gamete shows such striking morphological similarities throughout Metazoa that even the eggs of some invertebrates and vertebrates can appear virtually indistinguishable. In addition, while most invertebrate phyla are conservative with respect to the number, location, and structural complexity of their ovaries, annelids are not—primarily due to the high degree of lability within polychaetes (Eckelbarger 1992, 1994). Fauchald and Rouse (1997) have noted that the external morphological features of annelids, especially polychaetes, have been extensively investigated while studies of internal structures are far more limited, especially on the ultrastructural level. This observation applies to annelid ovaries and oogenesis because existing information is too limited to be very useful in addressing questions of homology. For example, comprehensive studies of ovarian structure and oogenesis in polychaetes are limited to only 0.3% of described species (Giangrande 1997), and nearly all of these are based on common shallow water species. Far fewer studies have been conducted on clitellates. In contrast to testes and sperm, ovaries and eggs have fewer morphological characters that could prove useful in phylogenetic analyses. However, there are a few characters worthy of consideration, including: 1) the presence or absence of definable ovaries; 2) the number and location of ovaries; 3) the existence of extraovarian vs. intraovarian oogenesis; 4) the release of previtellogenic oocytes into the coelom as solitary cells or in clusters; 5) the presence or absence of nurse cells, 6) the egg envelop morphology, and 7) the yolk platelet structure. Clitellates are notable in having clearly defined ovaries restricted to a few segments and in exhibiting hermaphroditism. In contrast, the majority of polychaetes have indistinct ovaries, they show wide variation in ovarian position, and they are generally dioecious. Intraovarian oogenesis is a plesiomorphic feature of the clitellates but is observed in about half the polychaete families studied with no obvious phylogenetic pattern. A few polychaete families have both intraovarian and extraovarian oogenesis and this variability is likely to be related to differences in life histories (Eckelbarger 1983). It is worth noting that within Phyllodocida, Nereididae, Alciopidae, Sphaerodoridae, Phyllodocidae, and Pholoidae, all share one unique feature with respect to oogenesis that may indicate a close relationship: they all lack a defined ovary and ovulation results in the

Oogenesis

!'

release of previtellogenic oocytes into the coelom in clusters surrounded by follicle cells. Fauchald and Rouse (1997) suggested that the Alciopidae were likely to have evolved from benthic ancestors resembling phyllodocids, and Rouse and Pleijel (2001) recently referred to them as Alciopini and members of the Phyllodocidae. The fact that both groups lack defined ovaries and possess oocyte clusters offers additional support. Nurse cells are derived only from the germ cell line (Huebner and Anderson 1976), and they appear to have multiple functions throughout the Metazoa (reviewed in Eckelbarger 1994). While the association of nurse cells with oocytes characterizes leeches, they are not encountered in oligochaetes and are relatively rare in polychaetes. However, nurse cells are common in many species of the Onuphidae (Paxton 1979) and Eunicidae (Anderson and Huebner 1968) and may be, in the opinion of Rouse (1992), an apomorphic character state for the Eunicida. While annelid nurse cells may be homologous, no common function has been established. Egg envelope morphology often shows show strong intrafamilial similarities in polychaetes (Eckelbarger 1984, 1988, 1992), but morphological variation has also been documented even within sibling species complexes (Eckelbarger and Grassle 1983). On the other hand, Blake and Arnofsky (1999) documented three distinct types of egg envelopes in spioniform polychaetes that are restricted to specific clades, demonstrating their value in phylogenetic analyses. Yolk body morphology often appears unique and consistent within some polychaete families (Eckelbarger, 2005), but there is significant variability in others. In addition, yolk bodies are biochemically complex structures (Eckelbarger 1986) that have been poorly studied and there is no evidence that they are homologous organelles. Indeed, yolk platelet morphology has even been documented to vary between closely related sibling species (Eckelbarger and Grassle 1983).

2.6 CONCLUSIONS Annelids show great diversity with respect to ovarian complexity, vitellogenic mechanisms, and reproductive patterns, which may reflect the life history specializations that have evolved in the group. The ovary and associated vitellogenic mechanisms play a pivotal role in the rate of egg production, the frequency of breeding, and the size and energy content of the egg and resultant consequences for larval development and dispersal. Efforts to comprehend the evolutionary forces that have molded annelid life history patterns must include the role played by the ovary and the diverse mechanisms of yolk synthesis that have arisen through selection. Unfortunately, our knowledge of annelid ovarian structure and oogenesis is very limited so it is difficult to correlate these features with life history evolution. In addition, it is difficult, at present, to apply what we know to phylogenetic analyses. In order to incorporate features of ovaries and oogenesis into these studies, additional research is required, particularly involving annelids with diverse reproductive and developmental patterns.

" Reproductive Biology and Phylogeny of Annelida

2.7 LITERATURE CITED Åkesson, B. 1962. The embryology of Tomopteris helgolandica (Polychaeta). Acta Zoologica 43: 135-199. Anderson, D. T. 1971. Embryology. Pp. 73-103. In R. O. Brinkhurst and B. G. M. Jamieson, Aquatic Oligochaetes of the World. Oliver and Boyd, Edinburgh. Anderson, D. T. 1973. Embryology and Phylogeny in Annelids and Arthropods. Pergamon Press, New York. 495 pp. Anderson, E. and Huebner, E. 1968. Development of the oocyte and its accessory cells in the polychaete, Diopatra cuprea (Bosc). Journal of Morphology 126: 163172. Blake, J. A. and Arnofsky, P. L. 1999. Reproduction and larval development of the spioniform Polychaeta with application to systematics and phylogeny. Hydrobiologia 402: 57-106. Bondi, C. and Facchini, L. 1972. Observations on the oocyte ultrastructure and vitellogenesis of Branchiobdella pentodonta Whitman. Acta Embryologiae Experimentalis 2: 225-241. Brinkhurst, R. O. and Jamieson, B. G. M. 1971. Aquatic Oligochaeta of the World. Oliver and Boyd, Edinburgh. 860 pp. Christie, G. 1984. The reproductive biology of a Northumberland population of Sphaerodorum gracilis (Rathke, 1843) (Polychaeta: Sphaerodoridae). Sarsia 69: 117121. Clark, R. B. and Olive, P. J. W. 1973. Recent advances in polychaete endocrinology and reproductive biology. Annual Review of Oceanography and Marine Biology 11: 175-222. Cognetti-Varriale, A. M. 1965. Richerche sulla biologia riproductiva dei Policheti: I. Gli ovari Exogoninae. Archivio Zoologico Italiano 50: 26-28. Dohle, W. 1999. The ancestral cleavage pattern of the clitellates and its phylogenetic deviations. Hydrobiologia 402: 267-283. Dumont, J. N. 1969. Oogenesis in the annelid Enchytraeus albidus with special reference to the origin and cytochemistry of yolk. Journal of Morphology 129: 317-344. Eckelbarger, K. J. 1979. Ultrastructural evidence for both autosynthetic and heterosynthetic yolk formation in the oocytes of an annelid (Phragmatopoma lapidosa: Polychaeta). Tissue and Cell 11: 425-443. Eckelbarger, K. J. 1980. An ultrastructural study of oogenesis in Streblospio benedicti (Spionidae), with remarks on diversity of vitellogenic mechanisms in Polychaeta. Zoomorphology 94: 241-263. Eckelbarger, K. J. 1983. Evolutionary radiation in polychaete ovaries and vitellogenic mechanisms and their role in life history patterns. Canadian Journal of Zoology 61: 487-504. Eckelbarger, K. J. 1984. Comparative aspects of oogenesis in polychaetes. In A. Fischer, and H. -D. Pfannenstiel (eds), Polychaete Reproduction. Progress in Comparative Reproductive Biology, Fortschritte der Zoologie 29: 123-148. Eckelbarger, K. J. 1986. Vitellogenic mechanisms and the allocation of energy to offspring in polychaetes. Bulletin of Marine Science 39: 426-443. Eckelbarger K. J. 1988. Oogenesis and female gametes. In W. Westheide and C. O. Hermans (eds), The Ultrastructure of Polychaeta, Microfauna Marina 4: 281-307. Eckelbarger, K. J. 1992. Oogenesis. Pp. 109-127. In F. W. Harrison and S. L. Gardiner (eds), Microscopic Anatomy of Invertebrates. Vol. 7, Chapter 2 (Polychaeta). WileyLiss Inc., New York.

Oogenesis

"

Eckelbarger, K. J. 1994. Diversity of metazoan ovaries and vitellogenic mechanisms: implications for life history theory. Proceedings of the Biological Society of Washington 107: 193-218. Eckelbarger, K. J. 2005. Oogenesis and oocytes. Hydrobiologia 535/536: 179-198. Eckelbarger, K. J. and Grassle, J. P. 1982. Ultrastructure of the ovary and oogenesis in the polychaete Capitella jonesi (Hartman, 1959). Journal of Morphology 171: 305320. Eckelbarger, K. J. and Grassle, J. P. 1983. Ultrastructural differences in the eggs and ovarian follicle cells of Capitella (Polychaeta) sibling species. Biological Bulletin 165: 379-393. Eckelbarger, K. J. and Grassle, J. P. 1984. Role of ovarian follicle cells in vitellogenesis and oocyte resorption in Capitella sp. I (Polychaeta). Marine Biology 79: 133-144. Eckelbarger, K. J. and Rice, S. A. 1988. Ultrastructure of oogenesis in the holopelagic polychaetes Rhynchonerella angelini and Alciopa reynaudii (Polychaeta: Alciopidae). Marine Biology 98: 427-439. Emanuelsson, H. 1969. Electron microscope observations on yolk and yolk formation in Ophryotrocha labronica La Greca and Bacci. Zeitschrift für Zellforschung und Mikroskopische Anatomie 953: 19-36. Fage, L. and Legendre, R. 1927. Pêches planctonique à la lumière, effectuées à Banyuls-sur-Mer et à Concarneau. I. Annelides Polychètes. Archives de Zoologie Experimentale Générale 67: 23-222. Fauchald, K. and Rouse, G. 1997. Polychaete systematics: Past and present. Zoologica Scripta 26: 71-138. Ferraguti, M. and Erséus, C. 1999. Sperm types and their use for a phylogenetic analysis of aquatic clitellates. Hydrobiologia 402: 225-237. Fernández, J., Téllez, V. and Olea, N. 1992. Hirudinea. Pp. 323-394. In F. W. Harrison and S. L. Gardiner (eds), Microscopic Anatomy of Invertebrates, Vol. 7, Annelida. Wiley-Liss, New York. Fischer, A. and Hoeger, U. 1993. Metabolic links between somatic sexual maturation and oogenesis in nereid annelids—a brief review. Invertebrate Reproduction and Development 23: 131-138. Fischer, A. and Weigelt, K.-R. 1975. Strukturelle Beziehungen zwischen jungen Oocyten und somatischen Zellen bei den Anneliden Platynereis und Piscicola. Sonderdruck aus Verhandlungsbericht der Deutschen Zoologischen Gesellschaft. 67: 319-323. Fischer A, Dorresteijn, A. W. C., and Hoeger, U. 1996. Metabolism of oocyte construction and the generation of histospecificity in the cleaving egg. Lessons from nereid annelids. International Journal of Developmental Biology 40: 421430. Franke, H. -D. 1999. Reproduction of the Syllidae (Annelida: Polychaeta). Hydrobiologia 402: 39-55. Garwood, P. R. 1981. Observations on the cytology of the developing female germ cell in the polychaete Harmothoe imbricata (L.). International Journal of Invertebrate Reproduction 3: 333-345. Giangrande, A. 1997. Polychaete reproductive patterns, life cycles and life histories: an overview. Annual Review of Oceanography and Marine Biology 35: 323-386. Grün, G. 1972. Über den Eidimorphismus und die Oogenese von Dinophilus gyrociliatus (Archinnelida) Zeitschrift für Zellforschung und Mikroskopische Anatomie 130: 70-92. Hagedorn, H. H. and Kunkel, J. G. 1979. Vitellogenin and vitellin in insects. Annual Review of Entomology 24: 475-505.

"

Reproductive Biology and Phylogeny of Annelida

Hartman, O. 1967. Polychaetous annelids collected by the USNS Eltanin and Saten Island cruises, chiefly from Antarctic Seas. Allan Hancock Monographs on Marine Biology 2: 1-387. Heacox, A. E. and Schroeder, P. C. 1981. A light and electron microscopic investigation of gametogenesis in Typosyllis pulchra (Berkeley and Berkeley) (Polychatea: Syllidae). II. Oogenesis. Cell and Tissue Research 218: 641-658. Heffernan, P. and Keegan, B. F. 1988. Quantitative and ultrastructural studies on the reproductive biology of the polychaete Pholoe minuta in Galway Bay. Marine Biology 99: 203-214. Hermans, C. O. and Schroeder, P. C. 1975. Annelida: Polychaeta. Pp. 1-213. In A. C. Giese and J. S. Pearse (eds), Reproduction of Marine Invertebrates, Vol. III, Annelids and Echiurans. Academic Press, New York. Hsieh, H. -L. 1984. Morphological studies on the reproduction and larval development of Kinbergonuphis simoni (Polychaeta: Onuphidae). M.Sc. thesis, University of South Florida, Tampa. Huebner, E. and Anderson, E. 1976. Comparative spiralian oogenesis—structural aspects; an overview. American Zoologist 16: 315-343. Hutchings, P. A. 1973. Gametogenesis in a Northumberland population of the polychaete Melinna cristata. Marine Biology 18: 199-211. Jamieson, B. G. M. 1981. The Ultrastructure of the Oligochaeta. Academic Press, London and New York. 462 pp. Jamieson, B. G. M. 1988. Oligochaete ultrastructure: Some comparisons with the Polychaeta. Pp. 397-428. In W. Westheide and C. O. Hermans (eds), The Ultrastructure of the Polychaeta. Gustav Fischer Verlag, New York. Jamieson, B. G. M. 1992. Oligochaeta. Pp. 217-322. In F.W. Harrison and S.L. Gardiner (eds), Microscopic Anatomy of Invertebrates. Wiley-Liss, New York. Jamieson, B. G. M. and Rouse, G. W. 1989. The spermatozoa of the Polychaeta (Annelida). An ultrastructural review. Biological Reviews 64: 93-157. Jamieson, B. G. M., Erséus, C. and Ferraguti, M. 1987. Parsimony analysis of the phylogeny of some Oligochaeta (Annelida) using spermatozoal ultrastructure. Cladistics. 3: 145-155. Jones, R. E. (ed.). 1978. The Vertebrate Ovary. Comparative Biology and Evolution. Plenum Press, New York. 853 pp. Jouin, C. 1968. Sexualité et biologie de la réproduction chez Mesonerilla Boaden (Archiannélides Nerillidae). Cahiers de Biologie Marine 9: 31-52. Korschelt, E. 1893. Über Ophryotrocha puerilis Clap. —Metsch. Und die polytrochen Larven eines anderen Anneliden (Harpochaeta cingulata, nov. gen. nov. spec.). Zeitschrift wiss Zoologie 57: 224-289. Kupriyanova, E. K., Nishi, E., Ten Hove, H. A. and Rzhavsky, A. V. 2001. Life history patterns in serpulimorph polychaetes: ecological and evolutionary perspectives. Annual Review of Oceanography and Marine Biology 39: 1-101. Lasserre, P. 1975. Clitellata. Pp. 215-275. In A. C. Giese and J. S. Pearse (eds), Reproduction of Marine Invertebrates, Vol. III, Annelids and Echiurans. Academic Press, New York. Lechenault, H. 1968. Etude cytochimique et ultrastructurale de l’ovocyte d’Eisenia foetida (Sav.). Zeitschrift für Zellforschung. 90: 96-112. Mann, K. H. 1962. Leeches (Hirudinea). Their Structure, Physiology, Ecology and Embryology. Pergamon Press, New York. 201 pp. Nielsen, C. 1998. Morphological approaches to phylogeny. American Zoologist 38: 942-952.

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"!

Olive, P. J. W. 1975. Reproductive biology of Eulalia viridis (Müller) (Polychaeta: Phyllodocidae) in the northeastern U.K. Journal of the Marine Biology Association of the United Kingdom 55: 313-326. Olive, P. J. W. 1983. Oogenesis in Annelida: Polychaeta. Pp. 357-422. In K. G. Adiyodi, and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates, Vol. 1, Oogenesis, Oviposition, and Oosorption, Wiley, New York. Olive, P. J. W. 1984. Environmental control of reproduction in polychaeta. Fortschritte der Zoologie. 29: 17-38. Olive, P. J. W. and Pillai, G. 1983. Reproductive biology of the polychaete Kefersteinia cirrata Keferstein (Hesionidae). II. The gametogenic cycle and evidence for photoperiodic control of oogenesis. International Journal of Invertebrate Reproduction 6: 307-315. Paxton, H. 1979. Taxonomy and aspects of the life history of Australian beachworms (Polychaeta: Onuphidae). Australian Journal of Marine and Freshwater Research 30: 265-294. Petersen, M. E. 1999. Reproduction and development in Cirratulidae (Annelida: Polychaeta). Hydrobiologia 402: 107-128. Pfannenstiel, H. D. 1978. Die Entwicklung der Kontakstruktur von Ei und Nahrzelle im Zuge der Oogenese von Ophryotrocha puerilis Claparède and Mecznikow (Polychaeta, Dorvilleidae). Zoomorphologie 90: 181-196. Rouse, G. W. 1992. Oogenesis and larval development in Micromaldane spp. (Polychaeta: Capitellida: Maldanidae). Invertebrate Reproduction and Development 21: 215-230. Rouse, G. W. 1999. Polychaete sperm: phylogenetic and functional considerations. Hydrobiologia 402: 215-224. Rouse, G. W. and Fitzhugh, K. 1994. Broadcasting fables: is external fertilization really primitive? Sex, size and larvae in sabellid polychaetes. Zoologica Scripta 23: 271-312. Ruthmann, A. 1964. Zellwachstum und RNS-Synthese im Ei-Nahrzellverband von Ophryotrocha puerilis. Zeitscrift der Zellforschungen 63: 816-829. Schroeder, P. C. and Hermans, C.O. 1975. Annelida: Polychaeta. Pp. 1-213. In A. C. Giese and J. S. Pearse (eds), Reproduction of Marine Invertebrates, Vol. III, Academic Press, New York. Sichel, G. 1966. Modificazioni ultrastructurali dell ooplasma in rapporta alla vitellogenese in Mercierella enigmatica Fauvel (Annelida: Polychaeta). Atti dell’ Accademia Gioenia di Scienze Naturali 18: 21-23. Westheide, W., McHugh, D., Purschke, G. and Rouse, G. 1999. Systematization of the Annelida: different approaches. Hydrobiologia 402: 291-307. Wilson, W. H. 1991. Sexual reproductive modes in polychaetes: classification and diversity. Bulletin of Marine Science 48: 500-516. Wourms, J. P. 1987. Oogenesis. Pp. 49-178. In A. C. Giese and J. S. Pearse (eds), Reproduction of Marine Invertebrates, Vol. 9, General Aspects: Seeking Unity in Diversity. Boxwood Press, Pacific Grove.

CHAPTER

3

Annelid Sperm and Spermiogenesis Greg W. Rouse

3.1 INTRODUCTION Ultrastructural studies of spermiogenesis and the morphology spermatozoa in annelids have been reviewed several times (Ferraguti 1983, 1999; Ferraguti and Erséus 1999; Franzén and Rice 1988; Jamieson and Rouse 1989; Rice 1992; Rouse 1999a). Here an introduction to the concepts and literature is given as well as an update on the more recent findings. All currently available ultrastructural descriptions of sperm ultrastructure in non-clitellate annelids are provided in Table 3.1. For information on studies on sperm in Clitellata see Chapter 8 where sperm of oligochaetes is comprehensively reviewed and Chapter 9 for information on Hirudinida.

3.2 SPERMIOGENESIS Most annelids lack permanent gonads. The origin and proliferation of the primordial germ cells are poorly understood, and this also applies to early stages of gamete development. The process of gamete proliferation most commonly attributed to annelids concerns liberation of spermatogonia (that divide mitotically to proliferate) or spermatocytes (that divide meiotically) into the coelom from germ cells lining the peritoneum. These patches of germ cells are generally found ventrally associated with the coelom lining near the ventral blood vessel and in males may be referred to as testes. Spermatogenesis is the process that commences with the primordial germ cells and ends with spermatozoa. Spermatogonia may or not divide or may divide numerous times to become clusters of spermaocytes. Spermatocytes undergo a reduction division to give rise to spermatids. The latter stage of the spermatogenesis, whereby the spermatids differentiate without division to produce spermatozoa, is distinguished as spermiogenesis (Olive 1983). The ‘testes’ of annelids usually contain only South Australian Museum Nth Terrace, Adelaide. S.A. 5000 Australia & Earth and Environmental Sciences, University of Adelaide SA. 5005 Australia

"$ Reproductive Biology and Phylogeny of Annelida Table 3.1 Sperm studies on Annelida with some ultrastructural component

Less inclusive taxa Clitellata

Arenicolidae Arenicola brasiliensis A. marina

Scolecida

Aciculata Amphinomida

Aciculata Eunicida

Capitellidae Capitella capitata Capitella spp. Capitellides spp. Capitomastus spp. Maldanidae Clymenella sp. Micromaldane spp. Cossuridae Cossura cf. longocirrata Opheliidae Armandia sp. Orbiniidae Haploscoloplos elongatus Naineris laevigata

Methanoaricia dendrobranchiata Questidae Questa ersei Scalibregmatidae Travisia japonica Amphinomidae Eurythoe complanata Diurodrilus D. subterraneus

Dorvilleidae Apodotrocha progenerans Dinophilus caudatus Ophryotrocha diadema O. gracilis O. hartmanni O. labronica O. notoglandulata O. puerilis Trilobodrilus spp.

Sperm

Reference

Intro

(Ferraguti 1999; Ferraguti and Erséus 1999). See text, Chapters 8 and 9

Ent Ent

(Sawada 1975) (Bentley and Pacey 1989; Meijer 1979; Pacey and Bentley 1992)

Intro Intro Intro Intro

(Franzén 1982a) (Eckelbarger and Grassle 1987) (Eckelbarger and Grassle 1987) (Eckelbarger and Grassle 1987)

Ent? Ent

(Rouse and Jamieson 1987) (Rouse 1992a)

Ect?

(Rouse and Tzetlin 1997)

Ect?

(Jamieson and Rouse 1989)

? Ent? Ent

(Rice 1992) (Giangrande and Petraroli 1994b) (Eckelbarger and Young 2002)

Intro

(Jamieson 1983a, 1983b)

Ect?

(Kubo and Sawada 1977)

Ect

(Rouse and Jamieson 1987)

Intro

(Kristensen and EibyeJacobsen 1995; Kristensen and Niilonen 1982) (Westheide 1988a) (Franzén 1977b) (Pfannesteil and Grünig (Pfannesteil and Grünig (Pfannesteil and Grünig (Berruti et al. 1978) (Pfannesteil and Grünig (Pfannesteil and Grünig (Scharnofske 1986)

1990) 1990) 1990) 1990) 1990)

Table 3.1 Contd. ...

Annelid Sperm and Spermiogenesis

"%

Table 3.1 Contd. ...

Aciculata Eunicida

Aciculata Phyllodocida

Eunicidae Marphysa mullawa Histriobdellidae Histriobdella homari Stratiodrilus novaehollandiae Lumbrineridae Lumbrineris sp. Onuphidae Diopatra sp. Hyalinoecia tubicola

Kinbergonuphis simoni Onuphis mariahirsuta Aphroditidae Laetmonice producta Chrysopetalidae Chrysopetalum debile Chrysopetalum sp. Dysponetus caecus Dysponetus pygmaeus Hesionides H. arenaria Hesionidae capricornia Lizardia hirschi Sirsoe methanicola Microphthalmus M. carolensis M. listensis M. nahantensis Myzostomida Myzostoma cirriferum Myzostoma sp. Nephtyidae Nephtys sp. Nereididae Neanthes japonica Nereis diversicolor N. irrorata N. limbata N. pelagica N. virens Playnereis brevicirrus P. dumerilii P. massiliensis Tylorrhynchus heterochaetus

Ect?

(Jamieson and Rouse 1989)

Int Int

(Jamieson et al. 1985) (Jamieson et al. 1985)

Ect?

(Rouse 1988b) (Jamieson and Rouse 1989) (Cotelli and Lora Lamia Donin 1975) (Hsieh and Simon 1990) (Jamieson and Rouse 1989) (Micaletto et al. 2003)

et et et et

al. al. al. al.

Ect? Ect? Intro? Intro?

(Tzetlin (Tzetlin (Tzetlin (Tzetlin

2002) 2002) 2002) 2002)

Intro

(Westheide 1984b)

Intro? Intro? Ect?

(Pleijel and Rouse 2000) (Pleijel and Rouse 2005) (Eckelbarger et al. 2001)

Intro Intro Intro

(Westheide and Rieger 1987) (Westheide 1984a) (Westheide and Rieger 1987)

Intro Intro

(Afzelius 1983, 1984; Eeckhaut and Jangoux 1991) (Mattei and Marchand 1988)

Ect?

(Rouse 1999a) Chapter 10

Ect Ect Ect Ect Ect Ect Ect Ect Ent Ect

(Sato and Osanai 1986) (Bertout 1976) (Defretin and Wissocq 1974) (Fallon and Austin 1967) (Defretin and Wissocq 1974) (Bass and Brafield 1972) (Kubo and Sawada 1977) (Pfannenstiel et al. 1987) (Lücht and Pfannensteil 1989) (Sato and Osanai 1983, 1990) Table 3.1 Contd. ...

"& Reproductive Biology and Phylogeny of Annelida Table 3.1 Contd. ...

Aciculata incertae sedis

Pholoidae Laubierpholoe swedmarki Pholoe minuta Taylorpholoe sp. Phyllodocidae Eulalia sp. Alciopa reynaudii Alciopina parasitica Krohnia lepidota Naiades cantrainii Plotohelmis tenuis Rhynchonerella angelini R. moebii Torrea candida Vanadis formosa Pilargidae Sigambra sp. Pisionidae Pisione remota Polynoidae Alentia gelatinsa Arctonoe spp. Harmothoe imbricata H. impar Lepidonotus sp. Sigalionidae Sigalion bandaensis Psammolyce sp. Syllidae Autolytus sp. Calamyzas amphictenicola Exogone dispar E. naidina Grubeosyllis clavata Petitia amphophthalma Sphaerosyllis hermaphrodita Typosyllis sp. Typosyllis pulchra Tomopteridae Tomopteris helgolandica Nerillidae Nerilla antennata

Canalipalpata Cirratuliformia

Acrocirridae Acrocirrus validus Macrochaeta clavicornis

Aciculata Phyllodocida

Int Ect?

Chapter 10 (Heffernan and Keegan 1988) (Rouse 1999a) Chapter 10

Ect? Ent Ent Ent Ent Ent Ent Ent Ent Ent

(Rouse 1988b) (Rice 1987) (Rice 1987) (Rice 1987) (Rice 1987) (Rice 1987) (Rice 1987) (Rice 1987) (Rice 1987) (Rice 1987)

Ect?

Chapter 10

Intro

(Westheide 1988b)

Ect Ent Ent Ect?

(Franzén and Rice 1988) (Pernet 2000) (Bentley and Serries 1992) (Bentley and Serries 1992) (Rouse 1988b) Chapter 10

Ect? Ect?

(Jamieson and Rouse 1989) (Rouse 1999a) Chapter 10

Ent? Ent? Ent Ent Ent? Intro Intro Ect? Ect?

(Franzén 1982a) (Franzén and Rice 1988) (Giangrande et al. 2002) (Giangrande et al. 2002) (Franzén 1974) (Buhrmann et al. 1996) (Kuper and Westheide 1997a, 1997b) (Jamieson and Rouse 1989) (Heacox and Schroeder 1981)

Ent?

(Franzén 1982b)

Intro

(Franzén and Sensenbaugh 1984)

Ect? Ect?

(Sawada 1984) see Chapter 11 Table 3.1 Contd. ...

Annelid Sperm and Spermiogenesis

"'

Table 3.1 Contd. ...

Canalipalpata Cirratuliformia

Canalipalpata Terebelliformia

Canalipalpata Spionida

Canalipalpata Sabellida

Cirratulidae Cirriformia tentaculata Cirriformia sp. Ctenodrilus sp. Flabelligeridae Flabelligera sp. Alvinellidae Alvinella caudata Alvinella pompejana Paralvinella grasslei P. pandorae pandorae P. pandorae irlandei P. palmiformis Ampharetidae Amphisamytha galapagensis Pectinariidae Cistenides okudai Terebellidae Nicolea zostericola

Ramex californica Streblosoma acymatum Chaetopteridae Chaetopterus pergamentaceus C. ‘variopedatus’ Mesochaetopterus minutus Magelonidae Magelona sp. Spionidae Boccardiella hamata Marenzelleria viridis Polydora ciliata P. ligni P. neoceaca P. socialis P. websteri Prionospio fallax P. cf. queenslandica Pseudopolydora paucibranchiata Streblospio benedicti Tripolydora sp. Oweniidae Owenia ‘fusiformis’ Sabellariidae Idanthyrsus pennatus

Ect? Intro

(Sawada 1984) (Jamieson and Rouse 1989) (Rouse 1999a) see Chapter 11

Ect?

(Rouse 1999a)

Ent Ent Ent Ent Ent Ent

(Jouin-Toulmond (Jouin-Toulmond (Zal et al. 1994) (McHugh 1995) (Jouin-Toulmond (Jouin-Toulmond

Ect?

(McHugh and Tunnicliffe 1994)

Ect?

(Sawada 1984)

Ent Ent Ect?

(Eckelbarger 1974; Rouse and McHugh 1994) (Rouse and McHugh 1994) (Jamieson and Rouse 1989)

Ect?

(Anderson and Eckberg 1983)

Ect? Ect?

(Jamieson and Rouse 1989) (Sawada 1984)

Ect?

(Rouse 1999a)

? Ect? Intro Intro Intro Intro Intro Ect? Ect?

(Rice 1992) (Bochert 1996) (Franzén 1974) (Rice 1981) (Williams 2000) (Rice 1981) (Rice 1981) (Franzén and Rice 1988) (Rouse 1988a)

? Intro Intro

(Rice 1992) (Rice 1981) (Rouse 1988a)

Ect

(Rouse 1988b)

Ect

This chapter

et al. 2002) et al. 1997)

et al. 2002) et al. 2002)

Table 3.1 Contd. ...

# Reproductive Biology and Phylogeny of Annelida Table 3.1 Contd. ...

Canalipalpata Sabellida

Phragmatopoma lapidosa P. californica Sabellaria alveolata S. cementarium Sabellidae Sabellinae Amphicorina bicoloris A. brevicollaris A. dentata A. mobilis A. paramobilis Amphiglena lindae A. mediterranea A. nathae A. pacifica A. terebro Bispira melanostigmata B. volutacornis Branchiomma bombyx B. luctuosum

Ect Ect Ect Ect

(Eckelbarger 1984) (Kopp 1985) (Pasteels 1965) (Franzén and Rice 1988)

Ent Ent Ent? Ent Ent Ent Ent Ent Ent Ent Ect? Ect? Ect? Ent

B. nigromaculata Demonax polarsterni Euchone pallida Jasmineira sp. Myxicola cf. sulcata Notaulax nudicollis Perkinsiana antarctica P. borsibrunoi P. littoralis P. riwo P. rubra Pseudopotamilla reniformis Sabella pavonina S. spallanzanii

Ect? Ect? Ect? Ect? Ent Ect? Ent Ect? Ect Ent Ect? Ect? Ect? Ect

Terebrasabella heterouncinata Sabellidae Fabriciinae Augeneriella basifurcata A. hummelincki A. pectinata Genus A. sp. Florida Genus A. sp. PNG Fabricia stellaris

Ent

(Rouse 1992b) (Rouse 1992b) (Rouse 1992b) (Rouse 1992b) (Rouse 1999a) (Rouse and Gambi 1998) (Rouse and Gambi 1998) (Rouse and Gambi 1998) (Rouse and Gambi 1998) (Rouse 1993a) (Rouse 1999a) (Nash and Keegan 2003) (Franzén and Rice 1988) (Licciano et al. 2002; Sordino and Gambi 1994) (Rouse 1999a) (Gambi et al. 2001) (Gambi et al. 2001) (Rouse 1999a) (Gambi et al. 2001) (Rouse 1999a) (Gambi and Patti 1999) (Gambi et al. 2000) (Gambi et al. 2000) (Rouse 1996b) (Chughtai 1986) (Chughtai 1986) (Graebner and Kryvi 1973) (Giangrande et al. 2000; Giangrande and Petraroli 1994a) (Fitzhugh and Rouse 1999; Simon and Rouse 2005)

Ent Ent Ent Ent Ent Ent

(Rouse (Rouse (Rouse (Rouse (Rouse (Rouse

1995) 1995) 1995) 1995) 1995) 1995) Table 3.1 Contd. ...

Annelid Sperm and Spermiogenesis

#

Table 3.1 Contd. ...

Canalipalpata Sabellida

Fabriciola brevibranchiata F. cri F. flammula F. liguronis F. mediaseta F. minuta F. parvus Fabricinuda bikinii F. trilobata Manayunkia aestuarina M. baicalensis M. mizu Novafabricia brunnea N. infratorquata N. tenuiseta Pseudofabricia aberrans Pseudofabriciola incisura P. quasiincisura P. peduncula Parafabricia ventricingulata Serpulidae Serpulinae Chitinopoma serrula Galeolaria caespitosa Hydroides dirampha Hydroides elegans Hydroides hexagonus Paraprotis dendrova

Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent Ent

(Rouse 1995) (Rouse 1995, 1996a) (Rouse 1993b) (Rouse 1993b) (Rouse 1995) (Rouse 1996c) (Rouse 1993b) (Rouse 1995) (Rouse 1995) (Rouse 1995) (Pylilo and Vishnjakov 1993) (Rouse 1995) (Rouse 1995) (Rouse 1995) (Rouse 1995) (Rouse 1995) (Rouse 1995) (Rouse 1995) (Rouse 1995) (Rouse 1992c)

Ent Ect? Ect? Ect? Ect? Ent

Pomatoleios krausii Serpula sp. Spirobranchus giganteus Serpulidae Filograninae Salmacina sp. Serpulidae Spirorbinae Metalaeospira tenuis Neodexiospira sp. Pileolaria potswaldi (as Spirorbis morchi) Pileolaria sp. Paralaeospira cf. levinsoni Protolaeospira tricostalis Protolaeospira capensis Romanchella quadricostalis Spirorbis spirorbis

Ect? Ect? Ect?

(Franzén 1982a) (Rouse 2005) This chapter (Mona et al. 1994) (Nishi 1992) (Colwin and Colwin 1961) (Rouse 1999b; Rouse 2005) This chapter (Nishi 1992; Sawada 1984) (Jamieson and Rouse 1989) (Nishi 1992)

Ent

(Rouse 1996c; Rouse 2005)

Ent Ent

(Rouse 2005) (Rouse 2005)

Ent Ent Ent Ent Ent Ent Ent

(Potswald 1967) (Rouse 2005) (Rouse 2005) (Rouse 2005) (Rouse 2005) (Rouse 2005) (Daly and Golding 1977; Picard 1980; Rouse 2005) Table 3.1 Contd. ...

#

Reproductive Biology and Phylogeny of Annelida

Table 3.1 Contd. ...

Canalipalpata Sabellida

Canalipalpata incertae sedis

Annelida incertae sedis

Siboglinidae Osedax rubiplumus Ridgeia piscesae Riftia pachyptila Siboglinum ekmani Polygordiidae Polygordius lacteus Protodrilida Parenterodrilus teanioides

Intro Ent Ent Intro

(Rouse et al. 2004) (Southward and Coates 1989) (Jones and Gardiner 1985) (Franzén 1973)

Ect?

(Franzén 1977b)

Intro

Protodriloides symbioticus Protodrilus adhaerens P. ciliatus P. gracilis P. haurakiensis P. helgolandicus P. hypoleucus P. jagersteni P. jouinae P. litoralis P.oculifer P. purpureus P. rubropharyngeous

Intro Intro Intro Intro Intro Intro Intro Intro Intro Intro Intro Intro Intro

P. submersus Saccocirrus sp. Echiura Bonellia viridis Hamingia arctica Urechis caupo Aeolosomatidae and Potamodrilus Aeolosoma litorale Aeolosoma marcusi Aeolosoma singulare Potamodrilus fluviatilus Hrabeiella H. periglandulata Parergodrilidae Parergodrilus heideri

Intro Intro

(Jouin-Toulmond and Purschke 2004) (Jouin 1978) (Nordheim von 1989) (Nordheim von 1989) (Nordheim von 1989) (Nordheim von 1989) (Nordheim von 1989) (Nordheim von 1989) (Nordheim von 1989) (Nordheim von 1989) (Nordheim von 1989) (Nordheim von 1989) (Nordheim von 1989) (Franzén 1974; Nordheim von 1989) (Nordheim von 1989) (Jamieson and Rouse 1989)

Intro Intro Ect

(Franzén and Ferraguti 1992) (Franzén and Ferraguti 1992) (Cross 1984)

Ent? Ent? Ent? Ent?

(Bunke 1986) (Gluzman 1994, 1997) (Marotta et al. 2003b) (Bunke 1985)

Intro

(Rota and Lupetti 1997)

Intro

(Purschke 2002a)

spermatogonia and stem cells (Olive 1983). Examples of groups with such simple testes are most Ampharetidae, Arenicolidae, Capitellidae, Clitellata, Opheliidae, Serpulidae, Syllidae and Terebellidae (Olive 1983), thus spanning the phylogenetic diversity of Annelida. Later stages of spermatogenesis within testes may occur in polychaetes such as Pectinariidae, Polynoidae and Flabelligeridae (Olive 1983) and clitellates such as lumbricids (see chapter 8). However, no case of fully differentiated

Annelid Sperm and Spermiogenesis

#!

sperm being found in testes has been documented (Olive 1983). In exceptional cases, such as some Phyllodocida and Echiura, it is thought there are no fixed testes and that the germ cell float freely in the coelom (Gould-Somero 1975; Olive 1983). So, in annelids, it can be generalized that the process of sperm maturation mainly occurs floating freely in the coelomic fluid (Olive 1983; Schroeder and Hermans 1975). In most annelids the sperm develop in syncytial masses that have been referred to as rosettes, morulae, or platelets, and these terms are often used interchangeably for bundles of spermatids or spermatocytes (Schroeder and Hermans 1975; Olive 1983). These clusters of spermatids are the result of the spermatogonia diving a number of times and staying attached and then as spermatocytes undergoing the reduction division resulting in a quadrupling of the number of cells attached to the syncytium. These clusters of spermatids in a syncytium can range from 8 to hundreds. On the other hand, in some polychaetes, a solitary spermatocyte undergoes the reduction division resulting in four spermatids. The two basic forms of spermiogenesis in annelids are illustrated here in Fig. 3.1. Figures 3.1A, B show spermatids developing in fabriciin Sabellidae. All members of this clade exhibit spermiogenesis where the spermatids develop in a large cluster attached to a central cytophore (Fig. 3.1A). The number of spermatids in a group varies according to the species. The spermatids usually develop with the acrosome closest to the cytophore and the tail and/ or midpiece furthest away (Fig. 3.1B). This situation is found in virtually all annelids that have a cytophores, except aeolosomatids and Potomodrilus (see below). The less common form of spermiogensis found in polychaetes, sperm developing tetrads with no central cytophore (Fig. 3.1B) is also found in Sabellida. This form of development is commonly (though not universally) seen in the sabellid subfamily Sabellinae and in many Serpulidae and has been used in as a character in cladistic analyses (Rouse and Fitzhugh 1994; Fitzhugh and Rouse 1999).

3.3 SPERMATOZOA 3.3.1 Sperm Terminology Annelids have a great range of sperm morphologies and traditionally these have been grouped as either ‘primitive sperm’ or ‘modified sperm’ based on the terminology of Retzius (1904; 1905) and Franzén (1956). Polychaetes show a range of sperm shapes (Fig. 3.2) but are traditionally referred to by these two terms only, whilst all clitellates would be classified as having modified sperm under this terminology. Franzén (1956) suggested that ‘primitive’ sperm with heads comprised of a simple acrosome, spherical nuclei and a small number of mitochondria and a free flagellum were associated with external fertilization (Fig. 3.2A). Modified sperm were any sperm that deviate from this pattern (e.g., Fig. 3.2B). Jamieson (1986)

#" Reproductive Biology and Phylogeny of Annelida

Fig. 3.1. Annelid spermatids. A. Interference contrast micrograph of spermatid bundle of Augeneriella alata (Sabellidae). Note central cytophore. B. Longitudinal section through spermatid bundle of Manayunkia aestuarina (Sabellidae). C. Tetrad of spermatids of Salmacina sp. (Serpulidae). Original.

Annelid Sperm and Spermiogenesis

##

introduced the neutral term ‘aquasperm’ and Rouse and Jamieson (1987) subsequently proposed a new system of classifying sperm based purely on function. Ect-aquasperm are released into the water and fertilize similarly released eggs. This term places no phylogenetic significance on this form of fertilization (e.g., Fig. 3.2A). Ent-aquasperm are again released freely into the ambient water but differ from ect-aquasperm in being gathered by, or in some other way reaching, the female (e.g., Fig. 3.2B). Introsperm have no contact with water when passed from male to female. This terminology was designed to avoid any a priori judgment of phylogenetic pattern. Additionally, a number polychaetes (and other animals e.g., see Rouse and Pitt 2000) have been found to have sperm with so-called ‘primitive’ morphology, yet do not fertilize freely spawned eggs in the classical sense. So among annelids, so-called ‘primitive’ sperm may be found in spermatophores as in some spionids (Richards 1970), or stored by

Fig. 3.2. A. Ect-aquasperm of Eurythoe complanata (Amphinomidae). The sperm is about to meet the egg. B. Ent-aquasperm of Manayunkia aestuarina. The sperm are released by the males and are taken up by the females and stored in spermathecae before fertilizing the eggs. Original. Abbreviations a, acrosome; e egg surface; m mitochondria; n nucleus.

#$ Reproductive Biology and Phylogeny of Annelida the female in spermathecae before fertilization e.g., alciopids (Rice and Eckelbarger 1989). In the echiurans Bonellai viridis and Hamingia arctica, where dwarf males live inside females and fertilization is internal, the sperm differ from classical ‘primitive sperm only in the fact that they have long acrosomes (Franzén and Ferraguti 1992). Rouse and Fitzhugh (1994) found after a cladistic analysis of some Sabellida that external fertilization and sperm with ‘primitive’ morphology were secondarily evolved in Sabellidae. The preceding examples make it quite clear that the term ‘primitive’ sperm fails to indicate plesiomorphic status for this sperm shape and also fails to be a good guide as to fertilization mechanism. Similarly the term ‘modified’ sperm is a catchall for sperm of many different shapes and only suggests that they have a spherical head. The terms ‘ect-aquasperm’, ‘ent-aquasperm’ and ‘introsperm’ are now being applied across various taxa in addition to polychaetes (Degaulejac et al. 1995; Hodgson and Chia 1993; Rouse and Pitt 2000; Sousa and Oliveira 1994) though this has not met with universal acceptance. For instance, Rice (1992) argued that it was difficult and potentially misleading to apply these terms and used the terms ‘primitive’ and ‘modified’ in describing polychaete sperm. Since the terms of Rouse and Jamieson (1987) require knowledge of reproductive mechanisms in the species concerned before the sperm can be classified it can be difficult to classify the sperm. However, it is certainly not a misleading terminology. Given the observation by Franzén (1956) that fertilization mechanisms do tend to correlate with sperm morphology (given the caveats outlined above), it is possible to tentatively assign sperm to a category, but as pointed out above, sperm shape is not an infallible guide to reproductive mode and this really needs to be used with caution.

3.3.2 Recent Descriptions of Annelid Sperm Ultrastructure Since the last review on the sperm of Clitellata (Ferraguti 1999; Ferraguti and Erséus 1999), there have been a number of further publications on clitellate sperm (Ferraguti et al. 1999; Ferraguti et al. 2002; Gelder and Ferraguti 2001; Marotta et al. 2003a). These papers have provided further evidence of the utility of sperm based characters for the phylogenetic analysis of clitellates initially demonstrated by Jamieson 1981, 1983c, 1984; Jamieson et al. 1987 (see also Chapter 8 of this volume). Since the last review of polychaete sperm by Rouse (1999a), there have been a number of interesting descriptions of polychaete sperm and fertilization mechanisms. These are studies were on Aeolosomatidae (Marotta et al. 2003b), Alvinellidae (Jouin-Toulmond et al. 2002), Aphroditidae (Micaletto et al. 2003), Chrysopetalidae (Tzetlin et al. 2002), Hesionidae (Eckelbarger et al. 2001; Pleijel and Rouse 2000, 2005), Orbiniidae (Eckelbarger and Young 2002), Parergodrilidae (Purschke 2002a), Protodrilida (Jouin-Toulmond and Purschke 2004), Sabellidae (Fitzhugh and Rouse 1999; Gambi et al. 2000; Gambi et al. 2001; Giangrande et al. 2000; Licciano et al. 2002; Nash and Keegan 2003; Simon and Rouse 2005), Siboglinidae (Hilario et al. 2005; Rouse et al. 2004), Spionidae (Williams 2000) and Syllidae (Giangrande et al. 2002).

Annelid Sperm and Spermiogenesis

#%

Rouse (1999a) did not discuss the description of the sperm of Hrabeiella periglandulata by Rota and Lupetti (1997). This taxon was referred to as a polychaete by Rouse and Pleijel (2001) with its placement among this assemblage listed as incertae sedis. Hrabeiella periglandulata is a terrestrial organism, but is clearly not a clitellate. Apart from the lack of a clitellum it also lacks the various reproductive features such as seminal receptacles or spermathecae (Rota and Lupetti 1997). It has been proposed however that H. periglandulata may be the sister group to Clitellata (Purschke 2003). Purschke (2003) based his argument on a series of proposed homologies based on the form of pharynx, cerebral sense organs and the central nervous system. The sperm of H. periglandulata, while filiform, shows no particular similarities with clitellates such as the presence of an acrosome tube, or interpolation of the mitochondria between the axonome and the nucleus (Rota and Lupetti 1997). Rota and Lupetti (1997) rejected any affinity with clitellates on this basis and compared the sperm with that of a number of polychaetes but found none that was particularly similar. However, this result does not actually rule out a close, or even sister group relationship, between Hrabeiella and Clitellata as proposed by Purschke (2003). The two sperm apomorphies above (and also a central sheath and tetragon fibres in the axonome) (see Ferraguti 1984) clearly support monophyly of Clitellata. That the sperm of H. periglandulata lacks these does not invalidate a close relationship with clitellates and Purschke’s (2003) hypothesis deserves close attention. A study of molecular sequence data for 18S rDNA, 28S rDNA and COI sequences failed to show a close relationship between clitellates and Hrabeiella (Jördens et al. 2004) and so the search for the sister group for Clitellata continues (see Chapter 1). Purschke (2002a) described the sperm of another enigmatic terrestrial annelid, Parergodrilus heideri. This taxon is placed, along with Stygocapitella subterranea in Parergodrilidae, and is currently regarded as incertae sedis among polychaetes by Rouse and Pleijel (2001). Parergodrilus heideri has internal fertilization, yet the sperm presents another example where sperm shape is misleading with respect to fertilization mechanism. The sperm of P. heideri is an almost classical ‘primitive’ sperm according to the terminology of Franzén (1977a), but it is found in a terrestrial polychaete with internal fertilization, and so can hardly be regarded as a primitive annelid. While the sperm acrosome, nucleus and midpiece are very slightly elongate, the sperm is, for instance, quite similar to sperm of many broadcast spawning polychaetes. The example of P. heideri provides yet more evidence for the lack of utility for the terms ‘primitive’ and ‘modified’ sperm.

3.4 SPERM OF SOME UNUSUAL ANNELIDS 3.4.1 Diurodrilus A detailed description of spermiogenesis and sperm ultrastructure of Diurodrilus subterraneus by Kristensen and Eibye-Jacobsen (1995) revealed

#& Reproductive Biology and Phylogeny of Annelida the sperm of this species to be quite unusual (Fig. 3.3A) as is the animal itself. The nucleus is small and capped by an extremely large acrosome. Several small mitochondria surround the nucleus anteriorly and the tail that is a free flagellum is attached at the base of the nucleus. Diurodrilus was originally referred to Dinophilidae by Remane (1925), with Diurodrilidae erected by Kristensen and Niilonen (1982). However, the latter author’s view that Diurodrilus and Dinophilidae are separate taxa does not preclude that Diurodrilus (and Dinophilidae) is part of Dorvilleidae. Eibye-Jacobsen and Kristensen (1994) did not consider Diurodrilus in their Dorvilleidae phylogenetic analysis and in a following study on sperm ultrastructure in Diurodrilus (Kristensen and Eibye-Jacobsen 1995) they considered the group to be of uncertain affinity. The sperm exhibits (at least) superficial similarities to Apodotrocha, even though detailed examination of the latter is lacking and in some Diurodrilus a posterior copulatory organ appears to be present (Kristensen and Eibye-Jacobsen 1995) that may be homologous to that in Dinophilus. In view of the current inclusions of such taxa as Dinophilus and Apodotrocha, Rouse and Pleijel (2001) included Diurodrilus in Dorvilleidae, although further studies are certainly required.

3.4.2 Aeolosomatidae and Potamodrilus Studies on sperm and spermiogenesis by Bunke (1985, 1986) on aeolosomatid and Potamodrilus sperm have shown that neither group has the clitellate apomorphies of an acrosome tube or the interpolatation of mitochondria between the centrioles and the nucleus. This led Bunke to conclude that these taxa should not be included in Clitellata (see Chapter 1). Spermiogenesis in both Aeolosoma litorale and Potamodrilus fluviatilis involves the spermatids developing synchronously in large groups (128 in the case of A. litorale) attached to cytophores (Bunke 1985, 1986). In both species the sperm show an unusual mode of development in that the acrosome develops furthest away from the cytophore and the flagellum closest to the cytophore. As seen in many other annelids, microtubules from a ring around the nucleus during spermiogenesis. The morphology of the sperm in both is essentially filiform (Fig. 3.3C, F). In P. fluviatilis no true midpiece is present and a few mitochondria can be found around the basal region of the nucleus (Fig. 3.3B, C). In A. litorale a small midpiece is formed but the mitochondria surround the anchoring apparatus and no part of the axoneme. In both species a number of unusual vesicles can be seen around the basal region of the nucleus (Fig. 3.3B, G). Spermiogenesis and sperm structure of the A. litorale and P. fluviatilis show no real indicators that would help to place these taxa within Annelida.

3.4.3 Siboglinidae As outlined in Chapter 1, Pogonophora (including Vestimentifera) have now been reclassified as members of the polychaete clade Sabellida (Rouse and Fauchald 1997), under the name Siboglinidae.

Annelid Sperm and Spermiogenesis

#'

Fig. 3.3. Sperm of Diurodrilus, Aeolosomatidae and Potamodrilidae. A. Diurodrilus subterraneus. Drawing of a mature sperm from a male specimen. Note very large acrosome in relation to the nucleus). B. Potamodrilus fluviatilis. Drawing of the base of the middle region of the sperm showing mitochondria along nucleus, unusual vesicles and the anchoring apparatus for the tail. C. Potamodrilus fluviatilis. Diagrammatic drawing of mature sperm showing characteristic hairpin bend at the base of the nucleus. D. Potamodrilus fluviatilis. Drawing of the base of the acrosome and the tip of the nucleus. E. Aeolosoma litorale. Drawing of the acrosome. F: Aeolosoma litorale. Schematic drawing of mature sperm. G. Aeolosoma litorale. Drawing of midpiece of spermatozoon. Note extremely short mitochondrial ring around the anchoring apparatus. A, modified from Kristensen, R. M. and EibyeJacobsen, D. 1995. Zoomorphology 115: 117-132, Fig. 1. B, C, D and F, modified from Bunke, D. 1985. Journal of Morphology 185: 203-216, Figs. 1-3. Abbreviations a, acrosome; d distal centriole; m mitochondria; n nucleus; p proximal centriole; v, vesicle.

$ Reproductive Biology and Phylogeny of Annelida Reproduction in Siboglinidae has been studied in some detail. Spermiogenesis and sperm ultrastructure has been studied with various levels of detail in the ‘frenulate’ Siboglinum ekmani by Franzén (1973) and the ‘vestimentiferans’ Riftia pachyptila (Gardiner and Jones 1985; Jones and Gardiner 1985) and Ridgeia piscesae (see Southward and Coates 1989) and is essentially similar in all three taxa (Fig. 3.4A-D). Spermiogenesis in general for the group has been summarized by Bakke (1983), Gardiner and Jones (1993) and Southward (1993). Spermatids develop in large morulae attached to a central cytophore. Usually all stages of development are found in socalled testes (probably coelomic spaces). The mature sperm are filiform with an elongate nucleus and flagellum. No true midpiece is present and two or three mitochondria wrap around the nucleus (Fig. 3.4A, B). Microtubules are present around the nucleus until very late in development. In S. ekmani they are involved in nuclear elongation (Franzén 1973) and are possibly involved in the final positioning of the acrosome in R. piscesae (see Southward and Coates 1989). The acrosome of Siboglinum ekmani is helical and lies at the apex of the nucleus (Fig. 3.4A) (Franzén 1973). Gardiner and Jones (1985) described nearly mature sperm of Riftia pachyptila as having an helical acrosome located off to the side of the nucleus, though they did note that it moves to a more anterior position in later stages observed with light microscopy (Fig. 3.4B). A study by Southward and Coates (1989) of sperm in spawned spermatozeugmata of Ridgeia piscesae indicates that there is a final maturation process that occurs after the emission of the spermatozeugmata. The acrosome slides over the anterior end of the nucleus and the subacrosomal space is thus occupied by the nucleus (Fig. 3.4C, D). The sperm nucleus of siboglinids can be slightly coiled (Siboglinum) or have deep indentations which are occupied by the mitochondria (Ridgeia, Riftia). The anterior portion of the nucleus of Riftia and Ridgeia shows markedly less electron density than the rest of the nucleus (Fig. 3.3B, D). The axoneme abuts the base of the nucleus and is anchored by two centrioles (Fig. 3.3A, B). The tail consists of a 9+2 axoneme and plasma membrane only. Franzén (1973) also noted in S. ekmani that a small proportion of the sperm found were smaller that the rest and that these could possibly be parasperm. Further investigation is required and two types of sperm were not noted by Gardiner and Jones (1985) and Jones and Gardiner (1985) in Riftia pachyptila. Siboglinids that were formally referred to as Pogonophora (also Perviata or Frenulata) shed spermatophores with long filaments into the surrounding seawater (Bakke 1990). These are then gathered by females and fertilization is thought to be internal in Siboglinum. In this genus larvae are brooded in the tube of the female, at least in the species for which there is currently information. An example of a spermatophore of Lamellisabella johanssoni is shown in Fig. 3.4E. The other main clade of Siboglinidae, Vestimentifera (also Obturata or Afrenulata), do not use spermatophores. Rather they form masses that appear to be spermatozeugmata, with the sperm embedded in a sticky matrix (Southward and Coates 1989). The

Annelid Sperm and Spermiogenesis

$

Fig. 3.4. Sperm and spermatophores of Siboglinidae (formerly Pogonophora and/ or Vestimentifera) and Myzostomida. A. Siboglinum ekmani (Frenulata). Diagram of longitudinal section through the head of ‘typical’ sperm. Spaces indicate gaps in the reconstruction of the sperm. Actual length of sperm head Fig. 3.4 Contd. ...

$

Reproductive Biology and Phylogeny of Annelida

spermatozeugmata have elongate tails in Ridgeia piscesae, as seen in the spermatophores of the other siboglinids (Southward and Coates 1989). It has been suggested that Riftia pachyptila is a free spawner (Carey et al. 1989), but Southward and Coates (1989) argue that this was an artifact because the observations were based on a shocked animal and the material emitted seen by Carey et al. (1989) was bundles of late spermatids. Observations of in situ spawning of Riftia pachyptila by Van Dover (1994) indicates that Southward and Coates (1989) were correct and that spermatozeugmata are spawned into the water which then attach to females. The females are then triggered to spawn and after fertilization the eggs are expelled by the female into the surrounding water where development occurs (Van Dover 1994). Van Dover (1994) argued that fertilization was probably internal, based on observations by Jones (1981) that sperm were found in the genital tracts of females. This has now been confirmed by Hilario et al. (2005).

3.4.4 Myzostomida The position of myzostomes is controversial (see chapters 1 and 11) but they are treated as annelids in this volume. A number of other ultrastructural studies have been conducted on spermiogenesis, sperm and spermatophores of myzostomids, though all have been restricted to the genus Myzostoma (see Table 3.1). Spermiogenesis occurs within spermiocysts, with all spermatids in a given spermiocyst at the same stage of development. Spermiogenesis is complicated compared with the process usually found in annelids, though no acrosome has ever been observed at any stage of development. In Myzostoma spp. the nucleus forms into a series of electron dense spheres surrounded by diffuse material. It is often Fig. 3.4 Contd. ...

is 30 µm Modified from Franzén, Å. 1973. Acta Zoologica 54: 179-192, Fig. 15. B. Riftia pachyptila. Diagram of nearly mature sperm still attached to cytophore. The acrosome at this stage is still lateral to the nucleus. Modified from Gardiner, S. L. and Jones, M. L. 1985. Transactions of the American Microscopical Society 104: 19-44 Fig. 58. C. Ridgeia piscesae (Vestimentifera). Immature sperm still attached to cytophore with acrosome in lateral position. Microtubules around the nucleus may be involved in the final movement of the acrosome over the nucleus. D. Ridgeia piscesae. Mature sperm from spermatozeugmata with acrosome now in final position over the apical end of the nucleus. Modified from Southward, E. C. and Coates, K. A. 1989. Canadian Journal of Zoology 67: 2776-2781, Figs.13. E. Spermatophore of Lamellisabella johanssoni (Frenulata). Main body of the spermatophore contains sperm attached to a long filament that is thought to assist in the uptake by females. Modified from Ivanov, A. V. 1963. Pogonophora. Academic Press, London, Fig. 65. F. Myzostomum cirriferum. Drawing of spermatozoon as seen with light microscope. The sperm head appears to be comprised a series of dark spheres. Modified from Jägersten, G. 1934. Zoologiska Bidrag från Uppsala 15: 1-22, Fig. 22. G. Myzostomum cirriferum. Spermatophore. Cysts with sperm are located in the ‘horn’ and ‘body’ regions of the spermatophore. H. Myzostoma cirriferum. Detail of a region of the ‘horn’ of a spermatophore. Note sperm within cysts. G and H, modified from Eeckhaut, I. and Jangoux, M. 1991. Zoomorphology 111: 49-58, Fig. 11.. Abbreviations a, acrosome; b body of spermatophore; f, filament of spermatophore; h ‘horn’; m mitochondria; n nucleus; spt spermatophore body; t, microtubules around nucleus.

Annelid Sperm and Spermiogenesis

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assumed that the electron dense spheres are chromatin (Afzelius 1983; Eeckhaut and Jangoux 1991) and this supported by the fact that granules stain positively for the Feulgen test (Eeckhaut and Jangoux 1991). Mattei and Marchand (1988) on the other hand suggest that the electron dense granules are in fact protein granules and the chromatin is restricted to the periphery of the nuclear region. The number of nuclear granules is usually 42 in M. cirriferum, though it can be as high as 50 (Afzelius 1983; Eeckhaut and Jangoux 1991). During spermiogenesis a number of small mitochondria fuse to form two (sometimes one or three) elongate rods that can be as long as the nuclear region (Afzelius 1984; Mattei and Marchand 1988). A manchette of 16-22 microtubules can be found (depending on the species studied) around the mitochondria during development and persist against the plasma membrane in the mature sperm. Mattei and Marchand’s (1988) interpretation of myzostome spermiogenesis also differs in other ways from the interpretations by Jägersten (1934) and Afzelius (1984). They suggest that the single centriole, a feature not previously observed, and attached axoneme initially migrate through the sperm forming a cytoplasmic canal. The canal then opens along much of its length leaving the flagellum outside the body of the sperm, but still in contact with it via ‘dense connectors’. The centriole, tipped with a ‘spine’, continues to migrate forward until the original posterior end of the flagellum is drawn up to the sperm body and the centriole and most of the flagellum lie ‘anterior’ to the body of the sperm (Fig. 3.4F). The sperm usually swims (in seawater at least) with the flagellum foremost (Afzelius 1983). This form of locomotion is not particularly effective and the sperm can also swim backwards, i.e., with the nucleus and mitochondria foremost and the tail trailing. This form of motion appears to be dependent on the action of the manchette of microtubules (Mattei and Marchand 1988). Reproduction in myzostomes is thought to mainly involve the use of spermatophores, though Wheeler (1898) reported external fertilization in Myzostoma glabrum. This process has been most carefully studied in M. cirriferum (Eeckhaut and Jangoux 1991; Jägersten 1939a, 1939b). Spermatophores in this species are up to 500 µm long and contain two forms of cysts. The cysts in the ‘horn’ and body regions of the spermatophores contain spermatozoa (Fig. 3.3G, H). The cysts in the ‘foot’ region of the spermatophore contain abortive germ cells. Spermatophores are placed on the skin of a receiving adult and the epidermis is lysed, allowing the contents of the spermatophore to pass into the receiver’s body. The sperm then make their way to the oocytes for internal fertilization.

3.5 ANNELID SPERM AND SYSTEMATICS Previous doubts about the use of sperm data as characters in polychaetes systematics (Jamieson and Rouse 1989; Rice 1992) were based on ideas of using sperm data alone for phylogenetic studies. While it is true that sperm ultrastructural characters have proved informative in clades with uniform

$" Reproductive Biology and Phylogeny of Annelida reproductive mechanisms such as clitellates (Jamieson 1981, 1983c, 1984; Jamieson et al. 1987; Ferraguti and Erséus 1999) there was thought to be a problem with polychaetes. Rice (1992: 150) summarized this as “variation in sperm structure within groups (such as polychaetes) may be so extensive that any number of phylogenies could be constructed depending upon which sperm type is considered to be ... plesiomorphic.” Jamieson and Rouse (1989) expressed a similar view and suggested that the supposed multiple origin of ‘modified’ forms of reproduction from external fertilization would preclude the use of sperm in systematics. These statements imply that the understanding the evolution of the tremendous variety of reproductive mechanisms among polychaetes is too problematic. However, it may be that this variability allows tests of hypotheses about the evolution of reproduction in marine invertebrates. If sperm (and other reproductive) characters are incorporated into data sets that include all morphological evidence then the problems nominated above can be eliminated. Under these circumstances the hypothesis that external fertilization is always primitive then becomes testable and homology assumptions about sperm morphology are open to reassessment. This then raises the question of what the plesiomorphic form of sperm shape and fertilization mode is for Annelida.

3.6 WHAT IS THE PLESIOMORPHIC SPERM TYPE IN ANNELIDA? Most previous influential systematizations of polychaetes (e.g. Fauchald 1977) recognise a taxon Phyllodocida, explicitly or implicitly accepting that this is a derived annelid clade. Basal annelids, according to Rouse and Fauchald (1997), are taxa such as Clitellata and simple-bodied forms like Questidae and Paraonidae, currently regarded as part of Scolecida. As pointed out in Rouse and Pleijel (2003) this rooting of Annelida was based on outgroup choices such as Mollusca and Sipuncula, and may well be misleading. However, if it is accepted then it would suggest that the filiform sperm type and complicated reproduction seen in taxa such as most Clitellata, involving fertilization in a cocoon after sperm transfer storage in spermathecae could perhaps be the basal condition for annelids. To accept this would mean that the basal condition for polychaetes such as Scolecida would also have to be filiform sperm and some sort of ‘internal’ fertilization. However, this is currently not known since a wide variety of reproduction from external fertilization to internal fertilization occurs in this group (Rouse and Pleijel 2001). To properly understand the placement of this also requires knowledge of the plesiomorphic condition in the sister group to Annelida. Accepting that Echiura is within Annelida, then most authors would regard Mollusca as the sistergroup to Annelida. The plesiomorphic reproductive condition or molluscs has been suggested to be internal fertilization and filiform sperm (Buckland-Nicks and Scheltema 1995). However, this argument was criticized by Rouse (1999b) as not being

Annelid Sperm and Spermiogenesis

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Fig. 3.5. Unrooted version of cladogram of Annelida. Modified from that shown in Rouse and Pleijel (2003). Major taxa of Annelida are shown with no implication that all are monophyletic. Some interesting minor groups are also shown. The placement of Clitellata and Echiura within Annelida is currently unknown. Some representative sperm types from each taxon are shown in order to demonstrate that establishing the plesiomorphic sperm shape (and fertilization mode) cannot be resolved until the root placement is established. Aeolosomatidae and Potamodrilidae, sperm of Potamodrilus fluviatilis modified from Bunke 1985, Fig. 1; Amphinomida, sperm of the amphinomid Eurythoe complanata modified from Jamieson and Rouse 1989, Fig. 1A; Cirratuliformia. Sperm of Cirriformia sp. modified from Jamieson and Rouse 1989, Fig. #, and Ctenodrilus sp. modified from Rouse 1999a, Fig. 7C; Clitellata, sperm of Capilloventer australis modified from Ferraguti et al. 1996, Fig. 1; Echiura, sperm Fig. 3.5 Contd. ...

$$ Reproductive Biology and Phylogeny of Annelida based on adequate consideration of all the data. Thus, even if we accept the topology of annelid phylogeny proposed by Rouse and Fauchald (1997) we cannot infer the basal reproductive mode and sperm shape for annelids. There are, however, several alternative hypotheses that suggest that the root for the annelid tree in a completely different location to that proposed by Rouse and Fauchald (1997), and this was recently reviewed by Purschke (2002b, and see Chapter 1). Storch (1968), Westheide (1997) and Conway Morris and Peel (1995) all presented hypotheses that would root the ‘crown’ Annelida tree with what is here considered a part of Phyllodocida, or the more inclusive group Aciculata, though none actually explicitly suggests what the relevant basal subgroup would be. Also the reproductive modes in these groups are also quite varied, ranging from external to internal fertilization and our knowledge of the phylogeny within Aciculata subgroups is not well resolved (Rouse and Pleijel 2001). Given this controversy it becomes difficult to estimate what the plesiomorphic reproductive condition for Annelida is. If we accept that the basic topology of annelid relationships postulated in Rouse and Fauchald (1997) is correct but do not root the tree then a diagram as shown in Fig. 3.5 is the result. This may represent the most conservative representation of our understanding of annelid relationships. On this diagram a range of sperm types has been plotted to represent the diversity of sperm and reproductive modes found in annelids. This is an extremely simplistic view since many subgroups of annelids show a wide range of reproductive modes and we have presently have little idea of how they have evolved.

3.7 ACKNOWLEDGEMENTS Thanks to Barrie Jamieson for conceiving this volume and for his comments on this chapter. This work was supported by the Australian Research Council and the South Australian Museum.

Fig. 3.4 Contd. ...

of Bonellia viridis modified from Franzén and Ferraguti 1992, Fig. 8; Eunicida, sperm of the onuphid Hyalinoecia tubicola modified from Jamieson and Rouse 1989, Fig. 1C, and the histriobdellid Stratiodrilus novohollandiae modified from Jamieson et al. 1985, Figs. 19-23; Parergodrilidae, sperm of Parergodrilus heideri modified from Purschke 2002a, Fig. 6, Hrabeiella sperm Hrabeiella periglandulata modified from Rota and Lupetti 1997, Figs. 26, 29, 32, 40; Phyllodocida, sperm of the polynoid Lepidonotus sp. modified from Jamieson and Rouse 1989, Fig. 3A, and the nereidid Platynereis massiliensis modified from Pfannenstiel et al. 1987, Fig. 4; Sabellida, sperm of the serpulids Galeolaria caespitosa modified from Jamieson and Rouse 1989, Fig. 4G and Chitinopoma serrula modified from Franzén 1982a, Fig. 12; Spionida, sperm of the spionids Tripolydora sp. and Prionospio cf. queenslandica modified from Rouse 1988a, Fig. 28; Scolecida, sperm of the capitellid Capitella capitata modified from Franzén 1982a, Fig. 19 and the arenicolid Arenicola marina modified from Jamieson and Rouse 1989, Fig. 2D; Terebelliformia, sperm of the terebellid Streblosoma acymatum modified from Jamieson and Rouse 1989, Fig. 4A, and the alvinellid Paralvinella pandorae pandorae modified from Jouin-Toulmond et al. 2002, Fig. 1.

Annelid Sperm and Spermiogenesis



%$3.8 LITERATURE CITED Afzelius, B. A. 1983. The spermatozoon of Myzostomum cirriferum (Annelida, Myzostomida). Journal of Ultrastructure Research 83: 58-68. Afzelius, B. A. 1984. Spermiogenesis in Myzostomum cirriferum (Annelida; Myzostomida). Videnskabelige Meddelelser fra Dansk naturhistorik Førening i Kjøbenhavn 145: 11-21. Anderson, W. A. and Eckberg, W. R. 1983. A cytological analysis of fertilization in Chaetopterus pergamentaceus. Biological Bulletin. Marine Biological Laboratory, Woods Hole, Mass. 165: 110-118. Bakke, T. 1983. Pogonophora. Pp. 377-385. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates, Volume II: Spermatogenesis and Sperm Function. John Wiley and Sons, Chichester. Bakke, T. 1990. Pogonophora. Pp. 37-48. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates. Volume IV, Part B Fertilization, Development, and Parental Care. John Wiley and Sons, Chichester. Bass, N. R. and Brafield, A. E. 1972. The life-cycle of the polychaete Nereis virens. Journal of the Marine Biological Association of the United Kingdom 52: 701-726. Bentley, M. G. and Pacey, A. A. 1989. A scanning and electron microscopical study of sperm development and activation in Arenicola marina L. (Annelida: Polychaeta). Invertebrate Reproduction and Development 15: 211-219. Bentley, M. G. and Serries, K. 1992. Sperm ultrastructure in two species of the polychaete genus Harmothoe (Polynoidae). Helgoländer wissenschaftliche Meeresuntersuchungen 46: 171-184. Berruti, G., Ferraguti, M. and Lora Lamia Donin, C. 1978. The aflagellate spermatozoon of Ophryotrocha: a line of evolution of fertilization among polychaetes. Gamete Research 1: 287-292. Bertout, M. 1976. Spermatogénèse de Nereis diversicolor O. F. Müller (Annèlide Polychète). I. Evolution du cytoplasme élaboration de l’acrosome. Journal de Microscopie et de Biologie Cellulaire 25: 87-94. Bochert, R. 1996. An electron microscopic study of spermatogenesis on Marenzelleria viridis (Verrill, 1873) (Polychaeta; Spiondae). Acta Zoologica 77: 191-199. Buckland-Nicks, J. and Scheltema, A. 1995. Was internal fertilization an innovation of early Bilateria? Evidence from sperm ultrastructure of a mollusc. Proceedings of the Royal Society of London-Series B: Biological 261: 11-18. Buhrmann, C., Westheide, W. and Purschke, G. 1996. Spermatogenesis and sperm ultrastructure in the interstitial syllid Petitia amphophthalma (Annelida, Polychaeta). Ophelia 45: 81-100. Bunke, D. 1985. Ultrastructure of the spermatozoon and spermiogenesis in the interstitial annelid Potamodrilus fluviatilis. Journal of Morphology 185: 203-216. Bunke, D. 1986. Ultrastructural investigation on the spermatozoon and its genesis on Aeolosoma litorale with considerations on the phylogenetic implications for the Aeolosomatidae. Journal of Ultrastructure and Molecular Structure Research 95: 113-130. Carey, S. C., Felbeck, H. and Holland, N. D. 1989. Observations of the reproductive biology of the hydrothermal vent tube worm Riftia pachyptila. Marine Ecology Progress Series 52: 89-94. Chughtai, I. 1986. Fine structure of spermatozoa in Perkinsiana rubra and Pseudopotamilla reniformis (Sabellidae: Polychaeta). Acta Zoologica 67: 165-171. Colwin, A. L. and Colwin, L. H. 1961. Fine structure of the spermatozoon of Hydroides hexagonus (Annelida), with special reference to the acrosomal region. The Journal of Biophysical & Biochemical Cytology 10: 211-230.

$& Reproductive Biology and Phylogeny of Annelida Conway Morris, S. and Peel, J. S. 1995. Articulated halkieriids from the lower Cambrian of north Greenland and their role in early protostome evolution. Philosophical Transactions of the Royal Society of London. Series B 347: 305-358. Cotelli, F. and Lora Lamia Donin, C. 1975. Ultrastructural analysis of mature spermatozoa of Hyalinoecia tubicola (O. F. Müller) (Annelida; Polychaeta). Monitore Zoologico Italiano 9: 51-66. Cross, N. 1984. Fertilization in Urechis caupo and in polychaetes. Fortschritte der Zoologie 29: 149-166. Daly, J. M. and Golding, D. W. 1977. A description of the spermatheca of Spirorbis spirorbis (L.) (Polychaeta: Serpulidae) and evidence for a novel mode of sperm transmission. Journal of the Marine Biological Association of the United Kingdom 57: 219-227. Defretin, R. and Wissocq, J.-C. 1974. Le spermatozoïde de Nereis irrorata Malmgren (Annélide Polychète). Journal of Ultrastructure Research 47: 196-213. Degaulejac, B., Henry, M. and Vicente, N. 1995. An ultrastructural study of gametogenesis of the marine bivalve Pinna nobilis (Linnaeus 1758). 2. Spermatogenesis. Journal of Molluscan Studies 61: 393-403. Eckelbarger, K. J. 1974. Population biology and larval development of the terebellid polychaete Nicolea zostericola. Marine Biology 27: 101-113. Eckelbarger, K. J. 1984. Ultrastructure of spermatogenesis in the reef-building polychaete Phragmatopoma lapidosa (Sabellariidae) with special reference to acrosome morphogenesis. Journal of Ultrastructure Research 89: 146-164. Eckelbarger, K. J. and Grassle, J. P. 1987. Spermatogenesis, sperm storage and comparative sperm morphology in nine species of Capitella, Capitomastus and Capitellides (Polychaeta: Capitellidae). Marine Biology 95: 415-429. Eckelbarger, K. J. and Young, C. M. 2002. Spermiogenesis and modified sperm morphology in the “seepworm” Methanoaricia dendrobranchiata (Polychaeta: Orbiniidae) from a methane seep environment in the Gulf of Mexico: Implications for fertilization biology. Biological Bulletin 203: 134-143. Eckelbarger, K. J., Young, C. M., Llodra, E. R., Brooke, S. and Tyler, P. 2001. Gametogenesis, spawning behavior, and early development in the “iceworm” Hesiocaeca methanicola (Polychaeta: Hesionidae) from methane hydrates in the Gulf of Mexico. Marine Biology 138: 761-775. Eeckhaut, I. and Jangoux, M. 1991. Fine structure of the spermatophore and intradermic penetration of sperm cells in Myzostoma cirriferum (Annelida, Myzostomida). Zoomorphology 111: 49-58. Eibye-Jacobsen, D. and Kristensen, R. M. 1994. A new genus and species of Dorvilleidae (Annelida, Polychaeta) from Bermuda, with a phylogenetic analysis of Dorvilleidae, Iphitimidae and Dinophilidae. Zoologica Scripta 23: 107-131. Fallon, J. F. and Austin, C. R. 1967. Fine structure of gametes of Nereis limbata (Annelida) before and after interaction. Journal of Experimental Zoology 166: 225242. Fauchald, K. 1977. The polychaete worms. Definitions and keys to the orders, families and genera. Natural History Museum of Los Angeles County. Science Series 28: 1-188. Ferraguti, M. 1983. Clitellata. Pp. 343-376. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates, Volume II. Spermatogenesis and Sperm Function. John Wiley and Sons, Chichester. Ferraguti, M. 1984. The comparative ultrastructure of sperm flagella central sheath in Clitellata reveals a new autapomorphy of the group. Zoologica Scripta 13: 201207.

Annelid Sperm and Spermiogenesis

$'

Ferraguti, M. 1999. Euclitellata. In B. G. M. Jamieson (ed.), Reproductive Biology of Invertebrates, Volume IXB. Progress in Male Gamete Ultrastructure and Phylogeny. Oxford and IBH Publishing Co., New Delhi. Ferraguti, M. and Erséus, C. 1999. Sperm types and their use for a phylogenetic analysis of aquatic clitellates. Hydrobiologia 402: 225-237. Ferraguti, M., Erséus, C., Kaygorodova, I. and Martin, P. 1999. New sperm types in Naididae and Lumbriculidae (Annelida: Oligochaeta) and their possible phylogenetic implications. Hydrobiologia 406: 213-222. Ferraguti, M., Erséus, C. and Pinder, A. 1996. The spermatozoon of Capilloventer australis and the systematic position of the Capilloventridae (Annelida, Oligochaeta). Australian Journal of Zoology 44: 469-478. Ferraguti, M., Marotta, R. and Martin, P. 2002. The double sperm line in Isochaetides (Annelida, Clitellata, Tubificidae). Tissue & Cell 34: 305-314. Fitzhugh, K. and Rouse, G. W. 1999. A remarkable new genus and species of fan worm (Polychaeta: Sabellidae: Sabellinae) associated with marine gastropods. Invertebrate Biology 118: 357-390. Franzén, Å. 1956. On spermiogenesis, morphology of the spermatozoon and biology of fertilization among invertebrates. Zoologiska Bidrag från Uppsala 31: 355-482. Franzén, Å. 1973. The spermatozoon of Siboglinum. Acta Zoologica 54: 179-192. Franzén, Å. 1974. Sperm ultrastructure in some Polychaeta. Pp. 267-278. In B. A. Afzelius (ed.), The Functional Anatomy of the Spermatozoon., Pergamon Press, Oxford. Franzén, Å. 1977a. Sperm structure with regard to fertilization biology and phylogenetics. Verhandlungen der Deutschen Zoologischen Gesellschaft 1977: 123-138. Franzén, Å. 1977b. Ultrastructure of spermatids and spermatozoa in Archiannelida. Zoon 5: 97-105. Franzén, Å. 1982a. Ultrastructure of spermatids and spermatozoa in three polychaetes with modified biology of reproduction: Autolytus sp., Chitinopoma serrula and Capitella capitata. International Journal of Invertebrate Reproduction 5: 185-200. Franzén, Å. 1982b. Ultrastructure of the biflagellated spermatozoon of Tomopteris helgolandica Greef, 1879. Gamete Research 6: 29-37. Franzén, Å. and Ferraguti, M. 1992. Ultrastructure of spermatozoa and spermatids in Bonellia viridis and Hamingia arctica (Echiura) with some phylogenetic considerations. Acta Zoologica 73: 25-31. Franzén, Å. and Rice, S. A. 1988. Spermatogenesis, male gametes and gamete interaction. Pp. 309-333. In W. Westheide and C. O. Hermans (eds), The Ultrastructure of Polychaeta., vol. Microfauna Marina 4, Gustav Fisher Verlag, Stuttgart. Franzén, Å. and Sensenbaugh, T. 1984. Fine structure of spermiogenesis in the archiannelid Nerilla antennata Schmidt. Videnskabelige Meddelelser fra Dansk naturhistorik Førening i Kjøbenhavn 145: 23-36. Gambi, M. C., Giangrande, A. and Patti, F. P. 2000. Comparative observations on reproductive biology of four species of Perkinsiana (Polychaeta: Sabellidae: Sabellinae). Bulletin of Marine Science 67: 299-309. Gambi, M. C. and Patti, F. P. 1999. Reproductive biology of Perkinsiana antarctica (Kinberg) (Polychaeta, Sabellidae) in the Straits of Magellan (South America): Systematic and ecological implications. Scientia Marina 63: 253-259. Gambi, M. C., Patti, F. P., Micaletto, G. and Giangrande, A. 2001. Diversity of reproductive features in some Antarctic polynoid and sabellid polychaetes, with

% Reproductive Biology and Phylogeny of Annelida a description of Demonax polarsterni sp. n (Polychaeta, Sabellidae). Polar Biology 24: 883-891. Gardiner, S. L. and Jones, M. L. 1985. Ultrastructure of spermiogenesis in the vestimentiferan tube worm Riftia pachyptila (Pogonophora: Obturata). Transactions of the American Microscopical Society 104: 19-44. Gardiner, S. L. and Jones, M. L. 1993. Vestimentifera. Pp. 371-460. In F. W. Harrison and M. E. Rice (eds), Microscopic Anatomy of Invertebrates, Volume 12: Onychophora, Chilopoda and Lesser Protostomata, Wiley-Liss, New York. Gelder, S. R. and Ferraguti, M. 2001. Diversity of spermatozoan morphology in two families of Branchiobdellida (Annelida: Clitellata) from North America. Canadian Journal of Zoology 79: 1380-1393. Giangrande, A., Licciano, M., Pagliara, P. and Gambi, M. C. 2000. Gametogenesis and larval development in Sabella spallanzanii (Polychaeta: Sabellidae) from the Mediterranean Sea. Marine Biology 136: 847-861. Giangrande, A. and Petraroli, A. 1994a. Observations on reproduction and growth of Sabella spallanzanii (Polychaeta, Sabellidae) in the Mediterranean Sea. Mémoires du Muséum National d’Histoire Naturelle 162: 51-56. Giangrande, A. and Petraroli, A. 1994b. Sperm morphology of Naineris laevigata (Polychaeta, Orbiniidae). Oebalia 20: 53-59. Giangrande, A., Sciscioli, M., Lepore, E., Mastrodonato, M., Lupetti, P. and Dallai, R. 2002. Sperm ultrastructure and spermiogenesis in two Exogone species (Polychaeta, Syllidae, Exogoninae). Invertebrate Biology 121: 339-349. Gluzman, C. 1994. A fine structural study of the “spermatozeugmata” of Aeolosoma marcusi (Oligochaeta?). Comunicaciones Biologicas 12: 345-355. Gluzman, C. 1997. Sperm cells in Aeolosoma marcusi (Annelida, Oligochaeta). BIOCELL 21: 137-142. Gould-Somero, M. 1975. Echiura. Pp. 277-311. In A. C. Giese and J. C. Pearse (eds), Reproduction of Marine Invertebrates, Vol. III, Annelids and Echiurans, Vol. 3, Academic Press, London. Graebner, I. and Kryvi, H. 1973. The spermiogenesis and mature sperm of Sabella penicillum (Polychaeta). An electron microscopical investigation. Norwegian Journal of Zoology 21: 211-226. Heacox, A. E. and Schroeder, P. C. 1981. A light and electron-microscopic investigation of gametogenesis in Typosyllis pulchra (Berkeley and Berkeley) (Polychaeta: Syllidae). I. Gonad structure and spermatogenesis. Cell and Tissue Research 218: 641-658. Heffernan, P. and Keegan, B. F. 1988. Quantitative and ultrastructural studies on the reproductive biology of the polychaete Pholoe minuta in Galway Bay. Marine Biology 99: 203-214. Hilario, A., Young, C. M. and Tyler, P. A. 2005. Sperm storage, internal fertilization, and embryonic dispersal in vent and seep tubeworms (Polychaeta: Siboglinidae: Vestimentifera). Biological Bulletin 208: 20-28. Hodgson, A. N. and Chia, F. S. 1993. Spermatozoon structure of some North American prosobranchs from the families Lottiidae (Patellogastropoda) and Fissurellidae (Archaeogastropoda). Marine Biology 116: 97-101. Hsieh, H.-L. and Simon, J. L. 1990. The sperm transfer system in Kinbergonuphis simoni (Polychaeta: Onuphidae). Biological Bulletin. Marine Biological Laboratory, Woods Hole, Mass. 178: 85-93. Ivanov, A. V. 1963. Pogonophora., Academic Press, London, 479pp. Jägersten, G. 1934. Studien über den histologischen Bau der männlichen Geschlectsorgane und die Ausbildung des Spermiums die Myzostomum. Zoologiska Bidrag från Uppsala 15: 1-22.

Annelid Sperm and Spermiogenesis

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Jägersten, G. 1939a. Über den Befruchtungsmechanismus der Myzostomiden. Arkiv för Zoologi 30B: 1-4. Jägersten, G. 1939b. Über die Morphologie und Physiologie des Geschlectsapparatus und den Kopulationsmechanismus der Myzostomiden. Zoologiska Bidrag från Uppsala 18: 163-242. Jamieson, B. G. M. 1981. The Ultrastructure of the Oligochaeta. Academic Press, London and New York. 462 pp. Jamieson, B. G. M. 1983a. Spermiogenesis in the oligochaetoid polychaete Questa (Annelida, Questidae). Zoologica Scripta 12: 179-186. Jamieson, B. G. M. 1983b. The ultrastructure of the spermatozoon of the oligochaetoid polychaete Questa sp. (Questidae, Annelida) and its phylogenetic significance. Journal of Ultrastructure Research 84: 238-251. Jamieson, B. G. M. 1983c. Spermatozoal ultrastructure: evolution and congruence with a holomorphological phylogeny of the Oligochaeta (Annelida). Zoologica Scripta 12: 107-114. Jamieson, B. G. M. 1984. A phenetic and cladistic study of spermatozoal ultrastructure in the Oligochaeta (Annelida). Hydrobiologia 115: 3-13. Jamieson, B. G. M. 1986. Onychophoran-euclitellate relationships: evidence from spermatozoal ultrastructure. Zoologica Scripta 15: 141-155. Jamieson, B. G. M., Afzelius, B. A. and Franzén, A. 1985. Ultrastructure of the acentriolar, aflagellate spermatozoa and the eggs of Histriobdella homari and Stratiodrilus novaehollandiae (Histriobdellidae, Polychaeta). Journal of Submicroscopic Cytology 17: 363-380. Jamieson, B. G. M., Erséus, C. and Ferraguti, M. 1987. Parsimony analysis of the phylogeny of some Oligochaeta (Annelida) using spermatozoal ultrastructure. Cladistics 3(2): 145-155. Jamieson, B. G. M. and Rouse, G. W. 1989. The spermatozoa of the Polychaeta (Annelida): An ultrastructural review. Biological Reviews 64: 93-157. Jones, M. L. 1981. Riftia pachyptila, new genus, new species, the vestimentiferan worm from the Galápagos Rift geothermal vents (Pogonophora). Proceedings of the Biological Society of Washington 93: 1295-1313. Jones, M. L. and Gardiner, S. L. 1985. Light and scanning electron microscopic studies of spermatogenesis in the vestimentiferan tube worm Riftia pachyptila (Pogonophora: Obturata). Transactions of the American Microscopical Society 104: 1-18. Jouin, C. 1978. Spermatozoide non flagellé et fécondation externe chez Protodriloides symbioticus (Giard) (Annélides Polychètes, Archiannélides). Vie Milieu (series AB) 28-29: 473-487. Jouin-Toulmond, C., Mozzo, M. and Hourdez, S. 2002. Ultrastructure of spermatozoa in four species of Alvinellidae (Annelida: Polychaeta). Cahiers de Biologie Marine 43: 391-394. Jouin-Toulmond, C. and Purschke, G. 2004. Ultrastructure of the spermatozoa of Parenterodrilus teanioides (Protodrilida: “Polychaeta”) and its phylogenetic significance. Zoomorphology 123: 139-146. Jouin-Toulmond, C., Zal, F. and Hourdez, S. 1997. Genital apparatus and ultrastructure of the spermatozoon in Alvinella pompejana (Annelida: Polychaeta). Cahiers de Biologie Marine 38: 128-129. Jördens, J., Struck, T. and Purschke, G. 2004. Phylogenetic inference regarding Parergodrilidae and Hrabeiella periglandulata (‘Polychaeta’, Annelida) based on 18S rDNA, 28S rDNA and COI sequences. Journal of Zoological Systematics and Evolutionary Research 42: 270-280.

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Kopp, J. C. 1985. A preliminary ultrastructural study of Phragmatopoma (Polychaeta) gametes. Invertebrate Reproduction and Development 8: 297-302. Kristensen, R. M. and Eibye-Jacobsen, D. 1995. Ultrastructure of spermiogenesis and spermatozoa in Diurodrilus subterraneus (Polychaeta, Diurodrilidae). Zoomorphology 115: 117-132. Kristensen, R. M. and Niilonen, T. 1982. Structural studies on Diurodrilus Remane (Diurodrilidae fam.n.), with description of Diurodrilus westheidei sp.n. from the Arctic interstitial meiobenthos, W. Greenland. Zoologica Scripta 11: 1-12. Kubo, M. and Sawada, N. 1977. Electron microscope study on sperm differentiation in Travisia japonica (Polychaeta). Annotationes Zoologicae Japonenses 50: 87-98. Kuper, M. and Westheide, W. 1997a. Sperm ultrastructure and spermatogenesis in the interstitial polychaete Sphaerosyllis hermaphrodita (Syllidae, Exogoninae). Invertebrate Reproduction and Development 32: 189-200. Kuper, M. and Westheide, W. 1997b. Ultrastructure of the male reproductive organs in the interstitial annelid Sphaerosyllis hermaphrodita (Polychaeta, Syllidae). Zoomorphology 117: 13-22. Licciano, M., Giangrande, A. and Gambi, M. C. 2002. Reproduction and simultaneous hermaphroditism in Branchiomma luctuosum (Polychaeta, Sabellidae) from the Mediterranean Sea. Invertebrate Biology 121: 55-65. Lücht, J. and Pfannensteil, H.-D. 1989. Spermatogenesis in Platynereis massiliensis (Polychaeta: Nereidae). Helgoländer wissenschaftliche Meeresuntersuchungen 43: 19-28. Marotta, R., Ferraguti, M. and Erséus, C. 2003a. A phylogenetic analysis of Tubificinae and Limnodriloidinae (Annelida, Clitellata, Tubificidae) using sperm and somatic characters. Zoologica Scripta 32: 255-278. Marotta, R., Ferraguti, M. and Martin, P. 2003b. Spermiogenesis and seminal receptacles in Aeolosoma singulare (Annelida, Polychaeta, Aeolosomatidae). Italian Journal of Zoology 70: 123-132. Mattei, X. and Marchand, B. 1988. La spermiogenèse de Myzostomum sp. (Procoelomata, Myzostomida). Journal of Ultrastructure and Molecular Structure Research 100: 75-85. McHugh, D. 1995. Unusual sperm morphology in a deep-sea hydrothermal-vent polychaete, Paralvinella pandorae (Alvinellidae). Invertebrate Biology 114: 161-168. McHugh, D. and Tunnicliffe, V. 1994. Ecology and reproductive biology of the hydrothermal-vent polychaete Amphisamytha galapagensis (Ampharetidae). Marine Ecology Progress Series 106: 111-120. Meijer, L. 1979. Hormonal control of oocyte maturation in Arenicola marina L. (Annelida, Polychaeta) II. Maturation and Fertilization. Development, Growth and Differentiation 21: 315-329. Micaletto, G., Gambi, M. C. and Piraino, S. 2003. Observation on population structure and reproductive features of Laetmonice producta Grube (Polychaeta, Aphroditidae) in Antarctica waters. Polar Biology 26: 327-333. Mona, M. H., Eissa, S. H. H., Abdel-Gawad, A. M. and Barbary, M.-S. 1994. Ultrastructural investigation of spermatogenesis in the tube-worm Hydroides dirampha (Polychaeta, Serpulidae). Journal of the Egyptian German Society of Zoology 13: 115-128. Nash, R. and Keegan, B. F. 2003. Reproductive cycle of Bispira volutacornis (Polychaeta: Sabellidae) on the west coast of Ireland. Marine Biology 143: 919-925. Nishi, E. 1992. Sperm morphology of serpulid polychaetes, Pomatoleios krausii (Baird), Spirobranchus giganteus corniculatus Pallas, Hydroides elegans Haswell and Salmacina dysteri (Huxley). Galaxea 11: 9-14.

Annelid Sperm and Spermiogenesis

%!

Nordheim von, H. 1989. Vergleichende ultrastrukturuntersuchungen der eu- und paraspermien von 13 Protodrilus arten (Polychaeta, Annelida) und ihre taxonomische und phylogenetische Bedeutung. Helgoländer wissenschaftliche Meeresuntersuchungen 43: 113-156. Olive, P. J. W. 1983. Annelida -Polychaeta (Chapter 16). Pp. 321-342. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates, Vol. II: Spermatogenesis and Sperm Function, John Wiley & Sons Ltd., Chichester. Pacey, A. A. and Bentley, M. G. 1992. An ultrastructural study of spermiogenesis and sperm morula breakdown in Arenicola marina (L.) (Annelida: Polychaeta). Helgoländer wissenschaftliche Meeresuntersuchungen 46: 185-199. Pasteels, J. 1965. La fécondation étudiée au microscope électronique étude comparative. Bulletin de la Société Zoologique de France (Evolution et Zoologie) 90: 195-224. Pernet, B. 2000. Reproduction and development of three symbiotic scaleworms (Polychaeta: Polynoidae). Invertebrate Biology 119: 45-57. Pfannenstiel, H.-D., Grunig, C. and Lucht, J. 1987. Gametogenesis and reproduction in nereidid sibling species (Platynereis dumerilii and P. massiliensis). Bulletin of the Biological Society of Washington 7: 272-279. Pfannesteil, H.-D. and Grünig, C. 1990. Spermatogenesis and sperm ultrastructure in the polychaete genus Ophryotrocha (Dorvilleidae). Helgoländer wissenschaftliche Meeresuntersuchungen 44: 159-171. Picard, A. 1980. Spermatogenesis and sperm-spermathecae relations in Spirorbis spirorbis (L.). International Journal of Invertebrate Reproduction 2: 73-83. Pleijel, F. and Rouse, G. W. 2000. A new taxon, Capricornia (Hesionidae, Polychaeta), illustrating the LITU (‘Least-inclusive taxonomic unit’) concept. Zoologica Scripta 29: 157-168. Pleijel, F. and Rouse, G. W. 2005. Lizardia hirschi, a new hesionid polychaete with external genital organs from the Great Barrier Reef. 208: 69-76. Potswald, H. E. 1967. An electron microscope study of spermiogenesis in Spirorbis (Laeospira) morchi Levinsen (Polychaeta). Zeitschrift für Zellforschung und Mikroscopische Anatomie 83: 231-248. Purschke, G. 2002a. Male genital organs, spermatogenesis and spermatozoa in the enigmatic terrestrial polychaete Parergodrilus heideri (Annelida, Parergodrilidae). Zoomorphology 121: 125-137. Purschke, G. 2002b. On the ground pattern of Annelida. Organisms Diversity & Evolution 2: 181-196. Purschke, G. 2003. Is Hrabeiella periglandulata (Annelida, “Polychaeta”) the sister group of Clitellata? Evidence from an ultrastructural analysis of the dorsal pharynx in H. periglandulata and Enchytraeus minutus (Annelida, Clitellata). Zoomorphology 122: 55-66. Pylilo, I. V. and Vishnjakov, A. E. 1993. Spermatogenesis in Manayunkia baicalensis (Polychaeta: Sabellidae). Issledovaniya-Fauny-Morei: 59-62. Remane, A. 1925. Diagnosen neuer Archianneliden. Zoologischer Anzeiger 65: 15-17. Retzius, G. 1904. Zur kenntnis der spermien der Evertebraten. I. Taf. 1-X11. Biologische Untersuchungen von Gustaf Retzius, Neue Folge 11: 1-32. Retzius, G. 1905. Zur kenntnis der spermien der Evertebraten. II. Biologische Untersuchungen von Gustaf Retzius, Neue Folge 11: 79-102. Rice, S. A. 1981. Spermatogenesis and sperm ultrastructure in three species of Polydora and in Streblospio benedicti (Polychaeta: Spionidae). Zoomorphology 97: 1-16. Rice, S. A. 1987. Reproductive biology, systematics and evolution in the polychaete family Alciopidae. Bulletin of the Biological Society of Washington 7: 114-127.

%" Reproductive Biology and Phylogeny of Annelida Rice, S. A. 1992. Polychaeta: Spermatogenesis and spermiogenesis. Pp. 129-151. In F. W. Harrison and S. L. Gardiner (eds), Microscopic Anatomy of Invertebrates, Volume 7: Annelida, vol. 7, Wiley-Liss, New York. Rice, S. A. and Eckelbarger, K. J. 1989. An ultrastructural investigation of spermatogenesis in the holopelagic polychaetes Vanadis formosa and Krohnia lepidota (Polychaeta: Alciopidae). Biological Bulletin. Marine Biological Laboratory, Woods Hole, Mass. 176: 123-134. Richards, S. L. 1970. Spawning and reproductive morphology of Scolelepis squamata (Spionidae: Polychaeta). Canadian Journal of Zoology 48: 1369-1379. Rota, E. and Lupetti, P. 1997. An ultrastructural investigation of Hrabeiella Pizl & Chalupsky, 1984 (Annelida). 2. The spermatozoon. Tissue and Cell 29: 603-609. Rouse, G. W. 1988a. An ultrastructural study of the spermatozoa from Prionospio cf. queenslandica and Tripolydora sp.: Two spionid polychaetes with different reproductive methods. Acta Zoologica 69: 205-216. Rouse, G. W. 1988b. An ultrastructural study of the spermatozoa of Eulalia sp. (Phyllodocidae), Lepidonotus (Polynoidae), Lumbrinereis sp. (Lumbrinereidae) and Owenia fusiformis (Oweniidae). Helgoländer Meeresuntersuchungen 42: 67-78. Rouse, G. W. 1992a. Ultrastructure of sperm and spermathecae in Micromaldane spp. (Polychaeta: Capitellida: Maldanidae). Marine Biology 113: 655-668. Rouse, G. W. 1992b. Ultrastructure of spermiogenesis and spermatozoa of four Oriopsis species (Sabellinae; Sabellidae; Polychaeta). Zoologica Scripta 21: 363379. Rouse, G. W. 1992c. Ultrastructure of the spermathecae of Parafabricia ventricingulata and three species of Oriopsis (Polychaeta: Sabellidae). Acta Zoologica 73: 141-151. Rouse, G. W. 1993a. Amphiglena terebro sp. nov. (Polychaeta: Sabellidae: Sabellinae) from eastern Australia; including a description of larval development and sperm ultrastructure. Ophelia 37: 1-16. Rouse, G. W. 1993b. New Fabriciola species (Polychaeta, Sabellidae, Fabriciinae) from the eastern Atlantic, with a description of sperm and spermathecal ultrastructure. Zoologica Scripta 22: 249-261. Rouse, G. W. 1995. Is sperm ultrastructure useful in polychaete systematics? An example using 20 species of the Fabriciinae (Sabellidae, Polychaeta). Acta Zoologica 76: 57-74. Rouse, G. W. 1996a. New Fabriciola and Manayunkia species (Fabriciinae, Sabellidae, Polychaeta) from Papua New Guinea. Journal of Natural History 30: 1761-1778. Rouse, G. W. 1996b. A new species of Perkinsiana (Sabellidae, Polychaeta) from Papua New Guinea; with a description of larval development. Ophelia 45: 101-114. Rouse, G. W. 1996c. Variability of sperm storage by females in the Sabellidae and Serpulidae (Polychaeta). Zoomorphology 116: 179-193. Rouse, G. W. 1999a. Polychaeta, including Pogonophora and Myzostomida. Pp. 81124. In B. G. M. Jamieson (ed.), Reproductive Biology of Invertebrates, Volume IXB. Progress in Male Gamete Ultrastructure and Phylogeny. Oxford and IBH Publishing Co., New Delhi. Rouse, G. W. 1999b. Polychaete sperm: phylogenetic and functional considerations. Hydrobiologia 402: 215-224. Rouse, G. W. 2005. Annelid sperm and fertilization biology. Hydrobiologia 535:167178. Rouse, G. W. and Fauchald, K. 1997. Cladistics and polychaetes. Zoologica Scripta 26: 139-204. Rouse, G. W. and Fitzhugh, K. 1994. Broadcasting fables: Is external fertilization really primitive? Sex, size and larvae in sabellid polychaetes. Zoologica Scripta 23: 271-312.

Annelid Sperm and Spermiogenesis

%#

Rouse, G. W. and Gambi, M. C. 1998. Sperm ultrastructure and spermathecal structure in Amphiglena spp. (Polychaeta: Sabellidae). Invertebrate Biology 117: 114-122. Rouse, G. W., Goffredi, S. K. and Vrijenhoek, R. C. 2004. Osedax: Bone-eating marine worms with dwarf males. Science 305: 668-671. Rouse, G. W. and Jamieson, B. G. M. 1987. An ultrastructural study of the spermatozoa of the polychaetes Eurythoe complanata (Amphinomidae), Clymenella sp. and Micromaldane sp. (Maldanidae), with definition of sperm types in relation to reproductive biology. Journal of Submicroscopic Cytology 19: 573-584. Rouse, G. W. and McHugh, D. 1994. Ultrastructure of spermatids and spermatozoa in Ramex californiensis and Nicolea zostericola (Terebellidae; Polychaeta). Ophelia 39: 225-238. Rouse, G. W. and Pitt, K. 2000. Ultrastructure of the sperm of Catostylus mosaicus and Phyllorhiza punctata (Scyphozoa, Cnidaria): Implications for sperm terminology and the inference of reproductive mechanisms. Invertebrate Reproduction & Development 38: 23-34. Rouse, G. W. and Pleijel, F. 2001. Polychaetes. Oxford University Press, London, 354 pp. Rouse, G. W. and Pleijel, F. 2003. Current problems in polychaete systematics. Hydrobiologia 496: 175–189. Rouse, G. W. and Tzetlin, A. B. 1997. Ultrastructure of the body wall and gametogenesis in Cossura cf. longocirrata (Cossuridae Polychaeta). Invertebrate Reproduction and Development 32: 41-54. Sato, M. and Osanai, K. 1983. Sperm reception by an egg microvillus in the polychaete, Tylorrhynchus heterochaetus. Journal of Experimental Zoology 227: 459469. Sato, M. and Osanai, K. 1986. Morphological identification of sperm receptors above egg microvilli in the polychaete, Neanthes japonica. Developmental Biology 3: 263270. Sato, M. and Osanai, K. 1990. Sperm attachment and acrosome reaction on the egg surface of the polychaete, Tylorrhynchus heterochaetus. Biological Bulletin. Marine Biological Laboratory, Woods Hole 178: 101-110. Sawada, N. 1975. Electron microscope studies on sperm differentiation in marine annelid worms. II. Sperm formation in Arenicola brasiliensis. Development, Growth and Differentiation 17: 89-99. Sawada, N. 1984. Electron microscope studies of spermiogenesis in polychaetes. Fortschritte der Zoologie 29: 99-114. Scharnofske, P. 1986. Ultrastructure of sperm morphology of Trilobodrilus axi and T. heideri (Dinophilidae, Polychaeta). Helgoländer wissenschaftliche Meeresuntersuchungen 40: 419-430. Schroeder, P. C. and Hermans, C. O. 1975. Annelida: Polychaeta. Pp. 1-213. In A. C. Giese and J. S. Pearse (eds), Reproduction of Marine Invertebrates. Volume III. Annelids and Echiurans., Vol. 3, Academic Press, New York. Simon, C. A. and Rouse, G. W. 2005. Sperm ultrastructure, spermiogenesis and spermathecal structure in Terebrasabella heterouncinata (Polychaeta: Sabellidae: Sabellinae). Invertebrate Biology 125: 39-49. Sordino, P. and Gambi, M. C. 1994. Reproductive biology and life cycle of Branchiomma luctuosum (Grube 1869) (Polychaeta; Sabellidae) in the Mediterranean Sea. Mémoires du Muséum National d’Histoire Naturelle 162: 640. Sousa, M. and Oliveira, E. 1994. An ultrastructural study of Crassostrea angulata (Mollusca, Bivalvia) spermatogenesis. Marine Biology 120: 545-551.

%$ Reproductive Biology and Phylogeny of Annelida Southward, E. C. 1993. Pogonophora. Pp. 327-369. In F. W. Harrison and M. E. Rice (eds), Microscopic Anatomy of Invertebrates, Vol. 12. Onychophora, Chilopoda and Lesser Protostomata., Wiley-Liss, New York. Southward, E. C. and Coates, K. A. 1989. Sperm masses and sperm transfer in a Vestimentiferan, Ridgeia piscesae Jones 1985 (Pogonophora; Obturata). Canadian Journal of Zoology 67: 2776-2781. Storch, V. 1968. Zur vergleichenden Anatomie der segmentalen Muskelsysteme und zur Verwandtschaft der Polychaeten-Familien. Zeitschrift für Morphologie und Ökologie der Tiere 63: 251-342. Tzetlin, A. B., Dahlgren, T. and Purschke, G. 2002. Ultrastructure of the body wall, body cavity, nephridia and spermatozoa in four species of the Chrysopetalidae (Annelida, “Polychaeta”). Zoologischer Anzeiger 241: 37-55. Van Dover, C. L. 1994. In situ spawning of hydrothermal vent tubeworms (Riftia pachyptila). Biological Bulletin. Marine Biological Laboratory, Woods Hole, Mass. 186: 134-135. Westheide, W. 1984a. The concept of reproduction in polychaetes with small body size: adaptations in interstitial species. Fortschritte der Zoologie 29: 265-287. Westheide, W. 1984b. Genesis and structure of the modified spermatozoon in the interstitial polychate Hesionides arenaria (Annelida). Biology of the Cell 50: 53-66. Westheide, W. 1988a. Genital organs. Pp. 263-279. In W. Westheide and C. O. Hermans (eds), The Ultrastructure of Polychaeta. Microfauna Marina, Vol. 4, Gustav Fisher Verlag, Stuttgart. Westheide, W. 1988b. The ultrastructure of the spermatozoon of Pisione remota (Annelida: Pisionidae) and its transformation in the recaptaculum seminis. Journal of Submicroscopic Cytology and Pathology 20: 169-178. Westheide, W. 1997. The direction of evolution within the Polychaeta. Journal of Natural History 31: 1-15. Westheide, W. and Rieger, R. M. 1987. Systematics of the amphiatlantic Microphthalmus listensis species-group (Polychaeta: Hesionidae): Facts and concepts for reconstruction of phylogeny and speciation. Zeitschrift für Zoologische Systematik und Evolutionsforschung 25: 12-39. Wheeler, W. M. 1898. The maturation, fecundation and early cleavages of Myzostoma glabrum Leuckart. Archive de Biologie, Liège 15: 1-77. Williams, J. D. 2000. Spermiogenesis and spermatozoon ultrastructure of Polydora neocaeca (Polychaeta: Spionidae) from Rhode Island. Invertebrate Reproduction & Development 38: 123-129. Zal, F., Desbruyères, D. and Jouin-Toulmond, C. 1994. Sexual dimorphisms in Paralvinella grasslei, a polychaete annelid from deep-sea hydrothermal vents. Compte rendus de l’Academie des sciences, Paris, Sciences de la Vie 317: 42-48.

Figure 4.1 in color(One) CHAPTER

4

Sexual Strategies and Mating Systems Gabriella Sella

4.1 INTRODUCTION The mating system of a species, i.e. the strategy employed in obtaining mates, is modeled by the selective pressures associated with reproduction through sperm versus reproduction through eggs. According to Bateman’s principle (1948), since eggs are more costly to be produced than sperm, the reproductive success of females is limited by access to resources necessary to produce eggs, whereas the reproductive success of males is limited by the availability of females (i.e. by eggs to fertilize). Therefore male strategies will be different from those of females and reproductive success of males can be highly variable, which can lead to conflicts of interests between the sexes. Up to now a natural selection perspective on the analysis of the structure of the mating system of a species, and consequently on strategies of allocation of energetic resources to male and female reproduction, has been applied to a limited number of animals, including Annelida, where the three sexual modes of reproduction (simultaneous hermaphroditism, sequential hermaphroditism and gonochorism) are all present. In this chapter the selective pressures will be described that maintain mating systems and sex allocation patterns of the best studied annelid model systems.

4.2 SIMULTANEOUS HERMAPHRODITISM Simultaneous hermaphrodites have functional male and female reproductive systems simultaneously active in the same individual for the greatest part of its life. Selective forces involved in life history patterns of many hermaphroditic animals are still poorly understood. Hermaphroditism is correlated with brooding, a sessile or sedentary habit,

Department of Animal and Human Biology, University of Turin, 10123 Turin, Italy

78 Reproductive Biology and Phylogeny of Annelida low density populations, and parasitic or commensal mode of life (Ghiselin 1969, 1974). However, these relationships are not very strict. For example, flatworms, earthworms, or serranid fishes are hermaphroditic but not sedentary, their populations are not at low density, and they do not brood their progeny. According to Charnov (1982), hermaphroditism is a resource allocation strategy evolved to optimize gamete production and related fitness returns. At the allocation level where either the male or the female fitness returns begin to diminish, individuals are selected to shift additional reproductive resources to the other sex and the simultaneous presence of both sexes in the same individual is favored. Diminishing fitness returns of the male function may arise when there are few mating opportunities due to low mobility or low density, while diminishing fitness returns of the female function are expected when progeny do not disperse (thus generating strong sib competition), or when the number of young that can be reared is limited. Since resources devoted to reproduction are limited, a hermaphroditic individual will optimize its reproductive success by limiting investment in that sexual function and diverting the remaining resources to the other sex. In simultaneous hermaphrodites a conflict of interests between the sexes is caused by a biased sex allocation, giving higher returns per investment in one sex function over the other. In externally and internally fertilizing species of annelids this conflict is solved in different ways that have been extensively studied in three groups: in externally fertilizing polychaetes of the genus Ophryotrocha (Premoli and Sella 1995), in the internally fertilizing oligochaete Lumbricus terrestris Linnaeus, 1774 (Michiels 1998; Michiels et al. 2000), and in leeches of the genus Helobdella (Kutschera and Wirtz 1986; Tan et al. 2004). Species of the genus Ophryotrocha live among detritus and fouling fauna of polluted harbors. Density of their populations is supposed to be low, as in most meiofauna species. Their spatial distribution is probably clumped because they tend to aggregate in spots where they converge following mucous trails. In the simultaneously hermaphroditic species Ophryotrocha diadema Åkesson, 1976 (Fig. 4.1), O. gracilis Huth, 1934, O. hartmanni Huth, 1934, mature individuals mate in pairs, following a time consuming courtship to acquire information on the partner’s degree of oocyte maturation. If the partner is accepted, courtship is achieved by pseudo-copulation, i.e. a form of external fertilization in which partners are in close contact. Partners of the same pair regularly take turns in assuming either the male (egg fertilizer) or the female (egg spawner) role, laying eggs every second day. Eggs are much more costly than sperm and therefore both partners are expected to prefer the male role, thus entering in a mating conflict. Reciprocal egg exchange represents a solution to this conflict. By means of this reciprocal egg exchange, the reproductive success of each partner is doubled. Reciprocity is conditional: a mate can fertilize the eggs of its partner only if the latter releases some of its eggs. Reciprocal egg exchange has been defined as egg trading by Fischer (1980), who originally described it for the black hamlet Hypoplectrus nigricans.

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Colour Figure

Fig. 4.1. A pair of hermaphroditic individuals of Ophryotrocha diadema. Both partners have laid eggs. The yellow egg coloration is determined by a dominant allele that controls presence of lutein in egg vitellum, the white egg color is due to the recessive allele. Original.

80 Reproductive Biology and Phylogeny of Annelida Reciprocity in egg exchange would not be evolutionarily stable if safeguards had not evolved against partners which do not reciprocate. In the hermaphroditic species of Ophryotrocha clutches with a small number of eggs are laid at short time intervals. This egg parceling can be considered a form of cheating prevention, since it reduces both the advantages of cheaters (they will fertilize few eggs), and the losses of cheated individuals (they will donate few eggs). As a result of this strategy, reproductive success of a cheater is lowered compared to that of a reciprocating partner. Contrary to hermaphroditic species, gonochoric and sequentially hermaphroditic species of Ophryotrocha lay more than a hundred eggs and spawn every one or two weeks (Premoli and Sella 1995). Co-operation involving repeated egg exchange within a mating pair requires that a mate recognizes its partner among other individuals. Lorenzi and Sella (2000) showed that an indirect mate recognition mechanism (possibly chemical cues from the mucous trails produced by the pair) has evolved, which guarantees that most egg exchanges are performed between the same two individuals. In O. diadema co-operation in egg trading with the same partner persists as long as the opportunity of pairing with a more attractive partner (in terms of egg maturation) is lacking. Frequency of ovigerous hermaphrodites in natural populations is probably not higher than 20% (Sella 1990). Cost of deserting a partner is therefore high: the potential benefit is devalued by the costs of search, courtship and predation (Sella and Lorenzi 2000). Since the probability of encountering a suitable partner is low, the pair bond is relatively stable. In captive populations 30% of the pairs are still stable after four consecutive egg layings (Sella and Ramella 1999). In contrast, in O. gracilis populations, where the proportion of ovigerous hermaphrodites is 48%, only 15% of the pairs are still stable after four consecutive spawnings (Sella et al. 1997). Internally fertilizing hermaphroditic Clitellata may show reciprocal insemination, i.e. both partners donate and receive sperm (see chapters 8 and 9). However, while in an egg trading mating system fertilizations are traded, in a sperm trading mating system hermaphrodites have only a limited control over the fate of their sperm in their partner. Instead of being used for egg fertilization, sperm received may be digested by the receiver. Among internally fertilizing hermaphrodites, digestion of superfluous sperm is very widespread (Michiels 1998). It probably originated because nutrients derived from the digested ejaculate are a compensation for costs of sperm production. In Lumbricus terrestris the spermathecae are known to resorb sperm (Grove 1925). Sperm digestion reduces the competitive ability of an ejaculate and may exert a selective pressure in favor of increased sperm investment (Greef and Michiels 1999), making the male function more and more costly. The only well studied mating systems of reciprocal sperm exchange in hermaphrodites with internal fertilization are those of the sea slugs Navanax inermis (Leonard and Lukowiak 1984) and Chelidonura sandrana (Anthes and Michiels 2004). The mating systems of Lumbricidae, although less well

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known, seem to also correspond to this model. The earthworm Lumbricus terrestris mates in pairs at night on the soil surface. Copulation lasts on average two hours and reciprocal exchange of sperm occurs simultaneously. Copulation is preceded by a long pre-copulatory behavior sequence (up to one hour and a half) during which both partners repeatedly visit each others burrows (Nuutinen and Butt 1997). Burrows are 1–3 m deep vertical tubes. Visiting worms normally retain their tail into their own burrow, and perform long lasting visits with their anterior segments. These repeated pre-copulatory visits have been interpreted in two different ways (Michiels et al. 2000): i) they may give information about reliability of sperm reciprocation and/or about body size of the partner. Sperm donors may prefer larger mates, which are expected to be more fecund; ii) they may be aimed at persuading the partner closer to the individual’s own burrow to mate, thus reducing the risk of having to stretch out over a long distance with risk of desiccation and predation. Insemination in Hirudinea is achieved by mutual exchange of spermatophores, which are attached to the body surface of the partner. After implantation of spermatophores, the body surface of the recipient undergoes histolysis and sperm are released into the body of the partner within three days. Hirudinea carrying two or three spermatophores have frequently been observed, indicating that copulation may be repeated several times with different partners (Kutschera 1992) and multiple paternity within broods may occur (Tan et al. 2004). When multiple ejaculates overlap, the receiver may develop mechanisms for control of received sperm. Often the receiver tries to rub off the spermatophore immediately after receipt or consume it, especially if it is in poor condition. Beside leeches, hypodermic impregnation is common in many other annelid taxa (See chapters 9 and 14), mainly in hermaphroditic ones (myzostomids, dinophilids, and other “archiannelids”). Why has it been adopted? According to Michiels (1998) to receive sperm is costly (manipulation by donor, transmission of diseases), and this may favor the evolution of very quick mating, like in Hirudinea (Kutschera and Wirtz 1986). Successful copulation, on the other hand, does not necessarily imply successful fertilization, received sperm often being used as a nutrient. Injury inflicted to the receiver by hypodermic impregnation (or like in several oligochaetes, by piercing the skin of the mating partner by means of specialized grooved chaetae (Koene et al. 2002), will temporarily reduce its mobility and hence its probability of successive matings. In this way the sperm donor may increase the probability of successful fertilization of its sperm. In mating systems of internally fertilizing hermaphrodites with reciprocal sperm transfer, according to Greeff and Michiels (1999), the combined effects of sperm digestion and sperm competition in the sperm storage organs can result in male investment (sperm and ejaculate production, maintenance of male genitalia, mating behavior) that matches or exceeds female investment. Following Charnov (1982), also in externally

82 Reproductive Biology and Phylogeny of Annelida fertilizing hermaphrodites, the fraction of reproductive resources that each hermaphrodite allocates to the male function will depend only on the mating group size i.e. the number of competing males. If there are no competing males, as in the pair mating situation of Ophrotrocha diadema, or in self-fertilizing species, e.g. Capitomastus minimus Langerhans, 1881 (as reported by Shroeder and Hermans 1975), Terebrasabella heterouncinata (Finley et al. 2001), Hediste limnicola Johnson 1901 (Fong and Pearse 1992), Platynereis massiliensis (Pfannenstiel et al. 1987), Spirorbis spirorbis, S. pagenstecheri (Janua pagenstecheri) (Gee and Williams 1962), Neanthes lichti = Nereis limnicola (Smith 1959), allocation to male gametes is the minimum necessary to fertilize all the eggs offered by the partner. As the number of competing males increases, allocation to sperm production will increase (as shown in the leech Helobdella papillornata by Tan et al. (2004), while female allocation conversely will diminish (as shown in O. diadema by Lorenzi et al. (2005)). In Branchiomma luctuosum Grube, 1869 (Sordino and Gambi 1992) and Spirorbis spirorbis Linnaeus, 1758 (Daly and Golding 1977), where sperm are released freely into water, there is probably strong sperm competition with unrelated sperm for egg fertilization, and the allocation to the male sex is high. Hermaphroditic polychaetes which are able to self-fertilize are reported to have reduced sperm production. In three meiofaunal Exogoninae species, Brania pusilloides (= B. pusilla), Grubeosyllis neapolitana (= G. clavata) and Sphaerosyllis hermaphrodita (Westheide 1990) there are a few anterior male segments followed by a series of female segments (Franke 1999), which indicate reduced male investment, probably due to low sperm competition and reduced mating group size. Many simultaneously hermaphroditic polychaetes and leeches pass through a protandrous or adolescent male phase before reaching the simultaneously hermaphroditic phase. The most likely selective pressure favoring protandry is sexual selection. In a sperm competition situation, male reproductive success is highly variable. Then, mating as male during adolescence may enhance the male lifespan reproductive success. In Ophryotrocha diadema it has been shown that two-thirds of adolescent males are able to sneak into hermaphroditic mating pairs and fertilize eggs, causing a 30% of fitness losses to mature hermaphrodites (Lorenzi and Sella 2003).

4.3 SEQUENTIAL HERMAPHRODITISM Sequential hermaphrodites start out as one sex, and then change to the other sex later in life. Among Annelida sequential hermaphrodites are known only in polychaetes and in the hirudinean genus Helobdella. Among polychaetes one-fifth out of the 73 hermaphroditic species listed by Shroeder and Hermans (1975) are sequentially hermaphroditic, the majority of them being sequentially protandrous and only three sequentially protogynous.

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Sequential hermaphroditism is generally explained by the sizeadvantage hypothesis, first advanced by Ghiselin (1969) and later extended by Charnov (1982). This hypothesis makes two assumptions: i) the reproductive success of an individual as a male or as a female is closely related to body size or age, and ii) the relationship between reproductive success and size or age is different for each sex. Protandry is expected if a larger body size increases female fecundity more than male fertility. Therefore small individuals are expected to be male and large individuals to be female, providing that the cost of sex reversal is not too high. Protogyny is expected when there is sexual selection by male combat and when single males can monopolize many matings. A large body size will favor the male sex. Among marine invertebrates, and especially among polychaetes living in mud in small clumped groups, the high frequency of protandry compared to protogyny may be explained by the strong correlation between body size and fecundity (a large body size will allow a female to produce more eggs than a small female), the frequency of random mating (reproductive success of small and large males will be the same under random mating, and this will cancel the advantages of protogyny), and progeny brooding by parents (Petraitis 1990). To discriminate between true gonochorism and sequential hermaphroditism may be difficult: a cue to infer whether a species is sequentially hermaphroditic or gonochoric is that population sex-ratios are always biased in favor of the first sex in sequentially hermaphroditic species. Some serpulid species, formerly considered gonochoric, have later been revealed to be protandric hermaphrodites. There is now a growing perception that sequential hermaphroditism is largely under-reported in this as well as other taxa (Kupryanova et al. 2001). Among polychaete worms, Ophryotrocha puerilis Claparède and Mecznichov, 1869, is the only protandric sequentially hermaphroditic species whose mating system has been studied so far. Population density fluctuates seasonally, but in the high density season (winter and spring) the clumped spatial distribution offers opportunities for social interactions (Sella and Ramella 1999). Isolated individuals begin sperm production when they have nine body segments and turn to the female sex when they reach 18 + 1 body segments. Reproductive success in females increases with body size, but not in males (Berglund 1986, 1990). Mate choice experiments showed that females prefer to mate with small males. Berglund advanced the hypothesis that this female preference is due to the fact that small males are less likely to change sex compared to large males. Therefore, by choosing small males as mates, females avoid a costly conflict over sex. A male will benefit from changing to the female sex after attaining the 18 segment body size because it will increase its reproductive success. In the mating system of O. puerilis both male competition and female choice are present and counteract each other. Females tend to hormonally inhibit oocytes production in males that have reached the body size turning point (Grothe and Pfannenstiel 1986). This generates a male biased sex ratio and

84 Reproductive Biology and Phylogeny of Annelida consequently a strong male competition for access to females. In this competition large males are generally winners (Berglund 1990), but females (often violently) reject them as mates (Berglund 1986), and therefore they do not gain access to females more often than small males. Ophryotrocha puerilis, Trypanosyllis (= Syllis) zebra Grube, 1870, Syllis amica (Policansky 1982), and leeches of the genus Helobdella (Kutschera and Wirtz 1986) are among the few alternating hermaphroditic species, i.e. that can change sex several times during lifetime. In O. puerilis, after several spawnings, and if both partners have reached the same size, they can change sex simultaneously more than once in their lifetime. Eggs being more costly to produce than sperm, females are more rapidly drained of resources than males and stop growing (Berglund 1990). Therefore for a female that has spawned eggs several times, it pays off to return for a while to the cheaper male sex. Males store up resources and grow more rapidly than females, and when they overtake females in body size they are ready to change sex and produce the costly eggs. Females that, for some unknown reason, never returned to the male sex produced significantly less eggs in the same time interval than females that returned for a while to the male sex (Premoli and Sella 1995).

4.4 GONOCHORISM Within Annelida gonochorism is present only in polychaetes, where it represents the most common form of sexual reproduction. The main sex allocation problem in populations of gonochoric species is how much of their reproductive resources parents must allocate to the production of sons and daughters. If sons and daughters have the same costs for parents the progeny sex ratio should be 1:1. Yet it is not rare to find population sexratios mildly or strongly female biased. Sex ratios in favor of the female sex may be due to sex specific differences in survivorship and/or growth of adults as in Polydora ligni Orth, 1971 and Polydora ciliata Johnson, 1828 (Zajak 1991). Sampling biases related to male or female different foraging behavior or mobility have also been suggested, as in the polynoids Branchipolynoe seepensis, Bathynoe cascadiensis and Gastrolepidia clavigera Schmarda, 1861 (Jollivet et al. 2000). A shift of population sex ratio toward females is expected when populations are subdivided in small breeding groups, and mating opportunities therefore occur between relatives only. Mothers, in such situations, should maximize their fitness by producing only the minimum number of sons to ensure that all their daughters will be fertilized. In contrast, in a large random mating population, one mother’s sons would compete primarily with sons that are not their brothers for mating with females that are not their sisters, and the sex ratio of the mother’s progeny should be at unity. This is the Hamilton’s (1967) Local Mate Competition hypothesis. In populations of Ophryotrocha labronica Bacci and La Greca, 1962 (Prevedelli and Simonini 2002), and of the deep-sea hydrothermal

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scale worm Branchipolynoe seepensis (Jollivet et al. 2000) there is generally a mildly female biased sex ratio. Low mobility of adults is supposed to favor subdivision of populations in small breeding groups where a certain level of inbreeding can be present, as circumstantially suggested by the heterozygote deficiency detected by population genetics methods (Sella and Robotti 1991; Jollivet et al. 2000). If this is the case, a female biased population sex ratio can reduce the local mate competition. The hypothesis that populations with a female biased sex ratio may result from population subdivision could be tested in other polychaete species with a female biased sex ratio, like the serpulid Paraprotis dendrova Uchida 1978, (Nishi and Yamasu 1992a); the terrestrial polychaete Parergodrilus heideri Reisinger, 1925 (Pursche 1999), or the nereidid Hediste diversicolor Müller, 1776 (Smith 1964 as reported by Sato 1999). The Local Mate Competition model seems to explain the mating system of the polychaete Dinophilus gyrociliatus Schmidt, 1857, quite well. Populations of this small interstitial polychaete are probably highly dispersed and at low density. Sex determination is chromosomic with male heterogamety of the XO-XX type, and a 1:1 sex ratio is therefore expected. Eggs are laid in capsules where the ratio of female to male eggs is generally 3:1. Males are dwarf without a digestive system and never come out of the egg capsula. They inoculate their sperm into their still immature sisters by hypodermic impregnation and then die. Females store sperm until they reach maturity and then begin to use their brother’s sperm to fertilize eggs. This generates a highly structured population since there is strict sib mating inside each capsula and absence of mate competition. There is a strong egg size dimorphism with males developing from small (40 µm egg diameter) eggs and females from large eggs (80 µm). Charnov (1987) estimated that 96% of reproductive resources are allocated to daughters compared to sons. A female egg gets about eight times the resources of a male egg, and males produce the minimum amount of sperm necessary to fertilize eggs of, on average, three sisters (Zunarelli-Vandini 1965). The congeneric species Dinophilus taeniatus Harmer, 1889, lives as free swimming in tidal pools where populations can reach very high densities. The lack of patchy distribution within each tide pool means random mating and absence of local mate competition. Mothers gain the same reproductive success through sons and daughters, and selection is expected to favor equal investment in progeny of both sexes. Population sex ratio is 0.5 and both sexes have the same body size (Jennings and Donworth 1986). In gonochoric animals sexual dimorphism is generally the result of sexual selection in mating systems where there is male–male competition for access to females. In the majority of gonochoric polychaetes sexual dimorphism is generally not pronounced. However, the large genital chaetae of males of Capitella sp. I (Petraitis 1990), and the conspicuous dorso-lateral processes of males of Ophryotrocha cosmetandra (Oug 1990), may be used in male competition. Male combat for access to females has been observed in polychaetes in the gonochoric Harmothoe imbricata

86 Reproductive Biology and Phylogeny of Annelida Rasmussen, 1956 (Daly et al. 1972), in Neanthes (=Nereis) arenaceodentata (Weinberg et al. 1990), in Ophryotrocha labronica (Berglund 1991), and in the sequentially hermaphroditic species O. puerilis (Berglund 1990).

4.5 MIXED STRATEGIES AND FACULTATIVE CHANGE OF GENDER Some animal populations consist of individuals that change and others that do not change sex, or of gonochoric and simultaneously hermaphroditic individuals. According to Ghiselin (1987) this variability in sex phenotypes could be maintained by a sort of balanced polymorphism, with different advantages to each strategy. Opportunistic sex change can be expected when the local habitat is characterized by strong fluctuations in food or mate availability. Some conditions may be more favorable for one sex or the other when there are sex specific differences in mating success, in resource exploitation, or in predation. Facultative change of gender may be a rewarding solution in species, like many polychaetes, that have wide dispersal of juveniles but low mobility of adults. If an individual is unable either to assess habitat quality before settlement or to move from the habitat it chose as a juvenile, then it may be the wrong sex for a particular habitat (Petraitis 1988). Sex in such cases is often environmentally determined or controlled by a polygenic sex determination mechanism, which is unstable and poorly canalized, and therefore easily modified by social and developmental factors (Premoli et al. 1996). The capitellid Capitella sp. I provides an exceptional model system for the study of facultative control of gender in a diploid species with female heterogamety. Populations of Capitella sp. I are often structured at low densities, but subject to explosive blooms in disturbed patches. They have a female biased sex ratio, and adults build temporary mucous tubes and show low mobility (Petraitis 1991). Fertilization is internal. Some males can develop eggs and function as either sex when individuals are reared in small groups with low proportion of females. However, low density alone cannot justify this change of sexual strategy, because it does not explain why only males and not females become hermaphrodites. Petraitis (1985) suggests that change from males to hermaphrodites is an adaptation to living in small groups with strong local mate competition. Males that cannot obtain mates become hermaphrodites and function primarily as females, thus avoiding mate competition. Petraitis (1988) demonstrated that a hermaphrodite offsets its loss in mating success through the male function by its success through the female function. It is likely that mate competition is important in other polychaetes with internal fertilization or pseudo-copulation, since other species, apart from Capitella, are known to have populations composed by a mixture of males, females and hermaphrodites, e.g. Polydora ligni Orth, 1971 (Radashevski 1989), Euclymene oerstedii Claparède, 1863 (Pilgrim 1964) and Salmacina dysteri (Nishi and Yamasu 1992b). The syllid Exogone enaidina (= E.

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gemmifera) Örsted, 1845 (Cognetti -Varriale and Zerneri 1965), Grubeosyllis (=Brania) clavata (Franke 1999) and Typosyllis (= Syllis) prolifera Krohn, 1852 (Franke 1986) are stolonizing free-spawners with a planktonic larval recruitment but with a sedentary habitat. Their populations tend to have a male biased sex ratio, even if laboratory experiments showed that the overall primary sex ratio is 1:1. In T. prolifera individuals that start their life as males are absolutely stable during successive reproductive cycles, while individuals that are primarily females can undergo irreversible sex change to the male phase at one of their subsequent reproductive cycles. Sex change does not depend on the sex of social partners but on population density; it is delayed or suppressed under high density population (Franke 1999). One may wonder why determination of female sex is labile in syllids. A possible answer is that having external fertilization, fertilization efficiency may be low unless large amounts of sperm are produced. A second possibility is that male stolons may be less costly than female stolons. In adverse and not yet investigated conditions, such as nutritional deficiencies, the first female reproductive effort could leave a very low residual reproductive potential for successive spawnings, and could thus induce a change to the male sex. In Grubeosyllis (= Grubea) clavata mature females spawn only when stimulated by a male pheromone. Oocytes are resorbed in the absence of this stimulus and worms become males. This suggests that a social influence is necessary to induce a worm to stay in the female phase (Durchon 1975).

4.6 PARENTAL CARE The pattern of parental care provided by each sex is an important aspect of the mating system of a species. Parental care is costly and to provide some form of care to its own brood therefore means that parents must trade energy allocated to this activity with energy devoted to other fitness components like survival, mating success, and fecundity. Often species that brood their progeny produce a much smaller number of eggs than nonbrooding congeners (Olive 1985). Parental care is frequently associated with hermaphroditism, small body size, copulation or pseudo-copulation, high certainty of paternity, interstitial habit, scarcity of food. However, the matter is still poorly understood. Parental care is observed in many polychaetes (see reviews by Wilson, 1991 and Giangrande 1997) and leeches. Protection of embryos has been achieved in polychaetes in three main ways: incubation of embryos in the parental tube (e.g. in Serpulidae), in a mucous capsule (e.g. in Sabellidae and Dorvilleidae), or by carrying young in chambers representing modifications of the adult body (e.g. in Nerillidae, Syllidae, many Serpulidae and Sabellidae), or under scales on the dorsal surface (e.g. Polynoidae). In hermaphroditic species of Ophryotrocha both parents care for eggs even if, at least in laboratory conditions, one parent is as good as two in caring for the brood (Sella 1991). The second parent could desert. This,

88 Reproductive Biology and Phylogeny of Annelida however, will happen only if the breeding sex ratio is 0.5, and if the deserting parent has a high probability of mating soon again. In populations of O. diadema ovigerous hermaphrodites make up only 20% of the breeding population, rendering the probability of re-mating soon very low. This explains why one of the two mates does not desert: bi-parental care is a by-product of the necessity of securing a partner for egg reciprocation. In O. gracilis populations frequency of ovigerous hermaphrodites rises to nearly 50%, and the breeding sex ratio is much more favorable to partner desertion than in O. diadema. Accordingly in the mating system of O. gracilis there is a low investment in parental care and the pair bonds are looser than in O. diadema (Sella et al. 1997). In different populations of the gonochoric O. labronica, when population sex ratios are 1:1, both parents are engaged in parental care; if population sex-ratio is female biased, the rarer (male) sex deserts and only females take care of eggs (Sella and Premoli 1995). Paternal care is unusual in marine invertebrates: it probably evolved when the advantages of caring for embryos outweighed the advantages of doing something else. Among polychaetes only three nereidid species are reported to have paternal care. Nereidids are generally semelparous with epitoky, both sexes spend about 70% of their energetic resources in reproduction (Porchet and Olive 1987), they die after spawning and the eggs develop without parental care. The mating systems of Neanthes (= Nereis) arenaceodentata and Perinereis massiliensis are different, and spawning occurs in pairs without formation of epitokes. Pair formation involves aggressive encounters between males, where winners, usually the largest of the two opponents, are more likely to obtain a mate. Both partners stay together in a mucus lined burrow. Females lay eggs and die, probably investing all their energetic resources in reproduction, while males care for eggs and reproduce more than once. Male costs of reproduction, including costs of egg care, are probably relatively low compared to female costs. Once inside the tube they do not suffer much sperm competition (although some fighting against intruders has been observed) and therefore they do not need to produce large amount of sperm (Weinberg et al. 1990). Brooding in Clitellata has evolved only in glossiphoniid leeches, where it consists of attaching cocoons and hatched larvae to the belly of the parent and providing resources for young. Predation pressure from water snails is considered as the most important factor in the evolution of parental care in leeches. In the eight species of Helobdella that have been studied in detail (H. triserialis Blanchard, 1849, H. striata Kutschera, 1985, H. californica Kutschera, 1988, H. stagnalis Linnaeus, 1758, H. conifera and three undescribed species; Kutschera and Wirtz 2001), attached larvae and young are carried around by parents. The adult leeches may attack water snails and Tubifex worms by inserting their proboscis into the soft parts of the snail or worm and sucking the body fluid, and the attached juveniles participate in the meal (Kutschera 1992). Up to now, this is the only known case of an invertebrate species for which it has been shown that parents

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collect food and offer it directly to their offspring. A consequence of this form of care-giving is that continued contact with offspring permits a finer degree of control over brood number and body size of offspring.

4.7 LITERATURE CITED Anthes, N. and Michiels, N. K. 2004. Do sperm trading simultaneous hermaphrodites always trade sperm? Behavioral Ecology 16: 188–198. Bateman, A. J. 1948. Intrasexual selection in Drosophila. Heredity 2: 349–368. Berglund, A. 1986. Sex change by a polychaete: effects of social and reproductive costs. Ecology 67: 837–845. Berglund, A. 1990. Sequential hermaphroditism and the size advantage hypothesis: an experimental test. Animal Behavior 39: 426–433. Berglund, A. 1991. To change or not to change sex: a comparison between two Ophryotrocha species (Polychaeta). Evolutionary Ecology: 128–135. Charnov, E. 1982. The Theory of Sex Allocation. Princeton University Press, Princeton, New Jersey, U.S.A. 355 pp. Charnov, E. 1987. Local mate competition and sex ratio in the diploid worm Dinophilus gyrociliatus. International Journal of Invertebrate Reproduction 12: 223–225. Cognetti-Varriale, A. M. and Zerneri, G. 1965. Ricerche sulla biologia riproduttiva dei Policheti.Gametogenesi e gonocorismo labile in Exogone gemmifera (Polychaeta, Syllidae). Archivio Zoologico Italiano 50: 59–65. Daly, J. M., Evans, S. M., and Morley, J. 1972. Changes in behavior associated with pair formation in the polychaete Harmothoë imbricata. Marine Behavior Physiology 1: 49–69. Daly, J. M. and Golding, D. W. 1977. A description of the spermatheca of Spirorbis spirorbis and evidence for a novel mode of sperm transmission. Journal of Marine Biology Association of the United Kingdom 57: 219–227. Durchon, M. 1975. Sex reversal in the Syllinae (Polychaeta:Annelida). Pp 41-47. In R. Reinboth (ed.), Intersexuality in the Animal Kingdom, Springer Verlag, Berlin. Finley, C. A, Mulligan, T., and Friedman, C. S. 2001. Life history of an exotic sabellid polychaete, Terebrasabella heterouncinata: fertilization strategy and influence of temperature on reproduction. Journal of Shellfish Research 20: 883–888. Fischer, E. A. 1980. The relationship between mating system and simultaneous hermaphroditism in the coral reef fish Hypoplectrus nigricans (Serranidae). Animal Behavior 28: 620–633 Fong, P. P. and. Pearse, J. S. 1992. Photoperiodic regulation of parturition in the selffertilizing viviparous polychaete Neanthes limnicola from central California. Marine Biology 112: 81–89. Franke, H. D. 1986. Sex ratio and sex change in wild and laboratory populations of Typosyllis prolifera (Polychaeta). Marine Biology 90: 197–208. Franke, H. D. 1999. Reproduction of the Syllidae (Annelida: Polychaeta). Hydrobiologia 402: 39–55. Gee, J. and Williams, G. B. 1965. Self and cross fertilization in Spirorbis borealis and S. pagenstecheri. Journal of the Marine Biology Association of the United Kingdom 45: 275–285. Giangrande, A.1997. Polychaete reproduction patterns, life-cycles and life-histories: an overview. Oceanography and Marine Biology Annual Review 35: 323–386. Ghiselin, M. T. 1969. The evolution of hermaphroditism among animals. Quarterly Review of Biology 44: 189–208.

90 Reproductive Biology and Phylogeny of Annelida Ghiselin, M. T. 1974. The Economy of Nature and the Evolution of Sex. University of California Press, Berkeley, California. 346 pp. Ghiselin, M. T. 1987. Evolutionary aspects of marine invertebrate reproduction. Pp: 608–665. In A. C. Giese and J. S Pearse (eds), Reproduction of Marine Invertebrates, Vol. IX, Blackwell Scientific Publications and the Boxwood Press, Palo Alto, California. Greeff, J. M. and Michiels, N. K. 1999. Sperm digestion and reciprocal sperm transfer can drive hermaphrodite sex allocation to equality. The American Naturalist 53: 421–430. Grothe, C. and Pfannenstiel, H. D. 1986. Cytophysiological study of neurosecretory and pheromonal influences of sexual development in Ophryotrocha puerilis. International Journal of Invertebrate Reproduction and Development 10: 227– 239. Grove, A. J. 1925. On the reproductive processes of the earthworm Lumbricus terrestris. Quarterly Journal of Microscopy Science 69: 245–291. Hamilton, W. D. 1967. Extraordinay sex ratios. Science 156: 477–488. Jennings, J. B. and Donworth, P. J. 1986. Observation on the life cycle and nutrition of Dinophilus taeniatus Harmer, 1889 (Annelida, Polycheta). Ophelia 25: 119–137. Jollivet, D., Empis, A., Baker, M. C., Hourdez, S., Comtet, T., Jouin-Toulmond, C., Desbruyères, D. and Tyler, P. A. 2000. Reproductive biology, sexual dimorphism and population structure of the deep sea hydrothermal vent scale-worm, Branchypolinoe seepensis (Polychaeta: Polynoidae). Journal of the Marine Biology Association of the United Kingdom. 80: 55–68. Koene, J. M., Sundermann, G. and Michiels, N. K. 2002. On the function of body piercing during copulation in earthworms. International Journal of Invertebrate Reproduction and Development 41: 35–40. Kupryanova, E. K., Nishi, E., Ten Hove, H. A. and Rzhavsky, A. V. 2001. Life-history patterns in serpulimorph polychaetes: Ecological and evolutionary perspectives. Oceanography and Marine Biology 39: 1–101. Kutschera, U. 1992. Reproductive behaviour and parental care of the leech Helobdella triserialis (Hirudinea: Glossiphoniidae). Zoologischer Anzeiger 2: 74–81. Kutschera, U. and Wirtz, P. 1986. Reproductive behaviour and parental care of Helobdella striata (Hirudinea, Glossiphoniidae): a leech that feeds its young. Ethology 72: 132–142. Kutschera, U. and Wirtz, P. 2001. The evolution of parental care in freshwater leeches. Theory in Biosciences 120: 115–137. Leonard, J. L. and Lukoviak, K. 1984. Male–female conflict in a simultaneous hermaphrodite resolved by sperm trading. The American Naturalist 124: 282–286. Lorenzi, M. C. and Sella, G. 2000. Is individual recognition involved in the maintenance of pair bonds in Ophryotrocha diadema (Dorvilleidae, Polychaeta)? Ethology, Ecology and Evolution 12: 197–202. Lorenzi, M. C. and Sella, G. 2003. Increased sperm allocation delays body growth in a protandrous simultaneous hermaphrodite. Biological Journal of the Linnean Society 78: 149–154. Lorenzi, M. C., Schleicherova, D., Sella, G., and Ramella, L. 2005. Outcrossing hermaphroditic polychaete worms adjust their sex allocation to social conditions. Journal of Evolutionary Biology 18: 1341-1347. Michiels, N. K. 1998. Mating conflicts and sperm competition in simultaneous hermaphrodites. Pp. 219–254. In T. R. Birkhead and A. P. Möller (eds), Sperm Competition and Sexual Selection. Academic Press, New York.

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Michiels, N. K., Holmer, A. and Vorndran, I. C. 2000. Precopulatory mate assessment in relation to body size in the earthworm Lumbricus terrestris: avoidance of dangerous liaisons? Behavioral Ecology 12: 612–629. Nishi, E. and Yamasu, T. 1992a. Brooding habit and larval development in the serpulid worm Paraprotis dendrova Uchida (Annelida, Polychaeta, Sedentaria). Bulletin of the College of Science University of the Ryulkyus 54: 83–92. Nishi, E. and Yamasu, T. 1992b. Brooding and development of a serpulid tube worm Salmacina dysteri (Huxley) (Polychaeta). Bulletin of the College of Science University of the Ryulkyus 54: 107–121. Nuutinen, V. and Butt, K. R. 1997. The mating behaviour of the earthworm Lumbricus terrestris (Oligochaeta: Lumbricidae). Journal of Zoology 242: 783–798. Olive, P. J. W. 1985. Covariability of reproductive traits in marine invertebrates: implications for the phylogeny of the lower invertebrates. Pp 42–59. In S. Conway Morris, D. George, R. Gibson, and H. M. Platt (eds) The Origins and Relationships of Lower Invertebrates. Clarendon Press, Oxford, U.K. Oug, E. 1990. Morphology, reproduction and development of a new species of Ophryotrocha (Polychaeta, Dorvilleidae) with sexual dimorphism. Sarsia 5: 191– 201. Petraitis, P. S. 1985. Females inhibit male propensity to develop into simultaneous hermaphrodites in Capitella species I (Capitellidae). The Biological Bulletin 168: 395–402. Petraitis, P. S. 1988. Occurrence and reproductive success of feminized males in the polychaete Capitella capitata (species type 1). Marine Biology 97: 403–412. Petraitis, P. S. 1990. Dynamics of sex change in a capitellid polychaete. Pp. 127–154. In M. Mangel (ed.), Sex Allocation and Sex Change: Experiments and Models. The American Mathematical Society, Providence, Rhode Island. Petraitis, P. S. 1991. The effects of sex ratio and density on the expression of gender in the polychaete Capitella capitata (species type 1). Evolutionary Ecology 5: 393– 402. Pfannenstiel, H. D., Grünig, C., and Lücht, J. 1987. Gametogenesis and reproduction in nereid sibling species (Platynereis dumerilii and P. massiliensis). Biological Society of Washington Bulletin 7: 272–279. Pilgrim, M. 1964. The functional anatomy of the reproductive system of the polychaetes Clymenella torquata and Caesicirrus neglectus. Proceedings of the Zoological Society of London 143: 443–464. Policansky, D. 1982. Sex change in plants and animals. Annual Review of Ecology and Systematics 13: 471–495. Porchet, M. and Olive, P. J. W. 1987. Aspects physiologiques des stratégies de reproduction chez les Annélides Polychètes. I. Semelparité et iteroparité. Année Biologique 113: 45–250. Premoli, M. C. and Sella, G. 1995. Sex economy in benthic polychaetes. Ethology, Ecology and Evolution 7: 27–48. Premoli, M. C., Sella, G. and Berra, G.P. 1996. Heritable variation of sex ratio in a polychaete worm. Journal of Evolutionary Biology 9: 4–854. Prevedelli, D. and Simonini, R. 2002. Relationships between body size and population growth rate in two opportunistic polychaetes. Journal of Marine Biology Association of the United Kingdom 82: 403–408. Purschke, G. 2002. Male genital organs, spermatogenesis and spermatozoa in the enigmatic terrestrial polychaete Parergodrilus heideri (Annelida, Parergodrilidae). Zoomorphology 121: 125–138.

92 Reproductive Biology and Phylogeny of Annelida Radashevsky, V.I. 1989. Ecology, sex determination, reproduction and larval development of commensal polychaetes Polydora commensalis and Polydora glycymerica in the Sea of Japan. Pp 137–164. In V. A. Sveshnikov (ed), Symbiosis in Marine Animals. Russian Academy of Sciences, Moscow. Sato, M. 1999 Divergence of reproductive and developmental characteristics in Hediste (Polychaeta: Nereidae). Hydrobiologia 402: 129–143. Schroeder, C. and Hermans, C. O. 1975. Annelida: Polychaeta. Pp. 1–169. In A. C. Giese and J. S Pearse (eds), Reproduction in Marine Invertebrates, Vol. III. Academic Press, New York. Sella, G. 1990. Sex allocation in the simultaneously hermaphroditic polychaete worm Ophryotrocha diadema. Ecology 71: 27–32. Sella, G. 1991. Evolution of biparental care in the hermaphroditic polychaete worm Ophryotrocha diadema. Evolution 45: 63–68. Sella, G., Premoli, M. C. and Turri, F. 1997. Egg trading in the simultaneously hermaphroditic polychaete worm Ophryotrocha gracilis. Behavioral Ecology 8: 83– 86. Sella, G. and Ramella, L. 1999. Sexual conflict and mating systems in the dorvilleid genus Ophryotrocha and the dinophilid genus Dinophilus. Hydrobiologia 402: 203– 213. Sella, G. and Lorenzi, M. C. 2000. Partner fidelity and egg reciprocation in the simultaneously hermaphroditic polychaete worm Ophryotrocha diadema. Behavioral Ecology 1: 260–264. Sella, G. and Robotti, C. 1991. Heterozygote deficiency at the phosphoglucose isomerase locus in a Tyrrhenian population of Ophryotrocha labronica (Polychaeta, Dorvilleidae). Ophelia, Suppl. 5: 641–645. Smith, R. I. 1959. Embryonic development in the viviparous polychaete Neanthes lighti Hartman. Journal of Morphology 87: 417–466. Sordino, P. and Gambi, M. C. 1992. Prime osservazioni sulla biologia riproduttiva e sul ciclo vitale di Branchiomma luctuosum (Grube 1869) (Polychaeta, Sabellidae). Oebalia supplement 17: 425–427. Tan, G. N., Govedich, F. R. and Burd, M. 2004. Social group size, potential sperm competition and reproductive investment in a hermaphroditic leech, Helobdella papillornata (Euhirudinea: Glossiphonidae). Journal of Evolutionary Biology 17: 574–580. Weinberg, J. R., Starczak, V. R., Müller, C., Pesch, G. C. and Lindsay, S. M. 1990. Divergence between populations of a monogamous polychaete with male parental care: premating isolation and chromosome variation. Marine Biology 107: 105–213. Wilson, W. H. 1991. Sexual reproductive modes in polychaetes: classification and diversity. Bulletin of Marine Science 48: 500–516. Zajak, R. D. 1991. Population ecology of Polydora ligni (Polychaeta, Spionidae). I. Seasonal variation in population characteristics and reproductive activity. Marine Ecology Progressive Series 77: 197–206. Zunarelli-Vandini, R. 1965. A possible way of origin of parthenogenetic strains of Dinophilus apatris (D. gyrociliatus). Experientia 21: 1–3.

CHAPTER

5

Early Annelid Development, A Molecular Perspective Steven Q. Irvine and Elaine C. Seaver

5.1

INTRODUCTION

The developmental biology of annelids has been actively studied for more than 100 years. Early in this history it was recognized that annelids are spiralians, sharing basic developmental patterns with a number of other protostome phyla, notably the mollusks. They are also, along with the vertebrates and arthropods, a major phylum of segmented eucoelomate animals. Recent revisions in metazoan phylogeny have placed the annelids as the most intensively studied, in terms of development, among the phyla of the Lophotrochozoa (Halanych et al. 1995; Tessmar-Raible and Arendt 2003). This recognition of a superphylum Lophotrochozoa has made study of annelid molecular development patterns important as an evolutionary comparison with the model systems from the other two major triploblastic clades, the Edysozoa, containing Drosophila, and Deuterostomia containing the chordates. Annelids have several strengths as organisms for developmental studies. Many species can be easily reared in the lab and spawned in large numbers. Many have minimal yolk and so have transparent embryos that cleave holoblastically. Many also have a stereotypic spiralian cleavage pattern, with relatively invariant cell lineages that have been traced in detail in certain species. These characteristics make their study complementary to that of insects, with syncytial cleavage, and vertebrates, with large regulative cell populations. In the case of the heavily studied glossiphoniid leeches, cell lineage can be traced throughout embryogenesis at a single cell level (Weisblat and Stent 1978; Weisblat and Shankland 1985). This characteristic has enabled powerful experimental techniques such as inhibition of various cell processes, intracellular lineage tracing, specific cell ablations, and even

Department of Biological Sciences, University of Rhode Island, Kingston, RI 02811, USA, and Kewalo Marine Laboratory, University of Hawaii, Honolulu, HI, 96813, USA

94 Reproductive Biology and Phylogeny of Annelida moving of whole clones of cells outside their normal territories. These methods are now being combined with molecular techniques to make for potent experimental manipulations. While annelids have a number of amenable characteristics, they are not easily adapted to genetic experimentation. To date, no mutant screens that might result in the identification of genes important to particular developmental processes, such as those done in flies, zebrafish, or mice, have been performed on any annelids. Therefore, molecular developmental biology of annelids is currently pursued using a candidate gene approach. This strategy is dependent on the discovery within the last twenty years that many, if not most, developmentally important genes are present as homologs in the different metazoan phyla. Therefore, genes whose sequence and function have been identified in a genetic model system, generally the fruit fly Drosophila melanogaster, can be candidates for a functional role in development in annelids. Using techniques dependent on certain highly conserved motifs in the nucleic acid sequences of the gene or its transcripts, the annelid homolog of the gene can be cloned. Once cloned, the function of the gene may be studied using a number of techniques. The majority of work to date has been in the determination of temporal and spatial gene expression patterns, either through in-situ hybridization to examine expression patterns of mRNA, or through antibody staining for expression patterns of proteins. These studies have in some cases been combined with experimental manipulations of the embryo, as mentioned above, to examine the role of the cellular environment on gene expression. The candidate gene approach has yielded much new insight into aspects of the molecular control of development. In many cases, as will become apparent below, expression patterns of the genes studied appear to be much different than those in flies and vertebrates. This makes for an interesting situation, but a difficult one without the option of direct functional gene discovery through genetics. New techniques, including genomic approaches, microarray technology, and methods of perturbing gene expression, such as RNA interference, may help the field around the genetic limitations to proceed further toward understanding the unique characteristics of annelid development. By far the majority of work in molecular and cellular development patterns in annelids has been done in a few species of glossiphoniid leeches, namely Helobdella robusta, Helobdella triserialis, Theromyzon rude, and the medicinal leech, Hirudo medicinalis, largely because of their utility as laboratory organisms. More recently, current techniques are being developed for a few polychaete (e.g. Platynereis (Tessmar-Raible and Arendt 2003) and Capitella (Seaver et al. 2005)) and oligochaete (e.g. see Arai and Shimizu 2001; Bely and Wray 2001) species. While most of the literature, therefore, deals with development in leeches, we will endeavor to include material from all three annelid classes. The first part of the chapter organizes information on the various genes that have been studied alongside the developmental processes with which

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they are associated. We then discuss two applications of the candidate gene approach, the Hox and engrailed genes. Finally, we make a summary comparison between the molecular developmental patterns seen in annelids with those of other well-studied taxa, and suggest some directions of future research.

5.2 THE EGG Since the early days of annelid embryology (e.g. Lillie 1902; 1909; Morgan 1910a), it has been known that the eggs of annelids contain maternal determinants, mRNAs and proteins, that are differentially localized to blastomeres during the first few cleavage divisions. These determinants have been shown to affect cell fate in a predictable way, when normally distributed to the early cleavage blastomeres, and led to the idea that annelids exhibit mosaic, or determinative development. Because of the apparent importance of these maternal factors, attention has been directed to understanding the organization of the egg. Polychaetes often have relatively small non-yolky eggs, in correspondence with planktotrophic life, involving indirect development through a feeding larval stage (Rouse 1999; 2000). However, most polychaetes have life histories involving lecithotrophy (Haszprunar et al. 1995 and see Chapter 6). Many clitellates have relatively large, yolky eggs (see Chapters 8 and 9), correlating with their direct development from embryo to juvenile, without a distinct metamorphosis (Shankland and Savage 1997). The main components of the annelid egg can be categorized as lipid-rich, and protein-rich components of the yolk, an egg cortex, and some arrangement of clear, or yolk-deficient cytoplasm (see Chapter 2). In both polychaetes and clitellates, the egg cytoplasm undergoes rearrangement after fertilization, before the first cleavage division. These rearrangements can be accompanied by rather dramatic shape changes in the fertilized egg, and have been studied for their likely involvement in the distribution of maternal determinants to the blastomeres. The classical idea is that yolk-free cytoplasm is distributed such that it makes its way to the teloblast precursors of the D-quadrant, which are destined to produce the major ectodermal and mesodermal components of the segmental tissues (see section 5.3, below). This teloplasm is also thought of as the dorsal determinant, since its presence correlates with the dorsal structures of the embryo. This correlation has been shown experimentally as outlined below. In the polychaete Chaetopterus, the egg cortex has been shown to contain 90-95% of the mRNA in the oocyte (Jeffery and Wilson 1983). There is evidence that the cortex undergoes extensive rearrangement after fertilization (reviewed in Eckberg and Anderson 1995). This reorganization appears to be mediated principally by microtubules, as judged by its blockage by microtubule inhibitors. In glossiphoniid leeches, such as Theromyzon rude and Helobdella triserialis, the yolk-free cytoplasm accumulates after fertilization in three

96 Reproductive Biology and Phylogeny of Annelida regions during the completion of meiosis: the animal and vegetal poles (teloplasm) and surrounding the egg nucleus (perinuclear plasm). As in polychaetes, biochemical disruption of microtubules and microfilaments has implicated microtubules as the main mediators of ooplasmic rearrangement (Astrow et al. 1989). However, detailed study of the complex cytoarchitecture of the leech egg shows that microfilaments form an integral part of cortical structures and must have some important roles in localization of cytoplasmic constituents (Fernandez et al. 1998; Cantillana et al. 2000). In the oligochaete Tubifex, the yolk-free cytoplasm accumulates after fertilization at the animal and vegetal poles, and has been termed pole plasm (Shimizu, 1982). In contrast to Chaetopterus and the leeches, the rearrangement has been shown to be a microfilament-dependent, rather than microtubule-dependent, process (Shimizu 1995). Thus, it appears that even though cytoplasmic rearrangement occurs in a superficially similar manner across annelids, the cellular mechanisms of the process have been modified in the oligochaete lineage, with a basic role shifting from microtubules to microfilaments in the oligochaetes. One approach to studying the significance of maternal determinants has been to experimentally redistribute cytoplasmic constituents to see how differences in their spatial distribution affects development (reviewed in Reverberi 1971). As a recent example, in the polychaete Platynereis dumerilii, Dorresteijn and Eich (1991) centrifuged fertilized eggs before first cleavage. This stratified the cytoplasm into lipid droplets, yolk granules and clear cytoplasm. The stratification was random with respect to the first cleavage plane, so that cytoplasmic constituents were abnormally distributed between the first few blastomeres. The centrifugation did not affect the location of the cleavage planes with respect to the polar bodies. The most dramatic finding was that a small but significant percentage of the embryos developed a second body axis — these embryos the authors termed “Janus monsters” (Fig. 5.1). Typically these double axis embryos are joined at the ventral side, correlating with the long-standing idea outlined above that one of the first four blastomeres (the D quadrant) contains determinants for the dorsal side of the embryo (Shankland and Savage 1997, and see section 5.3 below). In the Janus monster, the dorsal determinants are presumably distributed between two of the four-cell stage blastomeres, which go on to form two dorsal quadrants capable of organizing two independent body axes. In the leeches studied to date, cell fate can also be altered by centrifugation of eggs. In general, those cells that that inherit the teloplasm take on the characteristics of the teloblasts, which are normally derived from the D cell at the four-cell stage (Astrow et al. 1987, reviewed in Weisblat and Huang 2001). These results, like those in polychaetes outlined above, support the classical notion that the cells that inherit this plasm are conferred “organizer” properties inducing the dorso-ventral axis.

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Fig. 5.1. Drawing of a “Janus monster” twinned embryo developed after centrifugation of the uncleaved zygote. The twins are joined at the ventral side and have conjoined nerve cords in the anterior. The section is cut behind the duplicated peristomial cirri. Reproduced with permission from Dorresteijn, A. W. C. and Eich, P. 1991. Roux’s Archive of Developmental Biology 200: 342-35, Fig. 12, published by Springer-Verlag.

To date, no studies have been done to directly identify exactly the maternal cytoplasmic constituents that determine embryonic patterning. However, using the candidate gene approach several gene products have been found to be present in the egg. Table 5.1 lists known maternally expressed genes. The small amount of data currently available on maternal mRNAs or proteins do not show any obvious localization within the teloplasm. It is certainly possible that some of the genes listed in Table 5.1 are involved in the establishment of basic embryonic polarity. However, factors migrating with the teloplasm, which might be critical to conferring organizer properties to the D-quadrant have yet to be observed.

5.3

CLEAVAGE

All annelids undergo a common stereotypic holoblastic cleavage program known as spiral cleavage (Anderson 1966a,b; see also chapter 8). This

Cell signaling Translational repression TF-ZF TF-ZF TF-ZF TF-b-HLH TF-HB TF-HB

Dm — Wnt family genes Dm — nanos Dm — hunchback Dm — hunchback Dm — hunchback Dm — twist Msx family Dm — proboscipedia Vert — Hox2

Hro-WNT-A Hro-nos Htr-LZF2 T-hb Cc-hb Hro-twi Le-msx CHv-Hox2 yes yes yes yes yes yes yes yes

no no yes yes yes ? ? ?

Maternal Maternal transcript protein n/a n/a Peri-nuclear Peri-nuclear ?????? ? ? ?

Protein localization

Huang et al. 2001 Kang et al. 2002 Iwasa et al. 2000 Shimizu and Savage 2002 Werbrock et al. 2001 Soto et al. 1997 Master et al. 1996 Peterson et al. 2000

Reference

Species designations: b-HLH, basic-helix-loop-helix; Hro or Le, Helobdella robusta ; Htr, H. triserialis (leeches); T, Tubifex hattai (oligochaete); Cc, Capitella capitata; CHv, Chaetopterus variopedatus (polychaetes); Dm, Drosophila melanogaster (insect). Abbreviations: HB, homeobox; TF, transcription factor; Vert, vertebrate; ZF, zinc finger

Protein type

Probable homologs

Gene

Table 5.1 Maternal gene expression in annelids

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pattern of early cleavage is shared with a number of other lophotrochozoan taxa, including nemerteans, mollusks, and sipunculans, and is characterized by cell divisions regular in orientation, that generate uniquely identifiable cells with stereotypical cell fates. We will first describe the nature of the spiral cleavage program, and then examine the results of experimental manipulations of cleavage stage embryos and also relevant molecular data. For both the first and second cleavages in a spiralian embryo, the orientation of the cleavage plane is along the animal-vegetal axis (Fig. 5.2B, C). The first two cleavages produce blastomeres called macromeres denoted A-D at the four cell stage (Fig. 5.2E). The four macromeres A, B, C and D represent the four quadrants of the embryo, and the descendants of each blastomere contribute to corresponding domains of the adult body. That is, the A quadrant gives rise to roughly the left side of the adult head, the C quadrant to the right, D to the dorsal, and B to the ventral. The name to which ‘spiral cleavage’ refers becomes apparent at the third division when a set of generally smaller cells, the micromeres, is produced (Fig. 5.2F, G). The micromeres arise from an asymmetric cell division of each macromere towards the animal pole. In this division, the cleavage plane is oriented in an oblique angle relative to the animal-vegetal axis (Fig. 5.2G). Furthermore, in the next division of the macromeres (the fourth division), the cleavage plane is also oriented obliquely but in the opposite orientation by 90o. As the macromeres produce successive micromeres, there is an alternation of orientation in the oblique cleavage plane from clockwise to counterclockwise (Fig. 5.2H). Detailed characterizations of cell cleavage patterns were described for a number of polychaete species at the turn of the twentieth century, including Nereis (Wilson 1892), Capitella (Eisig 1899), Arenicola (Child 1900), Amphitrite (Mead 1897), Podarke (Treadwell 1901), Polygordius (Soulier 1902), Scoloplos (Delsman 1916) Chaetopterus (Lillie 1906), the leech Clepsine (Whitman 1878) the oligochaete Tubifex (Penners 1922; 1924) and more recently for the leech Theromyzon tessulatum (Sandig and Dohle 1988), and an earthworm (Storey 1989). The shared cleavage program among spiralians is accompanied by many similarities in cell fates of both micromeres and macromeres. For example, larval eyes generally arise from two of the first quartet micromeres, 1a and 1c. In addition, the primary mesoderm arises from a single fourth quartet micromere, 4d. The prototrochal cells come from primary trochoblasts that arise from descendants of the first quartet micromeres (1q2), and posterior ectoderm arises from 2d. (Fig. 5.2) Fate maps have been described for a number of annelids (Fig. 5.6) and show the similar relative positions of presumptive areas in the blastula, as described in more detail in section 5.4. More recent studies have utilized intracellular injections to perform detailed cell lineages in clitellates such as Tubifex (Goto and Shimizu 1999) and a number of studies in leeches (Weisblat and Stent 1978; Weisblat and Shankland 1985; Shankland 1987a; Shankland 1987b; Huang et al. 2002). However, due to the small size of cells in most polychaete species, such cell lineage studies using intracellular fills have lagged behind those performed

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Colour Figure

Fig. 5.2. Annelids undergo a spiral cleavage program. A, B, C, D and G are lateral views with the animal pole at the top. E, F and H, I, J, and K are animal views. A. Uncleaved zygote. B. The first cleavage plane is oriented along the animal vegetal axis, and for many species is accompanied by the formation of a polar lobe. C. The size of the blastomeres at the two cell stage is different for species that demonstrate unequal cleavage. The larger is always the CD cell. D. Polar lobe formation is also visible in the second division associated with the division of the CD blastomere and shunts material into only one of the two daughter cells (the D cell). E. In unequal cleaving species at the four cell stage, the D blastomere (darker shading) is larger than the other three. F. The first spiral cleavage is at the third division and occurs when each of the four macromeres divides towards the animal pole in a clockwise (dextral) direction (arrow). G. Lateral view showing the oblique orientation of the cleavage spindle relative to the animal-vegetal axis for the first spiral cleavage in an eight cell stage embryo. H.16 cell stage embryo. The cleavage spindle is oriented in a sinistral (counterclockwise) (arrow) to produce the 16 cell stage. The cells of the D quadrant are shaded. I. 33 cell embryo of Capitella sp. I. J. Bright field view of a 4-cell stage Capitella sp. I embryo. K. Bright field view of a 4-cell stage embryo of Capitella sp. I that has been labeled with an anti-histone antibody to show the position of the condensed chromosomes (in brown), immediately prior to cleavage from the 4 to the 8 cell stage. The darker shading in E-I shows the descendants of the D quadrant. A, animal pole; V, vegetal pole. Original.

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in clitellates. Within the context of such striking similarities in final cell fates among cellular homologues, there are species-specific differences, and it is noteworthy that variation in cleavage programs and descendant fates can occur even within close relationships, as has been shown to be the case for variations in micromere fates between two leech Helobdella species (Huang et al. 2002). When viewed from the animal pole, the similarity of the cleavage patterns of the micromere descendants from each of the four macromeres, representing the four embryonic quadrants, is obvious (Fig. 5.2H). Spiralian embryologists have long noted patterns of cell arrangements in the micromere cap, particularly the presence of the molluscan and annelid “cross” at late cleavage stages. The annelid cross forms from descendants of the first and second quartet micromeres at the animal pole (Fig. 5.2). The presence or absence of a molluscan or annelid cross has often been used as a phylogenetic marker to align other spiralians with either mollusks or annelids. One note of caution is that within annelids, not all species form a robust annelid cross (such as Chaetopterus, Mead 1897), and it may be more realistic to consider that there is a range in the patterns created by the arrangement of micromeres rather than there being two discrete categories of cleavage patterns (see Maslakova et al. 2004 for discussion), making it less useful as a phylogenetic marker. Within the context of a shared spiralian cleavage program, the relative sizes of cells and timing of cleavages can vary among annelid species. The exact arrangement and pattern of cells in the micromere cap are influenced by these differences, which in turn are influenced by relative amounts of yolk in the embryo and equal vs. unequal cleavage programs. Differences in cleavage patterns among annelids can occur as early as the first cleavage, in which an equal or unequal cleavage division produces two blastomeres of the same or different sizes. This difference is maintained through successive divisions, and at the four cell stage in unequal cleaving embryos one of the four blastomeres is typically larger than the other three (Fig. 5.2E). The larger cell is always the CD blastomere at the two-cell stage and the D blastomere at the four cell stage. The relative sizes of blastomeres within an embryo at the two and four cell stage determine whether an embryo is referred to as an equal (equal sizes) or unequal (unequal sizes) cleaving species. There are both equal and unequally cleaving annelids, and a discussion of which cleavage program is ancestral for the group (see Freeman and Lundelius 1992; Dohle 1999) must be regarded as unresolved at present (see Chapter 1). The relative timing of cell divisions themselves may have phylogenetic signal, as has been shown by Linberg for gastropod mollusks (Guralnick and Linberg 2001), and a similar analysis might yield valuable insight into some of the enigmatic relationships among annelids. In unequal cleaving forms, the larger D blastomere gives rise to a distinct set of descendants compared with those of the A, B and C blastomeres. Additionally, the D quadrant is unique in that it has organizing properties in the early embryo, which means it provides instructional cues to the other

102 Reproductive Biology and Phylogeny of Annelida blastomeres (Clement 1962). The cell division pattern of the D quadrant is often distinct from that of the other quadrants. The large conspicuous teloblastic cells present in leeches and some other clitellates also arise as D quadrant descendents (see section 5.5.1). The mechanism and timing by which the D quadrant is specified to undergo a unique developmental program has been studied in several unequal-cleaving annelid embryos. In these embryos, specification is produced by differential inheritance of cytoplasm by the D blastomere and is thought to take place by the four-cell stage (reviewed in Verdonk and Biggelaar 1983). Cytoplasm can be partitioned to a particular blastomere by two mechanisms, either asymmetric positioning of the cleavage furrow, such is the case in Nereis (Wilson 1892), leech (Sandig and Dohle 1988) and Tubifex (Penners 1922), or production of a polar lobe. The polar lobe is a transient structure that forms during cytokinesis and results from cytoplasmic extrusions towards the vegetal pole of the embryo (Fig. 5.2B). The entire contents of the polar lobe are specifically shunted to only one of the daughter cells. Polar lobes can be observed during multiple cleavages and are especially obvious in the first two embryonic divisions. Polar lobes are formed in a number of spiralian groups, two of the best studied examples are in the gastropod mollusk Illyanasa (Clement 1952) and the scaphapod Dentilium (Verdonk and Biggelaar 1983). In polychaetes, the polar lobes of Chaetopterus and Sabellaria have been described in detail. Sabellaria has a prominent polar lobe and its contents have been experimentally shown to be critical for normal development of the larvae and in the specification of the D quadrant (Hatt 1932; Render 1983) (Fig. 5.3A-C). When the polar lobe of the first division is removed, the resulting larva lacks structures such as the apical tuft and lateral post-trochal chaetae. When the polar lobe from the second division is removed, head structures such as the apical tuft form normally but the post-trochal chaetae fail to form (Render 1983). Thus, the contents from the polar lobe from the second division have less morphogenetic potential than the contents of the polar lobe from the first division. In contrast, when the first polar lobe is removed from Chaetopterus embryos, development is essentially normal (Henry 1986) (Fig. 5.3D, E). Differential inheritance of cytoplasm in Chaetopterus is achieved both by an asymmetric cleavage furrow and by production of a polar lobe. In this case, asymmetric cleavage is more functionally relevant and the polar lobe of Chaetopterus is relatively small. Thus, cytoplasmic determinants essential for D quadrant specification are primarily localized to the CD blastomere through positioning of the cleavage spindle and not into the polar lobe. This has been demonstrated by removing a vegetal region of the cytoplasm with resulting loss of larval structures such as eyes and lateral hooked bristles in Chaetopterus (Henry 1986). The nature of the contents of the polar lobe has intrigued researchers for many years; however, this remains a mystery. Polar lobe contents have been examined from both morphological and biochemical perspectives (Weber 1958), and currently there is no report of utilization of a subtractive hybridization technique. Thus, the exact nature

Fig 5.3. Comparison of polar lobe removals between Chaetopterus and Sabellaria. Original. A. Sabellaria has a prominent polar lobe and a cleavage furrow that is symmetrically positioned within the embryo. The larva of Sabellaria has several features that can be scored including an apical tuft, prototrochal band and chaetae in the post-trochal region. B. Removal at the first division polar lobe results in a partial larva that has a prototrochal band but lacks an apical tuft and chaetae. C. Removal of the polar lobe at the second division results in larvae that forms a normal apical tuft but lacks chaetae. D. Chaetopterus embryos have small polar lobes and an asymmetric cleavage furrow. E. Removal of the first division polar lobe in Chaetopterus embryos results in an essentially normal larvae containing lateral hook bristles, eyes and an apical tuft. Dashed lines denote positions of removal of the polar lobes. Original.

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104 Reproductive Biology and Phylogeny of Annelida of substances that function in the organizing properties of the D quadrant remain unknown. Equal cleaving species are less well studied in annelids than unequal cleaving forms and clitellates have only unequal cleaving forms, with perhaps one minor exception (Freeman and Lundelius 1992). In equal cleaving annelid embryos such as Hydroides (Wisely 1958), all four quadrants appear identical to one another and the determination of bilateral symmetry (or establishment of the secondary or dorsal-ventral axis) does not occur until a later stage of development following production of the 4th quartet micromeres (the 64-cell stage) (see Freeman and Lundelius 1992). The first embryonic axis, the animal-vegetal axis, is determined during oogenesis (see above). Bilateral symmetry is first manifest by an asymmetric division of 2d2, whose daughter cells have distinct relative sizes of their counterparts from the A, B and C quadrants. Prior to this, the quadralateral symmetry of the embryo is especially visible in an animal pole view, in which the presence of corresponding micromeres and their spatial arrangement reflect cell division activity from the four underlying macromeres (Fig. 5.2H). In mollusks, bilateral symmetry appears as the result of movement of the 3D macromere to an interior position, prior to the birth of the fourth quartet. The appearance of bilateral symmetry is closely tied to the specification of the D quadrant in equal cleaving forms. Compared with mollusks, there is very little information known about the exact mechanism of cell fate specification for equal cleaving annelids. In mollusks, the D macromere is specified as the result of a signal from the overlying first quartet micromeres where there is persistent cell-cell contact with one of the macromeres. Although D quadrant specification in equal cleaving annelids is likely to be occurring via a similar mechanism, this has not been directly experimentally examined. The developmental potential of blastomeres has been investigated by isolation of individual blastomeres at early cleavage stages, which are raised to trochophore stages, and then examined for presence of identifiable differentiated structures (Fig. 5.4). In addition, a number of studies have also reconstituted various combinations of blastomeres (i.e. C plus A), to examine possible inductive interactions between cells and their descendants. The results of these studies differ between equal and unequal cleaving embryos since blastomere differences are apparent at a much earlier stage for unequal cleaving forms. For example, in an early experiment performed by Wilson (1904) on Lanice, blastomeres isolated at the two cell stage showed a mosaic type of development in an unequal cleaving polychaete. In this case, the AB cell yielded a partial larva (containing an apical organ but no eyes nor post-trochal region), while the CD blastomere gave rise to a reasonably normal larva containing a prototroch, apical organ, eye and post-trochal region; thus the two cells have different developmental potential. Blastomere isolations and recombinations have been performed in several polychaetes including the

Fig. 5.4. Blastomere isolations during early stages of development in Sabellaria. A. Blastomeres separated at the two cell stage result in larvae with distinct morphological features. Larvae that develop from the CD blastomere have bilateral symmetry, prototrochal cilia, chaetae, an apical tuft, but lack apical cilia. Larvae resulting from AB isolates have prototrochal and apical cilia. B. Blastomeres isolated at the four cell stage result in three different larval morphologies. A and B blastomere isolations result in larvae with the same morphology which are partial larvae with prototrochal and apical cilia. C blastomere isolates result in partial larvae with prototrochal cilia and apical tufts. Larvae arising from isolated D blastomeres result in animals with bilaterally symmetric chaetae and prototrochal cilia. Dashed lines denote position of separation of blastomeres. Original.

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106 Reproductive Biology and Phylogeny of Annelida unequal cleaving Chaetopterus (Tyler 1930; Henry 1986), Sabellaria (Fig. 5.4) (Novikoff 1936; Render 1983), and Nereis (Costello 1945). In general, for isolated blastomeres, only embryos containing derivatives of the D quadrant macromere can form larvae that are bilaterally symmetric, have a dorsal ventral axis, and form mesodermal structures. These results underlie the importance of the ‘organizing’ properties of the D quadrant. Chaetopterus larvae raised from isolated blastomeres not containing a D quadrant, for example A or B, form an apical tuft, but do not generate eyes, bristles or endoderm (Henry 1986). What is clear is that 3D and its derivatives are always necessary for normal development. Additionally, a complex combination of influences determines cell fates in annelid embryos, even within a single embryo and the particular cellular mechanism can vary from cleavage to cleavage and cell to cell. Unequal cleaving embryos can be artificially forced to undergo an equal first division by gently compressing the eggs between two plates. Various manipulations leading to equalization of the first cleavage division have been performed in Chaetopterus (Titlebaum 1928; Tyler 1930; Henry and Martindale 1987), Nereis (Wilson 1896; Morgan 1910b), and Platynereis (Dorresteijn 1987) and in all cases the eggs develop into larvae with many duplicated structures. Thus, by forcing equal amounts of cytoplasm into the two daughter cells, the developmental potential is equalized to both halves of the embryo, and appears to result in the formation of two D quadrants. The results of these experiments are similar to those found when egg cytoplasm is equally distributed by centrifugation prior to first cleavage as described in section 5.2 (see Fig. 5.1 — Janus monster). The exact molecular nature of the cell signaling events by which the D quadrant is specified in equal cleaving embryos is largely unknown. However, recent experiments on mollusks and the serpulid polychaete Hydroides by Lambert and Nagy (2003) have shown a strong spatial and temporal correlation between the specification of the D quadrant and activation of the MAP Kinase (mitogen-activated protein kinase) signal transduction pathway (Fig. 5.5). The Erk 1/2 family of MAP Kinases are cytoplasmic components of a signal transduction pathway that are anchored by a scaffolding protein to the cytoplasmic face of the cell membrane and transduce extracellular signals in the responding cell through a series of phosphorylation events (Ferrell 1996). The presence of activated MAP Kinase is indicative that a cell is receiving a signal. In the equal cleaving Hydroides, MAP Kinase is activated specifically in only a single cell of the embryo, the micromere 4d (Lambert and Nagy 2003) (Fig. 5.5). Detection of activated MAP Kinase in 4d is delayed relative to its birth and coincides with the division of 2d2. This temporally corresponds to the appearance of bilateral symmetry in equal cleaving polychaetes. Thus, although the exact molecular identity of the signal itself that specifies the D quadrant (4d) remains unknown, it is likely that MAP Kinases are involved in transducing the signal from the cell surface into the cytoplasm.

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Activation of MAP Kinase in 4d of H. elegans

Fig. 5.5. The activated form of the signaling molecule MAP Kinase appears in a single cell (4d) in the H. elegans embryo after the birth of the 4th quartet micromeres. In this equal cleaving embryo, activation of MAP Kinase in one quadrant of the embryo coincides with the specification of the D quadrant. A. The dark reaction product marks the presence of activated MAP Kinase. Bright field. B. Same embryo with cell boundaries drawn. Vegetal view. From Lambert, J. D. and Nagy, L. M. 2003. Developmental Biology 263: 231-41. Fig. 7.

In annelids, only a small number of developmental regulatory genes to date are reported to be expressed during early cleavage stages and most have only been characterized in the leech embryo. These genes include the transcription factors Hr-nos (Pilon and Weisblat 1997), Le-msx (Master et al. 1996), Hb (Savage and Shankland 1996a), and the secreted signaling molecule Hr-WntA (Huang et al. 2001). In general, these genes are broadly expressed in all or most blastomeres during early cleavage stages (and often are additionally expressed at later stages in association with morphogenesis of particular processes). This is the case for msx, a homeobox-containing transcription factor, which is expressed in all blastomeres during early cleavage stages that inherit maternally deposited teloplasm (Huang et al. 2001). However, nanos (Hr-nos) expression shows differences in levels of protein among blastomeres and is expressed at higher levels in one D blastomere descendant, DNOPQ than another, DM. Since DNOPQ gives rise to the segmental ectoderm of the body and DM to the segmental mesoderm, it has been proposed that this difference may suggest a role for Hr-nos in the segregation between mesodermal and ectodermal fates, although this has not yet been functionally demonstrated. In the case of hunchback (Hb), RNA and protein expression has been examined in the leech embryo, and immunohistologically in both the oligochaete Tubifex and in the polychaete Capitella. Hb immuno-crossreactivity is present in all blastomeres during early cleavage stages in Capitella (Werbrock et al. 2001) and Tubifex (Shimizu and Savage 2002), but in the leech embryo, Hb

108 Reproductive Biology and Phylogeny of Annelida immuno-crossreactivity is localized to micromere cells (Iwasa et al. 2000) and is absent from the macromeres, potentially reflecting a molecular distinction between these two cell types. The Wnt genes are a family of secreted signaling molecules that have been implicated in a number of distinct developmental processes across metazoan species and there are at least eight members in the Wnt gene family in lophotrochozoans (Prud’homme et al. 2002). The gene expression of one Wnt gene family member, WNT-A, shows a dynamic and restricted expression pattern during early cleavage stages in the leech embryo (Huang et al. 2001). During early stages of the cell cycle of the 2 cell stage, WNT-A shows a stochastic pattern of expression: approximately half of the embryos show staining in only the larger CD blastomere and the other half in the AB cell. Immediately following this stochastic phase, localization becomes stabilized in 100% of the embryos and is initially restricted to the AB cell and then later in the cell cycle to the CD cell. When the two blastomeres are separated, WNT-A is expressed in both cells until each cell divides. The authors interpret this dynamic expression as the result of negative regulation through cell-cell contact that stabilizes the expression pattern. It is unclear what role this cell-cell signaling has in an embryo at a stage when there are cytoplasmic differences between AB and CD that probably account for the differences in identity between these two cells. It is possible that such dynamic WNT-A expression represents an evolutionary relic from an equal cleaving ancestor. The fact that in general the genes expressed during cleavage described to date are broadly expressed, suggests that there are still many molecular factors to be described that distinguish individual blastomeres. However, molecular studies of cleavage stages in annelids is in its infancy and the future promises some interesting results since current techniques will make it possible to examine the molecular basis of a number of fundamental questions in spiralian embryology, some of which were raised over a hundred years ago.

5.4 GASTRULATION AND GUT DEVELOPMENT For the purposes of this paper, gastrulation will be defined as the movement of cells to their organ-forming positions, and the associated formation of the three definitive layers and the primitive gut (Anderson 1973). This process must involve both the specification of cell fate and mechanisms for the proper spatial rearrangement of those cells. These two general aspects of gastrulation are probably highly interrelated, so that cell specification may be occurring as cells move, and cells may be moving because of events related to previously specified fates. We will first describe the general character of gastrulation in the various annelid groups and then look at molecular data that might pertain to gastrulation processes. In both polychaetes and clitellates the relative positions of presumptive areas of the blastula formed during cleavage are similar in all species (reviewed in Anderson 1973). However, the details of

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the fate map vary widely depending on the amount of yolk in the blastomeres, and whether development is indirect, as in some polychaetes, or direct, as in many polychaetes and all clitellates. Representative fate map diagrams are shown in Fig. 5.6. It has long been accepted that a common feature of annelid gastrulation is the generation of the segmental mesoderm and ectoderm by teloblastic growth (Brusca and Brusca 1990). Teloblasts are large stem cells producing a lineage of segmental founder cells (blast cells), which then give rise to segmentally reiterated progeny. The teloblasts are thought to reside in a posterior growth zone that generates tissue anteriorly. While this pattern is certainly well documented for clitellates, there is no definitive modern evidence for the presence of posteriorly located teloblastic cells in polychaetes, although their developmental patterns may be consistent with teloblastic growth. For the purposes of this review, we will present a summary of the classic view of teloblast/growth-zone-based gastrulation in polychaetes, with reservations and counter-evidence as presented in section 5.5 below. With the above in mind, in described species, there are several bilateral pairs of ectoteloblasts (four in glossiphoniid leeches), and one bilateral pair of mesoteloblasts, all derived from the D quadrant. The more or less large yolky prospective midgut cells are derived from the vegetal macromeres. The stomodeum is derived from the first or second quartet micromeres, commonly the 2b cell, while other anterior ectoderm, apart from that to be derived from the ectoteloblasts, comes from other micromeres of the first to third quartet (Fig. 5.6). Since polychaetes develop indirectly, through a trochophore larval stage, their blastula fate map and subsequent gastrulation movements include larval structures, especially the prototroch, not present in clitellates. In addition, it has been argued that polychaetes, but not clitellates, have ectomesoderm, derived from second or third quartet micromeres (Anderson 1966b; 1973).

Fig. 5.6. Diagrammatic blastula fate maps, drawn in left lateral view. A. The polychaete Scoloplos. B. The oligochaete Tubifex. From Anderson, D. T. 1973. Embryology and Phylogeny in Annelids and Arthropods. Pergamon, Oxford, Figs. 6b, 29a.

110 Reproductive Biology and Phylogeny of Annelida In the process of annelid gastrulation, varying combinations of invagination and epiboly are responsible for the cell movements that result in the arrangement of the gastrula. The relative extent of these two processes has been argued to depend, as in the arrangement of the fate map, mostly on the size of the various blastomeres, which is related to the amount of yolk in the embryos of different species (Okada 1957; Anderson 1973). This principle applies in both polychaete and clitellate gastrulation, with the proviso as above about the necessity to produce larval structures in the polychaetes. What follows is a generalization of annelid gastrulation. For more extensive reviews refer to Okada (1957), Anderson (1973), or the detailed modern papers on leech gastrulation (reviewed in Weisblat and Huang 2001). In polychaetes and clitellates (e.g., Lumbricus) with less yolky embryos that have coeloblastulae, gastrulation starts with invagination of the presumptive midgut cells (Fig. 5.7B). Larval ectoderm in polychaetes, or

Fig. 5.7. Diagrams of gastrulation in a representative polychaete and clitellate. A. Left lateral view, and B, slightly parasagittal section of gastrula of the polychaete Scoloplos. C. Development of the germinal bands in the leech Glossiphonia in posterodorsal view. D. Left lateral view of gastrulating embryo of the oligochaete Tubifex. Abbreviations: MI, left mesoteloblast; Mr, right mesoteloblast. From Anderson, D. T. 1973. Embryology and Phylogeny in Annelids and Arthropods. Pergamon, Oxford, Figs. 8d, 9c, 31d, 31e.

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provisional epithelium in clitellates, both derived from the animal micromere cap grows over the embryo towards the ventral side. The mesoteloblasts assume a posterior position in the blastocoel. Ectoteloblasts wind up located superficial to the mesoteloblasts just under the larval ectoderm or provisional epithelium. The blastopore generally becomes elongated in an anteroposterior direction with the posterior part closing off. The anterior opening becomes the mouth, and the stomodeal cells continue to invaginate to form the foregut. Later the posterior part of the blastopore reopens as the anus. For the polychaetes and most clitellates that have more yolky embryos, stereoblastula formation and gastrulation is characterized by epiboly rather than invagination (Fig. 5.7C). The micromere cap grows over the presumptive midgut cells and teloblasts. In clitellates, eventually the teloblasts begin producing the germinal bands, which will generate most of the segmental adult body. The posteriorly-located teloblasts produce segmental founder cells in an anterior to posterior sequence. The clonal progeny of these segmental founders become the germinal bands, which align along the ventral midline and grow dorsally to replace the micromere cap-derived provisional epithelium and become the adult body wall. The blastopore can be looked at as the point where the germinal bands meet to become the germinal plate, at the location of the presumptive stomodeum. The germinal bands then “zipper up” from anterior to posterior to cover the endodermal blastomeres (Fig. 5.7D). In looking for the molecular basis of cell fate and gastrulation, there have been two major approaches. First, cellular-level experiments have been done in the attempt to determine when cell fates are determined. Second, a candidate gene approach has been applied to look for annelid homologs of genes known to be involved in early embryonic patterning events in other groups, especially fruit flies (Table 5.2). As previously discussed, the distribution of cytoplasmic constituents during cleavage has been shown to be a major determinant of cell fate. Nelson and Weisblat (1991; 1992) performed experiments in the leech Helobdella which removed either vegetal or animal teloplasm before first cleavage. When animal teloplasm was removed the ectodermal teloblast mother cell (DNOPQ) assumed a fate similar to that of the mesodermal teloblasts, even though some vegetal teloplasm had relocated to the animal hemisphere. The ectodermal fate could be rescued by moving some of the vegetal teloplasm by centrifugation so that it contacted the cortex near the animal pole. These experiments confirmed earlier results indicating that animal and vegetal teloplasms are equivalent in their potentials, and suggest that it is the animal cortex interacting with teloplasm that distinguishes between ectodermal and mesodermal cell fates. We are not aware of experiments targeting the fate determination processes behind the endoderm or non-teloblastic ectoderm and mesoderm. It seems likely that these cell types are also determined by the inheritance of certain cytoplasm or cortex. However, it is also possible that cell-cell

112 Reproductive Biology and Phylogeny of Annelida signaling from other cells, such as the D-quadrant, is necessary for fate specification in these lineages. This case would be difficult to distinguish from the cell autonomous case, since the invariability of the spiral cleavage pattern would result in the same cell contacts regardless of whether fate were determined autonomously or by signaling. Using the candidate gene approach, researchers have found annelid homologs of many genes associated with gastrulation in flies and mice (listed in Table 5.2). In the leech, since ectodermal teloblast fate has been shown to be determined by animal teloplasm and cortex, one would expect to find genes differentially expressed in ectodermal precursor cells. Hro-nos, the leech homolog of nanos in Drosophila conforms to this expectation. As in flies, Hro-nos is a maternal transcript. Unlike the posterior localization of nanos in flies, however, Hro-nos is distributed throughout the uncleaved zygote. During early cleavage transcripts migrate to the teloplasms, eventually winding up distributed to the DNOPQ ectoteloblast precursor cell at fourth cleavage (Pilon and Weisblat 1997; Kang et al. 2002). Thus its distribution is associated with the cellular constituents shown to determine ectodermal cell fate in the segmental tissues of the body. Its differential distribution has been likened to the gradient of nanos protein in the fly embryo, which is involved in repressing translation of the gap gene hunchback, and sets up initial regionalization along the anterior-posterior axis. In the leech however, Hro-nos distribution has not been associated with any axial gradient, but rather with the determination of ectodermal cell fates (or possibly, more generally, ectodermal germinal band cell fates). Homologs of the gap gene hunchback have been cloned and characterized across several major annelid lineages. In the leech, Htr-LZF2 protein is present in the provisional epithelium and prostomium, but not in the germinal bands (Iwasa et al. 2000). On the other hand, Htr-LZF2 transcripts are detected in germinal band cells and their precursors, in addition to the prostomium and the provisional integument (Savage and Shankland 1996b). This situation is consistent with the notion that leech nanos and hunchback are interacting analogously to flies, with nanos protein inhibiting hunchback translation in the germinal band, presumably maintaining or establishing differential cell fates. The expression pattern in the polychaete Capitella is similar, with Cc-Hb protein detected mainly in the larval ectoderm, but apparently not in tissues attributed to teloblastic origins (Werbrock et al. 2001). However, in the clitellate Tubifex, T-hb protein is present in a subset of the ectodermal teloblasts (Shimizu and Savage 2002), suggesting that somewhere in the clitellate lineage the exclusion of hunchback from teloblasts was relaxed, presumably associated with other changes in the pathways of cell fate and pattern formation. A key aspect of gastrulation is mesoderm determination. The dorsal/twist/ snail pathway is associated with mesoderm determination in flies, and a homolog of snail is a marker for mesoderm in vertebrates. In flies dorsal protein is preferentially transported into ventral cell nuclei, and activates snail and twist expression in ventral cells. Snail and twist are also required

TF-ZF TF-HB

Hro-SNA Pd-gsc

Dm — hunchback

NK-2 — flies and vertebrates

Htr-LZF2 T-hb

Cc-hb

Lox-10

TF-HB

TF-ZF

TF-ZF TF-ZF

TF-T-box

Nardelli-Haefliger and Shankland 1993

Werbrock et al. 2001

Iwasa et al. 2000 Shimizu and Savage 2002

Arendt et al. 2001

Goldstein et al. 2001 Arendt et al. 2001

Goldstein et al. 2001

Soto et al. 1997

Arendt et al. 2001

Bruce and Shankland 1998

Kang et al. 2003

Master et al. 1996 Kang et al. 2002

Reference

Species designations: same as Table 5.1 and also, Pd, Platynereis dumerilii (polychaete). Abbreviations: same as Table 5.1 and also, rel, rel-domain

Pd-bra

Dm — snail Dm — goosecoid other gsc genes Dm — brachyenteron Mm — T/Brachyury Dm — hunchback Dm — hunchback

Early in margins of blastopore, later confined to posterior stomodeum and proctodeum (anus), prospective mesoderm Embryonic epithelium and CNS (not in teloblasts) Embryonic epithelium and subset of ectodermal teloblasts (one side only at any time point) All cells during gastrulation, then trochal regions, stomodeum, midgut, CNS Prostomial epidermis and supraesophogeal ganglion, later in endoderm prefiguring segmental restrictions of gut

Early in micromeres and blast cells, later in segmental repeats in germinal plate (not in CNS) Same as DL but later Trochophore in stomodeum

TF-b-HLH

TF-HB

TF-HB

TF-rel

Hro-hh

Dm — hedgehog Vertebrates - sonic hedgehog and others Htr-Lox22 Dm — orthodenticle other Otx genes Pd-otx Dm — orthodenticle other Otx genes Hro-twi Dm — twist (required for snail transcription) Hro-DL Dm — dorsal

Expression pattern Throughout germinal bands, precursors and descendants Early inherited mostly by DNOPQ cell, later segmentally reiterated in germ cell precursors Early in micromere-derived anterior germinal plate — later in foregut, posterior midgut, and at lower level in segmentally iterated pattern in germinal plate — later yet in hindgut only Early (st.8) in anterior dorsal surface ectoderm and foregut — later around mouth, in proboscis and CNS Trochophore in oral region and apical NS, anterior and posterior margins of peristomium; with prototroch then metatroch Present through gastrulation but localization unknown

Msx family Dm — nanos

Le-msx Hro-nos

Protein type

TF-HB Translational repression Cell signalling

Probable homologs

Gene

Table 5.2 Genes with possible roles in gastrulation or primary cell fate determination

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114 Reproductive Biology and Phylogeny of Annelida for the cell movements at the initiation of gastrulation in flies (Leptin et al. 1992), with snail expression occurring in involuting or invaginating cells in both insects and vertebrates (reviewed in Arendt and Nübler-Jung 1997; Hemavathy et al. 2000). From the apparent functional similarity of the snail pathway in both ecdysozoans (flies) and deuterostomes, one might expect that these genes would be involved in dorsoventral patterning and/or mesoderm specification in annelids as well. However, the expression data in leeches suggests a different role for these genes. Hro-DL protein is observed in all micromeres during gastrulation, and then in all the blast cells, but not in their parent teloblasts. Later, during segmentation, dorsal appears in segmentally iterated stripes along the anteroposterior axis (Goldstein et al. 2001). The expression of Hro-SNA protein exhibits a similar pattern but delayed somewhat compared with the onset of Hro-DL expression, consistent with the idea that dorsal is an activator of snail (Goldstein et al. 2001). Hro-twi, a maternal transcript (it is zygotic in flies), is present throughout gastrulation, but no localization data has been reported (Soto et al. 1997). Thus, no dorsoventral gradient, association with specifically ventral cell fates, or specifically mesodermal expression is seen, as would be expected from the fly or vertebrate data. Rather, in the leech, dorsal and snail must have some other function, as yet undetermined, during gastrulation in the micromeres and blast cells. In later development, the genes appear to be associated with cell diversification within segmental primordia. It is possible that dorsal and/or snail homologs in polychaetes might have some dorsoventral patterning or mesoderm specifying roles, if indeed this pathway is ancestral to bilaterians. It this were the case, the leech condition would be derived, and might be associated with the derived gastrulation patterns in clitellates. Arendt and co-workers have examined the expression patterns of three genes in the polychaete Platynereis dumerilii that are associated with gastrulation in other groups, Otx, brachyury, and goosecoid. Pd-otx is expressed in stomodeal cells, and in the prototrochal bands associated with the peristomium, the mouth-containing region of the head. Pd-otx transcripts are also seen at early larval stages in apical neurons and sensory cells (Arendt et al. 2001). Pd-otx expression prior to gastrulation was not reported. The expression in ciliary cells of the oral region is similar to the pattern seen in basal deuterostome larvae, while the expression in the anterior nervous system is present in many protostomes and deuterostomes. The stomodeal expression has not been reported elsewhere, to our knowledge. Otx expression has also been examined in the leech Helobella (Bruce and Shankland 1998). Here, Htr-Lox22 (Otx homolog) transcripts are detected during gastrulation in a ring at the dorsal side of the prostomium, which later demarcates the boundary between prostomial and foregut tissues. Early in foregut formation there is also strong expression in the stomodeum. Ht-Lox22 is also expressed in certain cells of the anterior and segmental CNS, but in leeches the expression is not as general a marker of anterior CNS as are Otx homologs in vertebrates and insects. Thus Otx may

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be an important player in the fate of stomodeal cells in annelids, and in demarcating the extent of the “head” region. Bruce and Shankland (1998) go on to argue that leeches have a radial organization of structures around the mouth, which is outlined by Htr-Lox22 expression, and may be a vestige of the derivation of the bilaterians from a radially organized ancestor. In Platynereis, this concentric pattern is not apparent at the stages examined in Arendt et al. (2001) and if anteroposterior elongation is indeed primitive for the protostome blastopore (Arendt and Nübler-Jung 1997), it may be that the radial oral symmetry of leeches is a derived condition. The polychaete homolog of the deuterostome gene brachyury is also expressed in the stomodeum, and more generally around the elongated blastopore, including the region of the proctodeum (prospective anus), as the gut forms. Pd-bra is restricted to the ventroposterior part of the stomodeum. Eventually expression is lost in the ventral midline cells, resolving to the ventral stomodeum and the proctodeum. This expression reappears later on the ventral midline of the midgut. Finally, the Pd-bra probe labels cells attributed as prospective posterior mesoderm (Arendt et al. 2001). Pd-gsc (Platynereis goosecoid homolog) is expressed at the anterior edge of the blastopore, destined for the stomodeum, in the early larva, and this stomodeal expression persists in later stages (Arendt et al. 2001). This pattern recalls the presence of goosecoid expression as a general marker for foregut roof in postgastrula vertebrate embryos, and in the larval foregut and nervous system in Drosophila. Thus, Otx, brachyury, and goosecoid, appear to be involved in the early specification of anterior structures of the gastrula. Otx has been widely viewed as one of the earliest markers of the most anterior CNS, from patterns seen in vertebrates and flies. In Platynereis and Helobdella, this neural function appears less prominent, Otx being strongly expressed in the stomodeum/foregut, another pattern widely conserved in bilaterians. The boundary of Otx expression demarcating head and trunk in the surface ectoderm appears to be a more unique attribute of the pattern in annelids vs other groups. Brachyury is expressed in four tissues at gastrulation in Platynereis, the stomodeum/foregut, proctodeum/hindgut, a set of ventral midline cells (which may wind up in the midgut later), and cells attributed as prospective visceral mesoderm. Notably, the fore and hindgut expression is similar to that seen in hemichordate and echinoderm (basal deuterostome) larvae. The Drosophila homolog brachyenteron is expressed in the hindgut, but not the foregut. In addition, it has been shown to have a role in the visceral mesoderm. In vertebrates, brachyury function has been shown to have a key role in axial mesoderm formation, and in basic gastrulation movements, but not in fore- and hindgut specification (Conlon and Smith 1999). Taken together, the data suggests that brachyury had two roles in the bilaterian ancestor, specification of prospective fore- and hindgut cells, and a subset of prospective mesoderm cells, and that polychaetes have retained these primitive functions, at least in part.

116 Reproductive Biology and Phylogeny of Annelida Hedgehog is a secreted signalling molecule known to have crucial functions in regulating segmental polarity in flies, and many patterning functions in vertebrates, including the induction of floor plate in the brain, and signaling polarity in the limb (Fietz et al. 1994). Hedgehog has been cloned from Helobdella and here its major roles appear to be in gut formation and specification of prospective gonadal tissue, but does not appear to have the segmental polarity role one might expect from the data in flies (Kang et al. 2003). In Helobdella it is first expressed in early gastrulation at the site of the future oral opening, and as gastrulation proceeds expression spreads throughout the proboscis, and also appears in the midgut and segmentally iterated cells of the germinal band. To test for function, cyclopamine was used to specifically block the hedgehog signalling pathway. This treatment resulted in severe disruption of development of the proboscis and crop, probably due to failure of the visceral mesoderm to form properly. In addition, gonads and coelomic mesenchyme failed to form. Thus, hedgehog signalling in the leech appears to be primarily required to properly specify a subset of the mesodermal cells required for gut, gonad, and mesenchyme formation. Homeobox genes have also been found to be expressed in the gut. Both the Lox3 cluster genes (3 tandem duplicates), of the Xlox class (WysockaDiller et al. 1995), and Lox10, an NK-2 gene (Nardelli-Haefliger and Shankland 1993) are expressed in segmentally iterated patterns in the midgut rudiment which prefigure the development of the segmental gut diverticula. Nardelli-Haefliger and Shankland (1993) show that the expression of Lox10 is dependent on the adjacent mesoderm, since ablation of mesoderm abolishes Lox10 expression and subsequent gut morphogenesis.

5.5

SEGMENTATION

Annelids are commonly referred to as the ‘segmented worms’, reflecting the view that this characteristic of their body plan is a core feature of the annelid body plan. The presence or absence of segmentation has been used as a disproportionately heavily weighted character for inclusion of species within the Annelida, although recent molecular analyses include some unsegmented forms (for example, see McHugh 1997). It is clear however, that possession of a segmented body plan is plesiomorphic for annelids. Within the context of a segmented body plan, there is a diversity of forms, from highly tagmatized to homonomous body plans, and a broad range in the number of body segments. Some species have an exact number of segments, such as in leeches, while others have an indeterminate number. A distinctive feature of annelids is their ability to generate segments at multiple life history stages as well as their ability to replace segments during regeneration. In this chapter, we will focus primarily on segment addition during development. Most annelids generate their first segments during larval life, often in a limited number, and the majority are added during the juvenile phase in a

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sequential fashion from anterior to posterior from a posterior growth zone. It has been proposed that the first larval segments are formed in a manner independent from those after metamorphosis. Iwanoff (1928) pushed this distinction one step further by saying that in serpulid polychaetes, the mesoderm of the larval segments arises from a separate ontological source compared with the mesoderm of post-larval segments, which originate from ecto-mesodermal origins in the posterior growth zone. In many species the first larval segments appear simultaneously or in a very narrow time window and become morphologically obvious as the chaetae protrude from the body wall. It has been noted that there is variation among species in whether the ectoderm or the mesoderm shows morphological segmentation earliest (see Anderson 1973). For example, serpulids, nereidids, tomopterids and eunicids show precocious morphological delineation of segments of the ectoderm, whereas in other groups the underlying mesoderm breaks into somitic blocks prior to ectodermal segmentation. This raises an interesting question of which germ layer drives segmentation in distinct species and is consistent with the possibility that there may be variation in which germ layer controls the formation of larval segments. In many clitellates and a few direct developing polychaetes, segments are generated during embryogenesis. The cellular origin of the segmental tissue in annelids is derived entirely from D quadrant descendants in the embryo. The segmented mesoderm arises from descendants of the 4d micromere and the ectoderm from 2d descendants. At the cellular level, the most detailed information about how segments are generated during development is known for clitellates, specifically in leeches, Tubifex (reviewed in Arai and Shimizu 2001), and the earthworm Eisenia (Storey 1989). Eisenia and Tubifex generate a variable number of segments and can regenerate lost segments. In contrast, leeches have a set number of segments and very poor regenerative capacities. Interestingly, the embryonic origin of segmental tissues is very similar among these species and even includes commonalities at the single cell level. Detailed cell lineages have been performed and reveal that the segmented tissues arise in a highly stereotypical manner and originate as a product of the divisions of the large teloblasts described in Sec. 5.4 above. There are four ectodermal teloblasts (N, O, P, Q) and one meso-teloblast (M) in each hemisegment. The teloblasts arise during cleavage as descendants of the D quadrant; M from 4d and N, O, P, and Q as a descendants of 2d. NOPQ is a teloblast precursor cell, which undergoes sequential divisions to give rise to N, O, P and finally Q, respectively. Each teloblast undergoes serially repeated highly asymmetric divisions and produces a single chain or bandlet of progeny called primary blast cells (Fig. 5.8). In the N and Q lineages, two types of primary blast cells are generated and each is born alternate to the other. In the M, O and P lineages only one type of primary blast cell is produced. Each primary blast cell also undergoes a stereotypic program of cell divisions that are uniquely identifiable for each lineage. The bandlets from each ecto-teloblast spatially come together to form the

118 Reproductive Biology and Phylogeny of Annelida

Fig. 5.8. Ablation of individual cells in the segment primordia of the ectoderm of the leech embryo reveals mosaic development. Each primary blast cell arising from the O teloblast (I) undergoes a stereotypical cleavage program (II) and gives rise to a predictable set of descendant cells (III), including nervous system elements, epidermis and part of the nephridia. A. Unoperated control showing the full complement of descendant cells which arise from a single O primary blast cell. The filled nucleus of one of the descendants of the primary blast cell denotes the single cell (called the O. aap cell, a daughter of the O. aa cell) that expresses the segment polarity gene engrailed. B. When the o.aa cell is ablated, the resulting clone of differentiated cells is missing the descendants (III) that normally arise from o.aa. Wavy arrow denotes laser ablation of the single cell o.aa. In ablations of the O lineage, there is loss of structures but no regulation to compensate for loss of descendents of the ablated cell. Original.

germinal band with the M bandlet situated directly beneath. At a slightly later stage, the two germinal bands from each side of the embryo merge to form the germinal plate. Soon after the bandlets form the germinal band, the primary blast cells begin to divide and ultimately produce a descendant clone of approximately 70 differentiated cells (Fig. 5.8), many of which can be identified as individual cells. Each teloblast lineage contributes to every segment. A single primary blast cell from the O, P, and M teloblasts contributes the complement of a single segment of descendant cells, while in the N and Q lineages, the descendants of the two distinct primary blast cells give rise to an equivalent of a single segment complement of descendants. One experimental advantage of the large clitellate embryos is that they are tractable by microinjection. Single cells can be labeled with intracellular tracers and followed through development to the stage when

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morphologically differentiated cell types are present. In addition to being able to describe fate maps at a precise level of resolution, it has been possible to ablate individual cells or groups of cells to investigate the relative contribution of mosaic patterning and cell-cell interactions in establishment of the segmented body plan. The identity of primary blast cells is determined in a cell autonomous manner by their teloblast of origin, except for O and P (Weisblat and Blair 1984). In the case of the O and P bandlets, the identity of primary blast cells is determined after their birth by interactions between the bandlets. If the P bandlet is ablated by killing the P teloblast, the O bandlet transfates to P. In addition, signaling from the Q bandlet is necessary for O bandlet identity (Huang and Weisblat 1996). Experimental manipulations have been performed to ascertain whether cell-cell interactions are critical for patterning along the anterior-posterior axis in the establishment of segmental units. In arthropods, notably Drosophila, establishment of segmental unit boundaries is the result of signaling between spatially opposed rows of cells. When individual primary blast cells are physically isolated from either their anterior, posterior, or both adjacent neighbors in either the O or P lineage by single cell ablations with a laser microbeam, they develop normally (Seaver and Shankland 2000). This reveals that each primary blast cell executes a cell autonomous developmental program. In addition, when single cells are ablated from primary blast cell clones in the midbody segments of the O and P lineages once the primary blast cells have undergone several divisions, the remainder of the clone produces expected descendants. Also, there is no regulation to compensate for the missing structures. Therefore, even as the segment primordia gets to a stage where it contains numerous cells, development proceeds via a mosaic pattern of development (Seaver et al. 2001) (Fig. 5.8). This is in contrast to what is observed in the anterior-most segments. Although in the four rostral segments, there is a similar set of segmental pattern elements or descendant cells as is found in the more posterior segments, they arise via a distinct developmental pathway (Kuo and Shankland 2004). In these segments the O and P pattern elements arise from a single ‘op’ blast cell and furthermore, cell-cell interactions among the granddaughter cells are necessary for normal development. A detailed understanding of the generation of segmented tissues in polychaetes at the cellular level is not well characterized as it is for some clitellates. Polychaete embryos and larvae are often much smaller, and it therefore becomes more difficult to describe development at the single cell level, let alone perform experimental manipulations. Even the larger yolky polychaete embryos are substantially smaller than the leech embryos that have been used as experimental systems (leech eggs can range from 400 µm to over a millimeter whereas polychaete eggs typically range from 50 µm to 250 µm) (see Freeman and Lundelius 1992). However, several polychaetes can be successfully reared in the laboratory (Fischer and Dorresteijn 2004), and with current techniques, it is likely that there will be substantial progress in this area in the next few years. One important difference

120 Reproductive Biology and Phylogeny of Annelida between segment formation in clitellates and polychaetes is that in polychaetes there are not the large morphologically obvious teloblastic cells located at the posterior end of the larva. Several researchers have noted that the generation of mesodermal segmental tissue in polychaete larvae does not arise from asymmetric divisions from a single stem-like cell. For example, in his description of the development of Hydroides, Shearer (1911) states that the mesodermal bands “…at first consist of groups of three or four cells; they divide in all directions so that after the first division it is not possible to speak of a pole-cell, the divisions always being equal.” In an independent study of Hydroides, Wilson (1890) also states that there is an absence of teloblasts and that at the posterior end of each mesodermal band there is a group of ‘about three cells’. In a more recent study in Hydroides and Capitella, examination of cell division patterns by incorporation of the nucleotide analog BrdU reveals that during larval segment formation there is not evidence for a localized posterior growth zone (Seaver et al. 2005). Instead, there are mitotically active lateral populations of cells. It is formally possible that teloblasts exist in polychaetes but are not larger than the surrounding cells. It will be important to be able to follow some of these cells and their descendants by lineage tracing to resolve this issue and determine if this may be widespread among polychaetes. Also, it is likely to be revealing to follow the cellular homologues of the clitellate cells M and NOPQ in polychaetes to determine whether or not descendants of these cells contribute to the segmented trunk tissues in the larvae and perhaps even in post-larval segments. Like other molecular studies in annelids, the characterization of segmentation has been limited to a candidate gene approach and in this case components of the segmentation pathway as originally defined in Drosophila have been characterized. Spatial and temporal expression patterns have served as a first step indication of whether a particular gene may have a role in the segmentation process. The expression pattern of orthologues of Drosophila segmentation genes have been characterized in the leech embryo and a smaller number reported for other clitellate and polychaete representatives. To date, representatives of the gap, pair-rule and segment polarity genes have been reported. Table 5.3 shows a list of Drosophila segmentation gene orthologues whose expression has been characterized in developing annelid embryos and larvae including, in leeches, en (Wedeen and Weisblat 1991; Lans et al. 1993), hh (Kang et al. 2003), eve (Song et al. 2002), hb (Savage and Shankland 1996a), hes (Song et al. 2004), in oligochaetes, en (Bely and Wray 2001), and in polychaetes, en and wg (Seaver et al. 2001; Prud’homme et al. 2003). One ‘note of caution’ is that the expression patterns of some of these genes are not as well conserved within arthropods, and therefore, may not have the same segmentation functions in basal arthropods as they do in Drosophila. Generally, the segment polarity genes have shown a higher level of ‘intra-arthropod’ conservation, especially the transcription factor engrailed, which will be discussed separately in section 5.6.1. Of the genes examined to date in

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Table 5.3 Drosophila segmentation genes whose expression has been characterized in annelids

Gene

Homolog

Species

Ht-en

engrailed

H. triserialis

Pl-en Ch-en Pdu-en

Pdu-wnt1

Hro-hh Hro-eve Hro-hes

Lzf2

Cc-hb

T-hb

Expression pattern

Segmental expression in ectoderm and mesoderm of forming segments engrailed P. leidyi Segmentally iterated cells; CNS engrailed Chaetopterus Segmentally iterated structures; sp. mesoderm engrailed P. dumerilii Segmental stripes during regeneration; ectodermal cells during larval segmentation wingless P. dumerilii Segmental stripes during regeneration; ectodermal cells during larval segmentation hedgehog H. robusta Midgut and foregut Even H. robusta Teloblasts and primary blast cells; skipped mitotically active cells Hairy/ H. robusta Macromeres, teloblasts, primary enhancer blast cells; association with mitotic of split apparatus hunchback H. triserialis Broad expression during cleavage; nonsegmental tissue during segmentation hunchback Capitella Broad expression during cleavage; sp. I nonsegmental tissue during segmentation hunchback T. hattai Broad expression during cleavage; subset of ectodermal teloblasts

Reference Wedeen and Weisblat 1991; Lans et al. 1993 Bely and Wray 2001 Seaver et al. 2001 Prud’homme et al. 2003; Seaver et al. 2001 Prud’homme et al. 2003; Seaver et al. 2001 Kang et al. 2003 Song et al. 2002 Song et al. 2004

Savage and Shankland, 1996; Iwasa et al. 2000 Werbrock et al. 2001

Shimizu and Savage 2003

annelids, the majority of them do not show an expression pattern that would implicate them in having a direct role in the process of segment formation. In the leech embryo, many of these genes are expressed at later stages during morphogenesis in association with the development of a particular structure. This is the case for hh whose expression is correlated with development of the gut (Kang et al. 2003). Other genes, such as hb, are broadly expressed at early cleavage stages and then limited to the nonsegmental tissue during segment formation (Iwasa et al. 2000; Werbrock et al. 2001; Shimizu and Savage 2002). In the case of eve, there is expression in cells undergoing mitosis including teloblasts and primary blast cells, however, there is no sign of pair- rule or segmental patterning (Song et al. 2002). Thus, the cumulative evidence favors the idea that arthropods and annelids utilize distinct molecular pathways in forming segments. It will be interesting to determine whether annelids share any commonalities in molecular mechanism of segmentation with vertebrates. Currently, we do not have a general understanding of the molecular mechanisms that control segment formation in annelids.

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5.6 OTHER GENE EXPRESSION PATTERNS DURING MORPHOGENESIS: TWO CASE STUDIES OF THE CANDIDATE GENE APPROACH 5.6.1 Engrailed Engrailed (en) is arguably the most widely studied member of the Drosophila segmentation gene cascade. En is a member of the segment polarity class of segmentation genes and a homeodomain-containing transcription factor. The expression pattern of en is highly conserved in arthropods, making it appealing as a molecular character of segmentation for comparison with non-arthropod taxa. In arthropods, it is expressed in the ectoderm of the posterior compartment of the forming segments. In Drosophila, en has functionally been shown to be an initiator of a signaling pathway that is involved in establishing differences within the segment along the anteriorposterior axis. Drosophila mutants for the en gene show mirror duplications of elements within the segment. The main motivation for examining expression of en in annelids has been to demonstrate support or lack thereof for the homology of segmentation between annelids and arthropods. The long-standing acceptance of the Articulata clade has assumed homology of segmentation between the two groups and in fact the presence of a segmented character was heavily weighted in grouping the two together and is still supported by some, e.g. (Scholtz 2002). However, most morphological and molecular evidence has separated annelids and arthropods into two of the three great clades of bilaterians, the Lophotrochozoa and Ecdysozoa respectively. These associations raise the possibility of independent origins of segmentation between the two groups (reviewed in Seaver 2003). In annelids, the expression of en during the process of segment formation has been examined in a range of clitellate and polychaete species. En expression was examined in the embryo of Helobdella triserealis as the first segmentation gene to be characterized in annelids (Wedeen and Weisblat 1991; Lans et al. 1993). Its expression was provocative in that it appears in a transverse band in the ectoderm (derivatives of the ectoteloblasts), initially as a single cell/segment early during embryonic segment formation. En is also expressed in the mesoderm lineage, initially as 2 cells/segmental clone. In a series of experiments that examined the potential role of cellular interactions in the establishment of segment polarity in the closely related Helobdella robusta, individual cells were ablated by laser microbeam and the development of the remainder of the segment was examined. In experiments where blast cell progeny that express en or their lineal precursors were removed, the remainder of the segmental clone gave rise to its normal complement of descendants. These results demonstrate that the establishment of segment polarity in the ectoderm appears to be independent of cell interactions along the anterior-posterior axis (Seaver and Shankland 2001). In addition, ablations of en-expressing cells in the N lineage, which gives rise to a large portion of the central nervous system, do not result in

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any defects in the separation of the CNS into discrete ganglia (Shain et al. 1998). In the clitellate Pristina leidyi, a freshwater species that undergoes fission and robust anterior and posterior regeneration (but does not readily produce sexually in the lab), expression of en has been characterized and is the only non-leech clitellate studied to date. In Pristina, the expression of en is similar during both fission and regeneration, and is present in a small number of cells that are located in the position of the CNS as well as additional segmentally iterated cells that do not correspond with previously identified structures (Bely and Wray 2001). The expression pattern of en has been examined in several polychaete species during formation of larval segments. These data represents a critical part of the story since it is hard to know if en expression patterns in clitellates are representative for annelids in general. At the cellular level, there are clearly distinctions in the way that the two groups generate segments. In Chaetopterus, Ch-en is expressed in a dynamic and complex pattern during all larval stages. Ch-en is present in lateral portions of the CNS, in some mesoderm, and in association with differentiation of particular structures such as the aliform notopodia in the B region of the body (Seaver et al. 2001). In Capitella, en is also expressed in segmentally iterated structures such as in a subset of neurons of the CNS and also in regions where cells are delaminating from the ectoderm epithelia (Seaver 2003). In both these species, expression does not appear as segmental stripes in the ectoderm prior to morphological segmentation, but instead is present in segmentally iterated structures. In contrast, in Platynereis, Pdu-en and Pdu-wg expression has reported expression in a stripe pattern reminiscent of the arthropod pattern in the regenerating adult and larvae (Prud’homme et al. 2003). Although it is reasonable to propose that the mechanisms by which segments are generated during regeneration may be the same as those that operate during development, this has not be rigorously established. The differences in expression of en among polychaetes may reflect the complexity of this grade of animals and future studies will be necessary to try to determine if there are in fact distinct mechanisms of segment formation even within the annelids.

5.6.2 Hox Genes The Hox genes, a subset of the homeobox transcription factors, have been a major focus of developmental biology because of their role in homeotic mutations in flies. Several aspects of the Hox genes make them especially intriguing. First, they are responsible for homeotic transformations of patterned structures in both flies and vertebrates. In these homeotic transformations, a defect in protein structure or regulation of a single gene acts like a “master switch” to turn, for example, a fly antenna into a leg. Second, all Hox genes contain a highly conserved DNA-binding motif, termed the homeodomain (or homeobox at the nucleic acid level). Homeobox genes have been found in all metazoan groups. Third, the Hox

124 Reproductive Biology and Phylogeny of Annelida genes have a clustered genomic organization, which is conserved in all groups. Finally, the spatial order of genes on the chromosome mirrors, generally, the spatial order of gene expression along the anteroposterior body axis, a principle called ‘colinearity’ (Manak and Scott 1994). The high level of sequence conservation within the homeobox makes it relatively easy to use molecular methods to clone Hox genes in animals, like annelids, where genetic methods are not possible. Therefore, with the interesting characteristics of the Hox genes touched on above, and the ready technology to clone them without the necessity of genetics, this gene family has attracted a great deal of attention among annelid developmental biologists. Hox gene number in annelids: Much has been made of the evolution of the Hox cluster itself in different taxa, and its effect on the evolution of developmental patterns. The notion is that the Hox cluster was built up over time by gene duplications in ancestral lineages, and that developmental patterning in various groups is affected by the number of Hox genes of each particular type present. One line of research has been to clone as many Hox gene fragments as possible from a species and to infer the probable state of the Hox cluster from these individual fragments using phylogenetic sequence analysis (Dick and Buss 1994; Snow and Buss 1994; Irvine et al. 1997; Rosa et al. 1999; Andreeva et al. 2001, and refs. in Table 5.4; Cho et al. 2003). As of this writing a Hox cluster has not been mapped in annelids, but Andreeva et al. (2001) showed linkage of Nereis Hox genes on a 2.5 megabase genomic fragment, consistent with a clustered organization. The first major finding from this work has been that annelids likely have the complete complement of Hox gene types (termed trans-paralogs, or orthologs) present in more “complex” animals like flies and vertebrates. A caveat to this view is that certain of the annelid genes may be the result of independent duplication within the annelid lineage of ancestral genes, making these Hox genes not strictly orthologous to those of flies and vertebrates (Balavoine et al. 2002). In polychaetes a single Hox gene has been found corresponding to each of the Hox genes in Drosophila, except that in Nereis there are two posterior genes, corresponding to the Drosophila Abd-B gene, an apparently ancestral trait of lophotrochozoans. Clitellates appear to have Hox gene complements somewhat more altered with respect to the fly cluster. In the leech, Hox2, Hox3, and posterior group orthologs have never been found, indicating possible gene loss. On the other hand, there are two genes each for the Hox4 and Hox5 groups, indicating gene duplication events in the leech lineage after divergence from polychaetes (Table 5.4). In Stylaria there is data indicating that there are two or more duplicates of the Hox1 gene (Snow and Buss 1994; Cho et al. 2003). Hox expression in polychaetes. Expression patterns have been examined in the polychaete Chaetopterus variopedatus for five Hox genes (Irvine and Martindale 2000) and for one gene in Nereis virens (Kulakova et al. 2002). Their names reflect their orthology assignments: CHv- Hox1, 2, 3,

Early Annelid Development, A Molecular Perspective Table 5.4 Annelid Hox genes Orthology Probable annelid orthologs group Polychaetes Leeches Gene Hox1 Nvi-lab CHv-Hox1 Htr-Lox7 Hox2 Nvi-pb CHv-Hox2 none found Hox3 Nvi-Hox3 CHv-Hox3 none found Hox4 Nvi-Dfd Htr-Lox6 CHv-Hox4 Htr-Lox18 Hox5 Nvi-Scr Htr-Lox20 CHv-Hox5 Hm-Lox1 Hox6 Nvi-Lox5 Htr-Lox5 Hox7 Nvi-Lox2 Htr-Lox2 Hox8 Nvi-Lox4 Hm-Lox4 Hox9-14 Nvi-Post1 Nvi-Post2 none found

125

References

Kourakis et al. 1997

Kourakis et al. 1997 Kourakis and Martindale 2001 Kourakis et al. 1997 Aisemberg and Macagno 1994 Kourakis et al. 1997 Nardelli-Haefliger and Shankland 1992 Wong et al. 1995

Species designations: same as Table 5.1 plus, Nvi, Nereis virens; Hm, Hirudo medicinalis. Polychaete References: Nvi (de Rosa et al. 1999); CHv (Irvine and Martindale 2000). Notes: i) The leech gene pairs Lox6/Lox18 and Lox20/Lox1 are thought to be paralogs resulting from independent duplication events within leeches. ii) Only Hox genes for which the entire homeobox has been sequenced are included. For other likely Hox genes from polychaetes and oligochaetes based on partial homeobox sequences refer to Dick and Buss 1994; Snow and Buss 1994; Irvine et al. 1997; Cho et al. 2003.

4, 5, being putative orthologs of orthology groups 1-5 in arthropods and vertebrates, while Nvi-Post1 represents an ortholog of the posterior group. mRNA expression of each of the Chaetopterus genes begins early in larval development — in fact, CHv-Hox2 is present as a maternal transcript (Peterson et al. 2000). In-situ hybridization (ISH) and probe excess titration shows abundant transcripts present for CHv-Hox1 and CHv-Hox2 in the early trochophore, with detectable expression of the other three genes slightly later. In each case, transcripts are present in bilateral cell populations at the posterior pole of the larva in the region coinciding with the “growth zone”, or teloblast location (Fig. 5.9A). The present data does not confirm that the expressing cells are indeed teloblasts. It should be noted that this early expression occurs long before morphological segmentation is evident. The ISH staining pattern in bilateral posterior cell populations persists in all larval stages through metamorphosis, even when discontinuous with staining in other regions of the larva. As larval development proceeds, transcripts of each of the five genes are detected extending from these posterior regions anteriorly in bilateral strips of ectoderm straddling the ventral midline, probably destined for the ventral nerve cord. By late larval stages it becomes apparent that the anterior boundaries of expression, with the exception of CHv-Hox2, follow the general rule of “colinearity”; the idea that anterior expression boundaries are the

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Colour Figure

Fig. 5.9 Photographs of Hox gene mRNA expression patterns as detected by whole mount in-situ hybridization. A. Ventral view of a stage L2 trochophore larva of the polychaete Chaetopterus variopedatus showing hybridization to a CHv-Hox3 riboprobe. Transcripts detected in bilateral posterior cell populations are typical of the pattern seen for CHv-Hox1-Hox5. B. Staining pattern of a Nvi-Post1 (Hox posterior group homolog) riboprobe in 86h. larva of the polychaete Nereis virens in ventrolateral view. Hybridization is apparent in 3 bilaterally paired groups of cells probably corresponding to mesodermal components of the parapodial rudiments, and precedes apparent morphological segmentation in the larva. The asterisk marks the mouth, and arrow denotes the prototroch. C, D. Expression of Hox genes in stage L5 larvae of C. variopedatus. CHv-Hox3 (C) and CHv-Hox5 (D) staining in the bilateral nerve cords and parapodia of the anterior tagma (bracket), the aliform notopodial rudiment (single arrowhead), and posterior ganglia (arrow). For CHv-Hox5, staining is restricted to the anterior of the B1 ganglion (double arrowhead), and single bilaterally located neurons in the B3 and B4 ganglia (double-headed arrows). E. Lateral view of a stage 10 embryo of the leech Helobdella robusta showing staining for a Lox6 (Hox4) riboprobe. Hollow arrowhead points to the R3 neuromere and solid arrowhead to epidermal structures in the same segment. F. Ventral view of the anterior portion of an H. triserialis embryo showing staining for a Lox20 (Hox5) probe. Transcripts are detected in segmentally iterated mesodermal septa (arrows). A, C, D, from Irvine, S. Q. and Martindale, M. Q. 2000. Developmental Biology 217: 333-351, Fig. 5e, 7c, 7e.; B, from Kulakova, M. A., Kostyuchenko, R. P., Andreeva, T. F. and Dondua, A. K. 2002. Mechanisms of Development 115: 177-179, Fig. 4.; and E, F, from Kourakis, M. J., Master, V. A., Lokhorst, D. K., Nardelli-Haefliger, D., Wedeen, C. J., Martindale, M. Q. and Shankland, M. 1997. Developmental Biology 190: 284-300, Figs. 6a, 7a.

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most important to regional or segmental specification, and that these boundaries are staggered along the anterior-posterior axis in rank order of the gene’s location from 3’ to 5’ on the chromosome. CHv-Hox2 has an anterior boundary at the anterior edge of the first segment, the CHv-Hox1 boundary is at the second segment, CHv-Hox3 at the third, CHv-Hox4 at the fourth, and CHv-Hox5 at the fifth. In these later stages, approaching metamorphosis, the strongest expression is in the nerve cord, with transcripts also found in various combinations for the different genes of the anterior parapodia, mid-body ciliary bands, and specialized parapodial structures (see Fig. 5.9C & D for examples). Apart from the staggered anterior boundaries of expression, several of the genes examined have defined posterior boundaries of expression. These posterior boundaries were found to coincide with major transitions in segmental morphology, and have been argued to have a possible role in tagmosis, or evolutionary changes in the functional regionalization of the body plan (Fig. 5.10B) (Irvine and Martindale 2001). Similar patterns have been observed in arthropods (Akam 2000; Hughes and Kaufman 2002). The only other published expression data for Hox genes in polychaetes as of this writing is ISH data for the Post1 gene in Nereis virens, a worm without the pronounced heteronomy in segmental morphology along the body axis seen in Chaetopterus (Kulakova et al. 2002). As mentioned above, two genes of the posterior class have been found in N. virens. Expression of Nvi-Post1 is detected in the earliest larval stages in large bilaterally disposed groups of cells posterior to the prototroch. Soon these cell populations resolve into three bilateral pairs of dorso-ventral bands of expression extending from the prototroch to the posterior (Fig. 5.9B). By the time morphological segmentation is visible in the metatrochophore, expression has become restricted to posterior spots similar to that subset of the later Chaetopterus pattern. The early stage expression in three bilateral anteroposterior groupings recalls the idea that polychaete larvae form three initial “larval” segments by subdivision of larval mesoderm, with subsequent segments being derived sequentially from the teloblastic growth zone (heteronomicity) (Iwanoff 1928; Irvine et al. 1999). Curiously, in other wellstudied taxa, such as vertebrates and flies, Hox genes of the posterior group are expressed only in posterior segments, rather than the expression in a broad range of the anterior-posterior axis seen in N. virens. This aspect of expression suggests that Nvi-Post1 has a function unconnected with the commonly invoked role in regional specification, but rather some more general role in larval segment formation. This possible general role may relate to the strong early expression seen for anterior Hox genes in Chaetopterus, since this also appears long before it can be spatially related to a region along the body axis. Another aspect of the Chaetopterus expression patterns worth noting is that CHv-Hox1 transcripts are robustly expressed at the foregut midgut boundary from the earliest larval stages. The expression in this part of the gut persists through metamorphosis. A transitory phase of expression in the

128 Reproductive Biology and Phylogeny of Annelida foregut is also seen for CHv-Hox2 in middle larval stages. Interestingly, the Drosophila homolog of CHv-Hox1, labial, is expressed in the gut in Drosophila, appearing in the anterior and posterior parts of the midgut rudiment at germband extension (Diederich et al. 1989). Proboscipedia, the Drosophila homolog of CHv-Hox2, is also expressed briefly in visceral mesoderm in the posterior foregut (Pultz et al. 1988). Thus, Chaetopterus may share with Drosophila a role for Hox1 and Hox2 genes in early development of the posterior foregut. Hox gene expression in glossiphoniid leeches. Hox gene expression has been examined in some detail in the leeches Helobdella and Hirudo, using mostly in-situ hybridization to mRNA, and in some cases immunohistochemistry (refer to Table 5.4 for references). Transcripts or protein are found predominantly in the ventral nerve cord (VNC), much as seen in the polychaete Chaetopterus (e.g. Fig. 5.9D). Unlike Chaetopterus, however, onset of expression is not reported until morphological segmentation is well underway (stage 8 in Helobdella). As seen in other taxa, anterior boundaries of expression in the VNC obey the “rule” of colinearity, with Lox7(Hox1) expressed in the first rostral ganglion (R1) and all along the body axis posteriorly, Lox6(Hox4) transcripts seen in posterior R2 and R3 to the caudal terminus, Lox20(Hox5) in posterior R3 and anterior R4, etc. (refer to Fig. 5.10A). Note that Hox2 and Hox3 orthologs have never been found in leeches (even though readily found in other annelids), and curiously, the anterior boundary for Lox6(Hox4) is located just one segment behind that for the Hox1 ortholog Lox7. Thus it appears that leeches may have lost their Hox2 and Hox3 genes, and furthermore that the anterior boundaries of expression of the other Hox genes seem to have shifted anteriorly as if to compensate for the loss. Several of the Hox genes examined in leeches have limited expression domains outside the VNC as well. Notably, Lox6(Hox4) is expressed in various peripheral sensory structures (Wong and Macagno 1998). Lox20(Hox5) is expressed only in parts of two rostral ganglia, but is expressed widely in mesoderm of the anterior third of the trunk after assembly of the germinal plate, especially in the intersegmental septa. Lox1, a possible Hox5 duplicate of Lox20, is expressed in nephridia and segmentally iterated cells of the body wall, while Lox5(Hox6) transcripts are seen transiently in segmentally iterated mesodermal cells. The Hox7/8 ortholog Lox2 is expressed in the genital primordia and dorso-ventral muscles, and another Hox7/8 gene, Lox4, shows transcripts in peripheral tissues in the mid-posterior trunk (Fig. 5.10A). While several of the genes are expressed from the anterior boundary caudally along the entire VNC, some genes have defined posterior boundaries of expression. In the cases of Lox2 and Lox4, the Hox7/8 orthologs, the posterior boundary is at the juncture between the midbody ganglia and fused caudal ganglion, and could be interpreted as defining the juncture between body tagma as is seen in Chaetopterus (Irvine and Martindale 2001).

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A

B

Fig. 5.10 Diagrams of Hox gene expression patterns in A. leeches, and B. the polychaete Chaetopterus. A. Diagrammatic ventral view of left half of a stage 10 Helobdella triserialis embryo with domains of Hox gene expression at mid-late stages of embryogenesis depicted in bars below for the genes listed at the left. Segments are numbered within the rostral (R), midbody (M) and caudal (C) domains. Expression in the central nervous system is shown by the dark bars, with expression in other organs shown below in lighter bars (refer also to notes below). He prefix to gene name refers to H. triserialis or H. robusta (Kourakis et al. 1997; Nardelli-Haefliger and Shankland 1992), while Hi refers to data from Hirudo medicinalis (Wysocka-Diller et al. 1989; Wong et al. 1995; Wong and Macagno 1998). B. Diagrammatic ventral view of left half of a stage L5 Chaetopterus variopedatus larva with domains of Hox gene expression depicted as in A (Irvine and Martidale 2000). Segments are numbered within the A, B, and C body tagma. Abbreviations: pr/pe, prostomium/peristomium; pyg, pygidium. Notes: 1, expression in eye primordia and 2, expression in peripheral nervous system in Hirudo. 3, expression in mesodermal septae. 4, segmentally iterated mesodermal expression. 5, expression in genital primodia and dorsoventral muscles 6, peripheral expression. 7, parapodial expression. 8, expression in parapodia and ventral plastron. 9, anterior parapodial and aliform notopodial rudiment expression. 10, expression as in 9 plus in ectoderm of setigers B2-C1.

130 Reproductive Biology and Phylogeny of Annelida Within the CNS, expression typically resolves to a subset of segmentally iterated neurons within each ganglion. This pattern has been documented in detail for Lox6 (Wong and Macagno 1998), Lox2 (Berezovskii and Shankland 1996), and Lox4 (Wong et al. 1995). The Lox2 and Lox4 studies showed that segment specific differences in particular leech neurons correlate in a consistent way with the expression of these gene products. Thus at the level of neuronal cell type, the Hox genes may regulate regional differences much as they have been implicated, as mentioned above, in broader regionalization of the body plan. For two of the orthology groups, Hox4 with Lox6 and Lox18, and Hox5 with Lox20 and Lox1, the leech genes are thought to be paralogs resulting from independent duplication within the leech lineage. In both cases, one of the genes of the pair obeys the colinearity principle in its expression pattern, while the other, Lox18 (Kourakis and Martindale 2001) and Lox1 (Aisemberg and Macagno 1994) respectively, are expressed all along the segmented body axis. This situation suggests that after duplication one of the resulting genes was released from the constraint of colinearity and evolved a divergent spatial expression pattern. An interesting analysis of Hox gene expression in the leech takes advantage of the phenomenon of bandlet slippage. In this type of experiment, laser ablation is used to kill a blast cell (segmental founder cell) early in development of the germinal bands, causing the segmental anlage derived from one teloblast to develop out of register with adjacent teloblast progeny. Thus, segmental founder cell progeny develop in a foreign segmental environment. Nardelli-Haefliger et al. (1994) looked at the effect of slippage on the expression of Lox2(Hox7/8) and found that the displacement of blast cell clones brought about a corresponding displacement of the Lox2 protein expression domain. Thus, Hox gene expression is dependent on the birth order of the parent blast cells, rather than position of cells, with respect to adjacent tissues within the segmental anlage. This finding is consistent with previous work that showed that the identity of segment-specific neuronal subtypes is determined at or near the time of birth of blast cells from the parent teloblast, and not altered if the cellular environment of the blast cell progeny is changed (Martindale and Shankland 1990). Hox gene expression in annelids — functional implications. From the summaries above, some commonalities are apparent between the Hox gene expression patterns seen in polychaetes and leeches (Fig. 5.10). The most immediately apparent characteristic is that in both groups the predominant expression is in the VNC — in some cases transcripts seen widely distributed within ganglia, and in others confined to a subset of segmentally reiterated neurons within ganglia. With certain exceptions, colinearity holds, with anterior boundaries of expression forming a nested series from anteriorposterior in the same rank order as the gene’s orthology group numbering. These nested expression domains are largely overlapping along the anterior posterior axis. In addition to the VNC expression, in both polychaetes and

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leeches various Hox genes are expressed in particular mesodermal and ectodermal structures of the trunk. Apart from these commonalities in annelid expression patterns, several differences between polychaete and leech expression patterns are also apparent. Most notably, the polychaete Hox genes are expressed much earlier with respect to the onset of morphologically apparent segmentation than are the leech genes, and may even be expressed in teloblasts. No expression in the teloblasts of leeches has been published to date, and the earliest expression is generally reported as occurring at late stage 8, when the germinal plate is already largely assembled. Another difference is that polychaetes have Hox2 and Hox3 orthologs expressed in colinear fashion, while these genes have not been found in leeches. As mentioned above, expression domains may have shifted anteriorly along with the apparent loss of these genes (Fig. 5.10A). Finally, the polychaete Chaetopterus has a robust expression of its Hox1 ortholog, and transient expression of Hox2, at the foregut-midgut boundary, a characteristic reminiscent of the pattern in flies, but not seen in leeches (or interestingly, in the published data from other arthropods (Irvine and Martindale 2001)). The colinear overlapping expression domains seen in annelids are much like those seen in the other segmented taxa — arthropods and vertebrates. However, there are significant differences in the extent of expression. In arthropods, particularly Drosophila, the Hox genes are considerably more broadly expressed in ectodermal and mesodermal segmental primordia (e.g. (Carroll et al. 1988; Jack et al. 1988; Pultz et al. 1988; Diederich et al. 1989). While expressed in arthropod neurogenic tissues, the predominance of expression in the VNC seen in annelids is not apparent in chelicerates, crustaceans, or primitive insects (e.g. Tear et al. 1990; Averof and Akam 1995; Damen et al. 1998; Telford and Thomas 1998; Abzhanov et al. 1999; Peterson et al. 1999). Rather, in these arthropods, expression is seen in ectodermal and mesodermal tissues contributing to body wall structures as much, or more prominently, than is the expression in the VNC. In vertebrates, on the other hand, the prominent sites of Hox gene expression are the nerve cord and the paraxial mesoderm. While the strong nerve cord expression is similar to that seen in annelids, the expression in annelids outside the VNC does not look like the mesodermal expression in vertebrates (eg. Burke et al. 1995; Prince et al. 1998). The vertebrate expression, like that in arthropods, tends to be more generalized within the somitic primordia, whereas in annelids the expression of a particular Hox gene, outside the VNC, tends to be associated with a particular structure. Examples are the presence of transcripts of the Hox7/8 ortholog Lox2 in genital primordia and dorso-ventral muscles of the leech, or CHv-Hox4 in peripheral tissues of the highly modified mid-body parapodia of Chaetopterus. Other annelid Hox genes show little or no expression outside the VNC. Generally speaking, initial expression domains in all three phyla are broader and as development proceeds become more restricted. In the annelids, however, even initial domains are limited as compared with

132 Reproductive Biology and Phylogeny of Annelida arthropods and vertebrates (leaving aside the early “growth zone” expression in polychaetes). Typically, early expression might be broadly distributed in a number of ganglia and become restricted later to subsets of segmentally reiterated neurons within ganglia, e.g. CHv-Hox5 (Irvine and Martindale 2000), or Lox6(Hox4) (Wong and Macagno 1998). From the studies on insects and vertebrates, the paradigm of Hox gene function has been characterized as a “combinatorial code”, with the particular combination of Hox genes present in a segment, on one level, specifying segmental identity, and in a particular cell, on another level, specifying cell type (at least in part see e.g. Akam 1998; Lohmann and McGinnis 2002). The data is consistent with a similar kind of Hox gene function in annelids in the VNC. Outside the VNC function is more difficult to infer. The general pattern suggests that Hox proteins are deployed in various structures along the body axis, but not in the generalized segmental pattern seen in arthropods and vertebrates.

5.7 SUMMARY AND PROSPECTS FOR FUTURE RESEARCH Taken together, the molecular developmental patterns seen in annelids differ significantly from those in vertebrates and Drosophila. It is worth reiterating a few major examples. Nanos is a primary determinant of posterior fates in the early Drosophila embryo. Its homolog Htr-nos in the leech is localized to the ectodermal teloblast precursor, and rather than establishing anteriorposterior polarity, it is associated with germinal band cell fate. Hunchback is also a key element of anterior-posterior axis specification in flies, but like Htr-nos, its annelid homologs do not show any relationship to axial patterning. Hedgehog is a segment polarity gene in flies, required for the proper patterning of the earliest segmental anlage. In vertebrates, hedgehog homologs act as morphogens patterning the limbs, somites, and other structures. Leech hedgehog is neither segmentally expressed, or a clear morphogen, but is involved in specifying subsets of mesoderm associated with the gut. Finally, the Hox genes in annelids share some basic characteristics with the expression patterns in flies and vertebrates, but differ in that their expression is largely limited to the central nervous system. These differences in molecular patterns suggest that fundamental developmental mechanisms differ in annelids, and possibly lophotrochozoans in general, as compared with vertebrates and insects. This situation provides an impetus to further work, since if lophotrochozoan molecular developmental patterns differ from the rest of the animal kingdom, understanding of the totality of animal developmental biology will depend on exploring developmental mechanisms in annelids — the major segmented lophotrochozoan phylum. Further understanding of annelid development on a molecular level also presents a considerable challenge, since genetic tools are not available for functional study. Fortunately, new technologies, such as gene knockdown using morpholinos,

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hold promise for exploring the function of genes discovered in other organisms. Beyond this, functional genomics, such as microarray techniques, offer an alternative to classical genetics for the discovery of molecules important to the particular developmental patterns in annelids.

5.8

ACKNOWLEDGEMENTS

We thank Dave Lambert for the picture of MAP Kinase staining in H. elegans. S. I. acknowledges the support of NIH Grant Number P20 RR016457 from the BRIN Program of the National Center for Research Resources.

5.9 LITERATURE CITED Abzhanov, A., Popadic, A. and Kaufman, T. C. 1999. Chelicerate Hox genes and the homology of arthropod segments. Evolution and Development 1: 77-89. Aisemberg, G. O. and Macagno, E. R. 1994. Lox1, an Antennapedia-class homeobox gene, is expressed during leech gangliogenesis in both transient and stable central neurons. Developmental Biology 161: 455-465. Akam, M. 1998. Hox genes: From master genes to micromanagers. Current Biology 8: R676-R678. Akam, M. 2000. Arthropods: Developmental diversity within a (super)phylum. Proceedings of the National Academy of Sciences USA 97: 4438-4441. Anderson, D. T. 1966a. The comparative early embryology of the Oligochaeta, Hirudinea and Onychophora. Proceedings of the Linnean Society of New South Wales 91: 10-43. Anderson, D. T. 1966b. The comparative embryology of the Polychaeta. Acta Zoologica 47: 1-42. Anderson, D. T. 1973. Embryology and Phylogeny in Annelids and Arthropods. Pergamon, Oxford. 495 pp. Andreeva, T. F., Kuk, C., Korchagina, N. M., Eikem, M. and Dondua, A. K. 2001. Cloning and analysis of structural organization of Hox genes in the polychaete Nereis virens. Russian Journal of Developmental Biology 32: 183-191. Arai, A. N. A. and Shimizu, T. 2001. Specification of ectodermal teloblast lineages in embryos of the oligochaete annelid Tubifex: involvement of novel cell-cell interactions. Development 128: 1211-1219. Arendt, D. and Nübler-Jung, K. 1997. Dorsal or ventral: similarities in fate maps and gastrulation patterns in annelids, arthropods and chordates. Mechanisms of Development 61: 7-21. Arendt, D., Technau, U. and Wittbrodt, J. 2001. Evolution of the larval foregut. Nature 409: 81-85. Astrow, S., Holton, B. and Weisblat, D. 1987. Centrifugation redistributes factors determining cleavage patterns in leech embryos. Developmental Biology 120: 270-283. Astrow, S. H., Holton, B. and Weisblat, D. A. 1989. Teloplasm formation in a leech, Helobdella triserialis, is a microtubule-dependent process. Developmental Biology 135: 306-319. Averof, M. and Akam, M. 1995. Hox genes and the diversification of insect and crustacean body plans. Nature 376: 420-423. Balavoine, G., de Rosa R. and Adoutte, A. 2002. Hox clusters and bilaterian phylogeny. Molecular phylogenetics and Evolution 24: 366-373.

134 Reproductive Biology and Phylogeny of Annelida Bely, A. E. and Wray, G. A. 2001. Evolution of regeneration and fission in annelids: insights from engrailed- and orthodenticle-class gene expression. Development 128: 2781-91. Berezovskii, V. and Shankland, M. 1996. Segmental diversification of an identified leech neuron correlates with the segmental domain in which it expresses Lox2, a member of the Hox gene family. Journal of Neurobiology 29: 319-329. Bruce, A. E. E. and Shankland, M. 1998. Expression of the head gene Lox22-Otx in the leech Helobdella and the origin of the bilaterian body plan. Developmental Biology 201: 101-112. Brusca, R. C. and Brusca, G. J. 1990. Invertebrates. Sinauer, Sunderland, MA. Burke, A. C., Nelson, C. E., Morgan, B. A. and Tabin, C. 1995. Hox genes and the evolution of vertebrate axial morphology. Development 121: 333-346. Cantillana, V., Urrutia, M., Ubilla, A. and Fernandez, J. 2000. The complex dynamic network of microtubule and microfilament cytasters of the leech zygote. Developmental Biology 228: 136-149. Carroll, S. B., DiNardo, S., O’Farrell, P. H., White, R. A. H. and Scott, M. P. 1988. Temporal and spatial relationships between segmentation and homeotic gene expression in Drosophila embryos: distributions of the fushi tarazu, engrailed, Sex combs reduced, Antennapedia, and Ultrabithorax proteins. Genes and Development 2: 350-360. Child, C. M. 1900. The early development of Arenicola and Sternaspis. Wilhelm Roux’s Archiv für Entwicklungsmechanik der Organismen 9: 587-722. Cho, S. J., Cho, P. Y., Lee, M. S., Hur, S. Y., Lee, J. A., Kim, S. K., Koh, K. S., Na, Y. E., Choo, J. K., Kim, C.-B. and Park, S. C. 2003. Hox genes from the earthworm Perionyx excavatus. Development Genes and Evolution 213: 207-210. Clement, A. C. 1952. Experimental studies on germinal localization in Ilyanassa. I. The role of the polar lobe in determination of the cleavage pattern and its influence on later development. Journal of Experimental Zoology 121: 563-626. Clement, A. C. 1962. Development of Ilyanassa following removal of the D macromere at successive cleavage stages. Journal of Experimental Zoology 149: 193-216. Conlon, F. L. and Smith, J. C. 1999. Interference with brachyury function inhibits convergent extension, causes apoptosis, and reveals separate requirements in the FGF and activin signalling pathways. Developmental Biology 213: 85-100. Costello, D. P. 1945. Experimental studies of geminal localization in Nereis. I. The development of isolated blastomeres. Journal of Experimental Zoology 100: 1960. Damen, W. G. M., Hausdorf, M., Seyfarth, E.-A. and Tautz, D. 1998. A conserved mode of head segmentation in arthropods revealed by the expression pattern of hox genes in a spider. Proceedings of the National Academy of Sciences USA 95: 10665-10670. Delsman, H. C. 1916. Eifurchung und Keimblattbildung bei Scoloplos armiger. Tijdschr. ned. dierk. Vereen. 14: 383-498. Dick, M. H. and Buss, L. W. 1994. A PCR-based survey of homeobox genes in Ctenodrilus serratus (Annelida: Polychaeta). Molecular Phylogenetics and Evolution 3: 146-158. Diederich, R. J., Merrill, V. K. L., Pultz, M. A. and Kaufman, T. C. 1989. Isolation, structure, and expression of labial, a homeotic gene of the Antennapedia Complex involved in Drosophila head development. Genes and Development 3: 399-414. Dohle, W. 1999. The ancestral cleavage pattern of the clitellates and its phylogenetic deviations. Hydrobiologia 402: 267-283.

Early Annelid Development, A Molecular Perspective

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Dorresteijn, A. 1987. A correlative study of experimentally changed first cleavage and Janus development in the trunk of Platynereis dumerilii (Annelida, Polychaeta). Roux’s Arch Developmental Biology 196: 51-58. Dorresteijn, A. W. C. and Eich, P. 1991. Experimental change of cytoplasmic composition can convert determination of blastomeres in Platyneries dumerilii (Annelida, Polychaeta). Roux’s Arch Developmental Biology 200: 342-351. Eckberg, W. R. and Anderson, W. A. 1995. Cytoskeleton, cellular signals, and cytoplasmic localization in Chaetopterus embryos. Current Topics in Developmental Biology 31: 5-39. Eisig, H. 1899. Zur Entwicklungsgeschichte der Capitelliden. Mittheilungen Aus der Zoologischen Station Zu Neapel 13: 1-292. Fernandez, J., Roegiers, F., Cantillana, V. and Sardet, C. 1998. Formation and localization of cytoplasmic domains in leech and ascidian zygotes. International Journal of Developmental Biology 42: 1075-1084. Ferrell, J. E. 1996. MAP kinases in mitogenesis and development. Curr Top. Dev. Biol 33: 1-60. Fietz, M. J., Concordet, J.-P., Barbosa, R., Johnson, R., Krauss, S., McMahon, A. P., Tabin, C. and Ingham, P. W. 1994. The hedgehog gene family in Drosophila and vertebrate development. Development 1994 Suppl. Fischer, A. and Dorresteijn, A. 2004. The polychaete Platynereis dumerilii (Annelida): a laboratory animal with spiralian cleavage, lifelong segment proliferation and mixed benthic/pelagic life cycle. Bioessays 26: 314-325. Freeman, G. and Lundelius, J. W. 1992. Evolutionary implications of the mode of D quadrant specification in coelomates with spiral cleavage. Journal of Evolutionary Biology 5: 205-247. Goldstein, B., Leviten, M. W. and Weisblat, D. A. 2001. Dorsal and Snail homologs in leech development. Development Genes and Evolution 211: 329-337. Goto, A. K. K. and Shimizu, T. 1999. Cell lineage analysis of pattern formation in the Tubifex embryo. International Journal of Developmental Biology 43: 317-327. Guralnick, R. P. and Linberg, D. R. 2001. Reconnecting cell and animal lineages: What do cell lineages tell us about the evolution and development of Spiralia. Evolution 55: 1501-1519. Halanych, K. M., Bacheller J. D., Aguinaldo A. M., Liva S. M., Hillis D. M. and Lake, J. A. 1995. Evidence from 18S ribosomal DNA that the lophophorates are protostome animals. Science 267: 1641-3. Haszprunar, G., Salvini-Plawen, L. and Reiger, R. M. 1995. Larval planktotrophy — a primitive trait in the Bilateria? Acta Zoologica 76: 141-154. Hatt, P. 1932. Essais experimentaux sur le localisations germinales dans l’oeuf d’une annelide (Sabellaria alveolata L.). Archives d’Anatomie Microscopique et de Morphologie Experimentale 28: 81-98. Hemavathy, K., Ashraf, S. I. and Ip, Y. T. 2000. Snail/Slug family of repressors: slowly going into the fast lane of development and cancer. Gene 257: 1-12. Henry, J. J. 1986. The role of unequal cleavage and the polar lobe in the segregation of developmental potential during first cleavage in the embryo of Chaetopterus variopedatus. Roux’s Archives of Developmental Biology 195: 103-116. Henry, J. J. and Martindale, M. Q. 1987. The organizing role of the D quadrant as revealed through the phenomenon of twinning in the polychaete Chaetopterus variopedatus. Roux’s Archives of Developmental Biology 196: 499-510. Huang, F. Z., Bely, A. E. and Weisblat, D. A. 2001. Stochastic WNT signalling between nonequivalent cells regulates adhesion but not fate in the two-cell leech embryo. Current Biology 11: 1-7.

136 Reproductive Biology and Phylogeny of Annelida Huang, F. Z., Kang, D., Ramirez-Weber, F. A., Bissen, S. T. and Weisblat, D. A. 2002. Micromere lineages in the glossiphoniid leech Helobdella. Development 129: 71932. Huang, F. Z. and Weisblat, D. A. 1996. Cell fate determination in an annelid equivalence group. Development 122: 1839-47. Hughes, C. L. and Kaufman, T. C. 2002. Hox genes and the evolution of the arthropod body plan. Evolution and Development 4: 459-499. Irvine, S. Q., Chaga, O. and Martindale, M. Q. 1999. Larval ontogenetic stages of Chaetopterus: Developmental heterochrony in the evolution of chaetopterid polychaetes. Biological Bulletin 197: 319-331. Irvine, S. Q. and Martindale, M. Q. 2000. Expression patterns of anterior Hox genes in the polychaete Chaetopterus: Correlation with morphological boundaries. Developmental Biology 217: 333-351. Irvine, S. Q. and Martindale, M. Q. 2001. Comparative analysis of Hox gene expression in the polychaete Chaetopterus: Implications for the evolution of body plan regionalization. American Zoologist 41: 640-651. Irvine, S. Q., Warinner, S. A., Hunter, J. D. and Martindale, M. Q. 1997. A survey of homeobox genes in Chaetopterus variopedatus and analysis of polychaete homeodomains. Molecular Phylogenetics and Evolution 7: 331-345. Iwanoff, P. P. 1928. Die Entwicklung der Larvalsegmente bei den Annelides. Zeitschrift für Morphologie und Ökologie der Tiere 10: 62-161. Iwasa, J. H., Suver, D. W. and Savage, R. M. 2000. The leech hunchback protein is expressed in the epithelium and CNS but not in the segmental precursor lineages. Development Genes and Evolution 210: 277-288. Jack, T., Regulski, M. and McGinnis, W. 1988. Pair-rule segmentation genes regulate the expression of the homeotic selector gene, Deformed. Genes and Development 2: 635-651. Jeffery, W. R. and Wilson, L. J. 1983. Localization of messenger mRNA in the cortex of Chaetopterus eggs and early embryos. Journal of Embryology and Experimental Morphology 75: 225-239. Kang, D., Huang, F., Li, D., Shankland, M., Gaffield, W. and Weisblat, D. A. 2003. A hedgehog homolog regulates gut formation in leech (Helobdella). Development 130: 1645-1657. Kang, D., Pilon, M. and Weisblat, D. A. 2002. Maternal and zygotic expression of a nanos-class gene in the leech Helobdella robusta: primordial germ cells arise from segmental mesoderm. Developmental Biology 245: 28-41. Kourakis, M. J. and Martindale, M. Q. 2001. Hox gene duplication and deployment in the leech Helobdella. Evolution and Development 3: 145-153. Kourakis, M. J., Master, V. A., Lokhorst, D. K., Nardelli-Haefliger, D., Wedeen, C. J., Martindale, M. Q. and Shankland, M. 1997. Conserved anterior boundaries of Hox gene expression in the central nervous system of the leech Helobdella. Developmental Biology 190: 284-300. Kulakova, M. A., Kostyuchenko, R. P., Andreeva, T. F. and Dondua, A. K. 2002. The Abdominal-B-like gene expression during larval development of Nereis virens (polychaeta). Mechanisms of Development 115: 177-179. Kuo, D. H. and Shankland, M. 2004. A distinct patterning mechanism of O and P cell fates in the development of the rostral segments of the leech Helobdella robusta: implications for the evolutionary dissociation of developmental pathway and morphological outcome. Development 131: 105-15. Lambert, J. D. and Nagy, L. M. 2003. The MAPK cascade in equally cleaving spiralian embryos. Developmental Biology 263: 231-41.

Early Annelid Development, A Molecular Perspective

137

Lans, D., Wedeen, C. J. and Weisblat, D. A. 1993. Cell lineage analysis of the expression of an engrailed homolog in leech embryos. Development 117: 857-871. Leptin, M., Casal, J., Grunewald, B. and Reuter, R. 1992. Mechanisms of early Drosophila mesoderm formation. Development Supplement 2: 23-31. Lillie, F. R. 1902. Differentiation without cleavage in the egg of the annelid Chaetopterus pergamentaceous. Archiv für Entwicklungsmechanik der Organismen 14: 477-499. Lillie, F. R. 1906. Observations and experiments concerning the elementary phenomena of embryonic development in Chaetopterus. Journal of Experimental Zoology 3: 153-268. Lillie, F. R. 1909. Polarity and bilaterality of the annelid egg. Experiments with centrifugal force. Biological Bulletin 16: 54-79. Lohmann, I. and McGinnis, W. 2002. Hox genes: It’s all a matter of context. Current Biology 12: R514-R516. Manak, J. R. and Scott, M. P. 1994. A class act: conservation of homeodomain protein functions. Development 1994 Supplement 61-77. Martindale, M. Q. and Shankland, M. 1990. Intrinsic segmental identity of segmental founder cells of the leech embryo. Nature 347: 672-674. Maslakova, S. A., Martindale, M. Q. and Norenburg, J. L. 2004. Vestigal prototroch in a basal nemertean, Carinoma tremaphoros (Nemertea; Palaeonemertea). Evolution and Development 6: 219-226. Master, V. A., Kourakis, M. J. and Martindale, M. Q. 1996. Isolation, characterization, and expression of Le-msx, a maternally expressed member of the msx gene family from the glossiphoniid leech, Helobdella. Developmental Dynamics 207: 404-419. McHugh, D. 1997. Molecular evidence that echiurans and pogonophorans are derived annelids. Proceedings of the National Academy of Sciences USA 94: 80068009. Mead, A. D. 1897. The early development of marine annelids. Journal of Morphology 13: 227-326. Morgan, T. H. 1910a. Cytological studies of centrifuged eggs. Journal of Experimental Zoology 9: 593-656. Morgan, T. H. 1910b. The effects of altering the position of the cleavage planes in eggs with precocious specification. Archiv für Entwicklungsmechanik der Organismen 29: 205-224. Nardelli-Haefliger, D., Bruce, A. E. E. and Shankland, M. 1994. An axial domain of HOM/Hox gene expression is formed by morphogenetic alignment of independently specified cell lineages in the leech Helobdella. Development 120: 1839-1849. Nardelli-Haefliger, D. and Shankland, M. 1992. Lox2, a putative leech segment identity gene, is expressed in the same segmental domain in different stem cell lineages. Development 116: 697-710. Nardelli-Haefliger, D. and Shankland, M. 1993. Lox10, a member of the NK-2 homeobox gene class, is expressed in a segmental pattern in the endoderm and the cephalic nervous system of the leech Helobdella. Development 118: 877-892. Nelson, B. H. and Weisblat, D. A. 1991. Conversion of ectoderm to mesoderm by cytoplasmic extrusion in leech embryos. Science 253: 435-438. Nelson, B. H. and Weisblat, D. A. 1992. Cytoplasmic and cortical determinants interact to specify ectoderm and mesoderm in the leech embryo. Development 115: 103-115. Novikoff, A. B. 1936. Transplantation of the polar lobe in Sabellaria vulgaris. Anatomical Abstracts Philadelphia 67: 57.

138 Reproductive Biology and Phylogeny of Annelida Okada, K. 1957. Annelida. Pp. 192-241. In M. Kumé and K. Dan (eds.) Invertebrate Embryology. NOLIT, Belgrade. Penners, A. 1922. Die Furchung con Tubifex rivulorum. Lam. Zool. Jb. Anat. Ont. 45: 323-368. Penners, A. 1924. Die Entwicklung des Keimstreifs und die Organbildung bei Tubifex rivulorum. Lam. Zoologische Jahrbücher, Abteilung für Anatomie und Ontogenie der Tiere 45: 251-308. Peterson, K. J., Irvine, S. Q., Cameron, R. A. and Davidson, E. H. 2000. Quantitative assessment of Hox complex expression in the indirect development of the polychaete annelid Chaetopterus sp. Proceedings of the National Academy of Sciences USA 97: 4487-4492. Peterson, M. D., Rogers, B. T., Popadic, A. and Kaufman, T. C. 1999. The embryonic expression pattern of labial, posterior homeotic complex genes and the teashirt homologue in an apterygote insect. Development Genes and Evolution 209: 7790. Pilon, M. and Weisblat, D. A. 1997. A nanos homolog in leech. Development 124: 1771-1780. Prince, V. E., Joly, L., Ekker, M. and Ho, R. K. 1998. Zebrafish hox genes: genomic organization and modified colinear expression patterns in the trunk. Development 125. Prud’homme, B., Lartillot, N., Balavoine, G., Adoutte, A. and Vervoort, M. 2002. Phylogenetic analysis of the Wnt gene family. Insights from lophotrochozoan members. Current Biology 12: 1395. Prud’homme, B., de Rosa, R., Arendt D., Julien J. F., Pajaziti R., Dorresteijn A., Adoutte A., Wittbrodt J., and Balavoine, G. 2003. Arthropod-like expression patterns of engrailed and wingless in the annelid Platynereis dumerilii suggest a role in segment formation. Current Biology 13: 1876-1881. Pultz, M. A., Diederich, R. J., Cribbs, D. L. and Kaufman, T. C. 1988. The proboscipedia locus of the Antennapedia complex: a molecular and genetic analysis. Genes and Development 2: 901-920. Render, J. A. 1983. The second polar lobe of the Sabellaria cementarium embryo plays an inhibitory role in apical tuft formation. Roux’s Archives of Developmental Biology 192: 120-129. Reverberi, G. 1971. Annelids. Pp. 126-163. In G. Reverberi (ed.) Experimental Embryology of Marine and Fresh-water Invertebrates. North-Holland, Amsterdam. de Rosa, R., Grenier, J. K., Andreeva, T., Cook, C. E., Adoutte, A., Akam, M., Carroll, S. B. and Balavoine, G. 1999. Hox genes in brachiopods and priapulids and protostome evolution. Nature 399: 772-776. Rouse, G. 1999. Trochophore concepts: ciliary bands and the evolution of larvae in spiralian Metazoa. Biological Journal of the Linnean Society 66: 411-464. Rouse, G. 2000. The epitome of hand waving? Larval feeding and hypotheses of metazoan phylogeny. Evolution and Development 2: 222-233. Sandig, M. and Dohle, W. 1988. The cleavage pattern in the leech Theromyzon tessulatum (Hirudinea, Glosiphoniidae). Journal of Morphology 196: 217-252. Savage, R. and Shankland, M. 1996a. Identification and characterization of a hunchback orthologue, Lzf2, and its expression during leech embryogenesis. Developmental Biology 175: 205-217. Scholtz, G. 2002. The Articulata hypothesis — or what is a segment? Organisms, Diversity and Evolution 2: 197-215. Seaver, E. C. 2003. Segmentation: mono- or polyphyletic? International Journal of Developmental Biology 47: 583-95.

Early Annelid Development, A Molecular Perspective

139

Seaver, E. C., Paulson, D. A., Irvine, S. Q. and Martindale, M. Q. 2001. The spatial and temporal expression of Ch-en, the engrailed gene in the polychaete Chaetopterus, does not support a role in body axis segmentation. Developmental Biology 236: 195-209. Seaver, E. C. and Shankland, M. 2000. Leech segmental repeats develop normally in the absence of signals from either anterior or posterior segments. Developmental Biology 224: 339-353. Seaver, E. C. and Shankland, M. 2001. Establishment of segment polarity in the ectoderm of the leech. Development 128: 1629-1641. Seaver, E. C., Thamm, K. and Hill, S. D. 2005. Growth patterns during segmentation in the two polychaete annelids, Capitella sp. I, Hydroides elegans: comparisons at distinct life history stages. Evolution and Development 7: 312-326. Shain, D. H., Ramirez-Weber, F.-A., Hsu, J. and Weisblat, D. A. 1998. Gangliogenesis in leech: morphogenetic processes leading to segmentation in the central nervous system. Development Genes & Evolution 208: 28-36. Shankland, M. 1987a. Differentiation of the O and P cell lines in the embryo of the leech I: Sequential commitment of blast cell sublineages. Developmental Biology 123: 85-96. Shankland, M. 1987b. Differentiation of the O and P cell lines in the embryo of the leech II: Genealogical relationship of descendant pattern elements in alternative developmental pathways. Developmental Biology 123: 97-107. Shankland, M. and Savage, R. M. 1997. Annelids, the Segmented Worms. Pp. 219236. In S. F. Gilbert and A. M. Raunio (eds) Embryology: Constructing the Organism. Sinauer, Sunderland, MA. Shearer, C. 1911. On the development and structure of the trochophore of Hydroides uncinatus (Eupomatus). Quarterly Journal of Microscopical Science, London 56: 543-591. Shimizu, T. 1995. Role of the cytoskeleton in the generation of spatial patterns in Tubifex eggs. Current Topics in Developmental Biology 31: 197-235. Shimizu, T. and Savage, R. M. 2002. Expression of hunchback protein in a subset of ectodermal teloblasts of the oligochaete annelid Tubifex. Development Genes and Evolution 212: 520-525. Snow, P. and Buss, L. W. 1994. HOM/Hox-type homeoboxes from Stylaria lacustris (Annelida: Oligochaeta). Molecular Phylogenetics and Evolution 3: 360-364. Song, M. H., Huang, F. Z., Chang, G. Y. and Weisblat, D. A. 2002. Expression and function of an even-skipped homolog in the leech Helobdella robusta. Development 129: 3681-92. Song, M. H., Huang, F. Z., Gonsalves, F. C. and Weisblat, D. A. 2004. Cell cycledependent expression of a hairy and Enhancer of split (hes) homolog during cleavage and segmentation in leech embryos. Developmental Biology 269: 183195. Soto, J. G., Nelson, B. H. and Weisblat, D. A. 1997. A leech homolog of twist: evidence for its inheritance as a maternal mRNA. Gene 199: 31-37. Soulier, A. 1902. Les premiers stades embryologiques de la Serpule. Travaux de l’Institut de zoologie de l’Université de Montpellier 9: 1-78. Storey, K. G. 1989. Cell lineage and pattern formation in the earthworm embryo. Development 107: 519-531. Tear, G., Akam, M. and Martinez-Arias, A. 1990. Isolation of an abdominal-A gene from the locust Schistocerca gregaria and its expression during early embryogenesis. Development 110: 915-925.

140 Reproductive Biology and Phylogeny of Annelida Telford, M. J. and Thomas, R. H. 1998. Expression of homeobox genes shows chelicerate arthropods retain their deutocerebral segment. Proceedings of the National Academy of Sciences USA 95: 10671-10675. Tessmar-Raible, K. and Arendt, D. 2003. Emerging systems: between vertebrates and arthropods, the Lophotrochozoa. Current Opinion in Genetics and Development 13: 331-340. Titlebaum, A. 1928. Artificial production of Janus embryo of Chaetopterus. Proceedings of the National Academy of Sciences USA 14: 245-247. Treadwell, A. L. 1901. The cytogeny of Podarke obscura (Verrill). Journal of Morphology 17: 399-486. Tyler, A. 1930. Experimental production of double embryos in annelids and mollusks. Journal of Experimental Zoology 57: 347-402. Verdonk, N. H. and Biggelaar, J. A. M. van den 1983. Early development and formation of the germ layers. Pp. 91-122. In N. H. Verdonk, J. A. M. van den Biggelaar and A. S. Tompa (eds.) The Mollusca. Academic Press, New York. 3. Weber, R. 1958. Uber die submikroskopische Organisation und die biochemische Kennzeichnung embryonaler Entwicklungstadien von Tubifex. Archiv für Entwicklungsmechanik der Organismen 150: 542-580. Wedeen, C. J. and Weisblat, D. A. 1991. Segmental expression of an engrailed-class gene during early development and neurogenesis in an annelid. Development 113: 805-819. Weisblat, D. A. and Blair, S. S. 1984. Developmental interdeterminacy in embryos of the leech Helobdella triserialis. Developmental Biology 101: 326-335. Weisblat, D. A. and Huang, F. Z. 2001. An overview of glossiphoniid leech development. Canadian Journal of Zoology 79: 218-232. Weisblat, D. A. and Shankland, M. 1985. Cell lineage and segmentation in the leech. Philosophical Transactions of the Royal Society of London B 312: 40-56. Weisblat, D. A. S. R. T. and Stent, G. S. 1978. Cell lineage analysis by intracellular injection of a tracer enzyme. Science 202: 1295-1298. Werbrock, A. H., Meiklejohn, D. A., Sainz, A., Iwasa, J. H. and Savage, R. M. 2001. A polychaete hunchback ortholog. Developmental Biology 235: 476-488. Whitman, C. O. 1878. Embryology of Clepsine. Quarterly Journal of Microscopical Science 18: 215-315. Wilson, E. B. 1890. The origin of the mesoblast-band in annelids. Journal of Morphology 4: 205-219. Wilson, E. B. 1892. The cell lineage of Nereis. A contribution to the cytogeny of theannelid body. Journal of Morphology 6: 361-466. Wilson, E. B. 1896. On cleavage and mosaic work. Archiv für Entwicklungsmechanik der Organismen 3: 19-26. Wilson, E. B. 1904. Mosaic development in the annelid egg. Science 20: 748-750. Wisely, B. 1958. The development and settling of a serpulid worm, Hydroides norvegica gunnerus (Polychaeta). Australian Journal of Marine and Freshwater Research 9: 351-361. Wong, V. Y., Aisemberg, G. O., Gan, W.-B. and Macagno, E. R. 1995. The leech homeobox gene Lox4 may determine segmental differentiation of identified neurons. Journal of Neuroscience 15: 5551-5559. Wong, V. Y. and Macagno, E. R. 1998. Lox6, a leech Dfd ortholog, is expressed in the central nervous system and in peripheral sensory structures. Development Genes and Evolution 208: 51-55. Wysocka-Diller, J., Aisemberg, G. O. and Macagno, E. R. 1995. A novel homeobox cluster expressed in repeated structures of the midgut. Developmental Biology 171: 439-447.

CHAPTER

6

Annelid Larval Morphology Greg W. Rouse

6.1.

INTRODUCTION

Larvae can be broadly defined as a structural state that occurs between the onset of the morphogenesis following embryonic development (cleavage, blastula, gastrula) and the metamorphosis to the adult body form (Hickman 1999). This covers larvae that are feeding or non-feeding, brooded by the parent in some form, or freely developing in the water. This definition also allows for larvae that go through a rapid or catastrophic metamorphosis to the adult form (e.g. Owenia) as well as those that gradually change into the adult (many directly-developing annelids). This is quite useful in the context of this chapter, since developing stages of annelids are marvellously variable in form. There have been previous comprehensive reviews that have highlighted this diversity (e.g. Schroeder and Hermans 1975; Bhaud and Cazaux 1982, 1987), and it is somewhat unfortunate that this diversity has not been generally appreciated. This may be because one particular (and rare) kind of annelid larval form, the ‘classical’ trochophore with opposed-band feeding (see below), tends to star in biology textbooks and in discussions on animal evolution (reviewed by Nielsen 1995). In this chapter the diversity of annelid larvae is reviewed, terminology outlined and the evolution of the various larval forms and ciliary bands is discussed.

6.2.

CILIATED BANDS OF LARVAE

Annelids show a wide range of ciliary tufts or bands. The following definitions and list of taxa are derived from Bhaud and Cazaux (1982; Bhaud and Cazaux 1987) and Rouse (1999; 2000c). The terms are discussed in the order of appearance on the larval body from anterior to posterior (see Fig. 6.1). Apical tuft. The apical organ, found in many animal larvae, is a group of sensory cells that subsequently become part of the brain (Nielsen 1995). The apical organ can often be identified by the presence of a group of cilia South Australian Museum Nth Terrace, Adelaide. S.A. 5000 Australia & Earth and Environmental Sciences, University of Adelaide SA. 5005 Australia

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Reproductive Biology and Phylogeny of Annelida

known as the apical tuft (Fig. 6.1). The distribution of the apical organ appears to be widespread and is present in most annelids where detailed anatomical studies have been done. The presence of an apical tuft is a guide as to the presence of an apical organ, but this is potentially misleading since for instance, Polygordiidae that have an apical organ, lack an apical tuft, except in the very earliest stages (Woltereck 1904). The apical tuft also tends to disappear relatively early in development of other polychaetes. An apical tuft is not found in clitellates and also appears to be absent in some Cirratulidae, Histriobdellidae, Lopadorhynchidae, Orbiniidae, Sabellidae, and Tomopteridae (see Rouse 1999). Akrotroch. Häcker (1896) introduced the term akrotroch to describe a ciliated region between the prototroch and the apical organ. An akrotroch is defined here as any ring of cilia that lies anterior to the prototroch (i.e., on the episphere), and is not associated with the apical organ. An akrotroch can be distinguished from a meniscotroch (see below) in being a complete ring

Fig. 6.1. Diagrammatic representation of a ventral view of an annelid larva showing most ciliary bands described in this paper. Note that no larval form has ever been described with all of these bands. Abbreviations: a apical tuft, ak akrotroch, b buccal opening (actual or incipient), f ciliated feeding groove, m metatroch, me meniscotroch, ms mesotroch, n neurotroch, o oral brush, p prototroch, t telotroch. Not shown are gastrotroch or nototroch. These are segmental ciliary bands (as the mesotroch appears to be; see text). Original.

Annelid Larval Morphology

"!

around the episphere (Figs. 6.1, 6.4). Akrotrochs have only been described in the larvae of annelids and have been found in Cirratulidae, Dorvilleidae, Eunicidae, Lumbrineridae, Onuphidae, Orbiniidae and Syllidae (Rouse 1999). Meniscotroch. Bhaud and Cazaux (1982) defined a meniscotroch as a crescent-shaped area of short cilia on the episphere. They further distinguished it by the cilia of the central part of the meniscotroch being longer, bent and forming a pointed brush (Bhaud and Cazaux 1982). In contrast, Lacalli (1980) distinguished this particular ciliated patch as being separate from the meniscotroch, and called it the frontal organ. This is not accepted here. Bhaud and Cazaux (1982) regarded a meniscotroch as being absent in families such as Hesionidae, Polynoidae, Pholoidae and Sigalionidae, even though they have a ciliated patch or patches on the episphere. Rouse (1999) used the more general definition with a further refinement in that the meniscotroch is always on the ventral surface of the episphere, in line with the mouth and does not need to a central pointed brush (Figs. 6.1, 6.4). A meniscotroch following this definition, is known from taxa in 13 polychaete families, all of them members of Phyllodocida; Aphroditidae, Chrysopetalidae, Glyceridae, Goniadidae, Hesionidae, Lopadorhynchidae, Nephtyidae, Pholoidae, Phyllodocidae, Pilargidae, Pisionidae, Polynoidae, Sigalionidae and possibly Nereididae (see Rouse 1999). The meniscotroch is potentially used in feeding (B. Pernet pers. comm.). Prototroch. The term prototroch, a name coined by Kleinenberg (1886), is a single equatorial ring of (usually compound) cilia that is derived from a group of specific cells, called trochoblasts (Damen and Dictus 1994) and lies anterior to the mouth (see Chapter 5). While there is variation in the specific cellular composition of the prototroch, it is essentially the same in annelids, entoprocts, molluscs, and sipunculans. Embryological studies have identified at least primary trochoblasts (embryonic cells 1a2-1d2) in 12 polychaete families (see Rouse 1999). While for most polychaetes no detailed cell lineage studies have been performed to assess whether a true prototroch exists it can be reasonably assumed that a distinct ciliated band in the immediately preoral region of a larvae corresponds to a prototroch (Figs. 6.1, 6.2, 6.3, 6.4, 6.7, 6.8, 6.9). The prototroch divides the larva into an anterior portion, the episphere, and a posterior portion, the hyposphere (Bhaud and Cazaux 1982). Annelids where a prototroch can said to be absent are Clitellata (Fig. 6.6), Aeolosomatidae and Histriobdellidae. Further investigation of Chaetopteridae, Myzostomida and Sternaspis is also warranted (Rouse 1999). While some authors studying development of clitellates have identified trochoblasts (see reviews by Dawydoff 1959; Needham 1990), others including Anderson (1966; 1971) and Jamieson (Chapter 8, this volume), do not give this credence. Metatroch. A metatroch has been defined as a post-oral band of cilia that beats in a posterior to anterior direction, in opposition to the prototroch (Nielsen 1995; Strathmann 1993). Rouse (1999) expanded the definition with

"" Reproductive Biology and Phylogeny of Annelida

Fig. 6.2. Classical opposed-band trochophore of serpulid polychaetes. A. Differential interference contrast micrograph of Spirobranchus giganteus trochophore showing complete gut. B. Scanning Electron Micrograph (SEM) of Spirobranchus giganteus trochophore in side view showing apical tuft and prototroch. C. Scanning Electron Micrograph (SEM) of Spirobranchus giganteus trochophore in posterior view showing prototroch, metatroch and ciliated food groove. Arrow indicates mouth. Original. Abbreviations: a, apical tuft; e, eye; g, gut; m, metatroch; p, prototroch.

reference to annelids, to encompass any ciliary band that lies behind the mouth (Figs. 6.1, 6.2, 6.3, 6.4), but is still presegmental (i.e., is situated on the peristomium). A variety of annelids are stated to have a metatroch in the traditional sense (i.e. associated with food gathering): Amphinomidae (Fig. 6.3E), Echiura (Fig. 6.3A; now considered annelids, see Chapter 1), Opheliidae (Fig. 6.3B), Oweniidae (Fig. 6.3D), Polygordiidae (Figs. 6.6A, B, 6.9), Sabellariidae (Fig. 6.3C) (Pernet 2003), and Serpulidae (Fig. 6.2), though it should be noted that larvae of some Echiura (Fig. 6.5A) and Opheliidae lack a metatroch (Fig. 6.4E) and there is some doubt about Amphinomidae (Rouse 2000a). It would appear that the metatroch in all of these taxa are peristomial structures. Some Saccocirridae (Fig. 6.3F) may have members that use the metatroch for feeding (Pierantoni 1906), but further investigation is required. Additionally, there are a number of other polychaete groups that have a metatroch, but it is not used in feeding. These are: Ampharetidae, Dorvilleidae, Eunicidae (Fig. 6.5C), Lumbrineridae, Myzostomida (possibly not annelids, see Chapter 1), Onuphidae, Orbiniidae

Annelid Larval Morphology

"#

Fig. 6.3. Other opposed-band annelid trochophores. A. Trochophore of Echiuris sp.. Modified from Hatschek, B. 1881. Arbeiten aus dem Zoologischen Institute der Universität Wien und der Zoologischen Station in Triest 3: 1-34, Fig. 12. Feeding has been shown to occur via an opposed-band system in some larval Echiura, others are lecithotrophic. B. Five segment larva of Armandia brevis (Opheliidae). Modified from Hermans, C. O. 1978. Pp. 113-126. In F.-S. Chia and M. E. Rice (eds), Settlement and Metamorphosis of Marine Invertebrate Larvae, Elsevier, New York, Fig. 4.. This opheliid feeds via an opposed-band system (Miner et al. 1999); others are lecithotrophic or use some other feeding mechanism. C. Trochophore of Sabellaria alveolata (Sabellariidae). Modified from Wilson, D. P. 1929. Journal of the Marine Biological Association of the United Kingdom 16: 221-268, Fig. 1.8. Sabellariids use an opposedband feeding system (see Pernet 2003). D. Larva of Owenia fusiformis (Oweniidae) that has been shown to use opposed-band feeding. Modified from Wilson, D. P. 1932b. Philosophical Transactions of the Royal Society of London. Series B 221: 231-334, Fig. 29.1. E. Upper diagram shows a dorsal view of ‘rostraria’ larva of an amphinomid polychaete with purported feeding tentacles. Modified from Sveshnikov, V. A. 1978. Morphology of larval polychaetes, Akademiia Nauk SSSR, Moscow, 1-151, Fig. 1E. The lower diagram shows a frontal view of a rostraria larva with ciliary rows on feeding tentacles indicating opposedband feeding occurs. Arrows indicate path of food taken from parental tentacles on its way to the buccal opening. Modified from Jägersten, G. 1972. Academic Press, New York, 282, Fig. 45. Other amphinomids do not have feeding larvae. F. Ventral view of larva of Saccocirrus papillocercus. Modified from Pierantoni, U. 1906. Mittheilungen aus der Zoologischen Station zu Neapel 18: 46-72, Fig. 28. Saccocirridae may have members that use the metatroch for feeding. Other saccocirrid larvae feed using extensible oral arms. Abbreviations: an, anus; b, buccal opening; c, chaetae; e, eye; g, gut; m, metatroch; me, meniscotroch; p, prototroch.

"$ Reproductive Biology and Phylogeny of Annelida (Fig. 6.4F), Protodrilidae, Sabellidae and some Serpulidae. Other polychaetes have erroneously been cited as having a metatroch, or an opposed-band ciliary feeding system (and hence implying the presence of a metatroch). These are Capitellidae (Figs. 6.5C, 6.8), Siboglinidae (Figs. 6.4J, 6.10) and Syllidae (Fig. 6.9) (Rouse 2000a). Oral brush. An oral brush is a bundle of long cilia at the posterior base of the prototroch, on the left side of the mouth (Fig. 6.1), of larvae of the scaleworm taxa Pholoidae (Fig. 6.9) Polynoidae (Figs. 6.4H, 6.9) and Sigalionidae (Fig. 6.9) (Phillips and Pernet 1996). The oral brush extends posteriorly, and is used to help ingest particles up to 60 µm in diameter. It appears that the brush is an alternative way of facilitating feeding in polychaete larvae that are planktotrophic, but lack a metatroch (Phillips and Pernet 1996). Ciliated food groove. Where the metatroch has been shown to be involved in food gathering, there is often a third ciliary band between the prototroch and metatroch (Figs. 6.1, 6.2). This is commonly called the ciliated food groove (Strathmann 1993; Nielsen 1995) and acts to transport food. Among annelids it is present in larval sabellariids (Pernet 2003) oweniids, and some echiurids and serpulids. Amphinomids and polygordiids may also have this feature. Neurotroch. The term neurotroch was introduced by Gravely (1909) to identify a ventral longitudinal band of cilia running from behind the mouth to near the anus (Fig. 6.1). It is sometimes referred to as a gastrotroch, but this term is used to describe ventral transverse ciliary rings on segments in polychaete larvae (Bhaud and Cazaux 1982). The occurrence of a neurotroch is sporadic among annelids. It is present in at least some members of 30 polychaete families, including Siboglinidae, and has been shown to be absent in 19 others (Rouse 1999, 2000a) as well as Clitellata. Mesotroch. This refers to a complete transverse ciliated ring (or rings) on the middle of the larval body (Bhaud and Cazaux 1982). The definition makes no distinction between the presegmental region or segmental region and so it is difficult to see how they can be distinguished from gastrotrochs and nototrochs. Recent use of the term mesotroch can be found in Irvine et al. (1999) with reference to Chaetopterus larvae and here the ciliary bands are clearly segmental in later larvae (Figs. 6.5H), though initially it may appear non-segmental (Fig. 6.4A) and this was erroneously interpreted by Rouse (1999) as a possible metatroch (see Chapter 13). Gastrotroch and nototroch. Transverse ciliary rings on segments in polychaete larvae (Figs. 6.4B, F, 6.5G). Gastrotrochs are ventral and nototrochs are dorsal (Bhaud and Cazaux 1982). These terms are usually applied to spioniform larvae. Telotroch. A telotroch is a ring of cilia lying at the posterior of a larva (Fig. 6.1) that appears to have a locomotory function (Strathmann 1993). A telotroch is present in most annelid larvae. It is generally present where larval development is known, though it often appears in larvae at a rather late stage of development. It has not been shown in any developmental

Annelid Larval Morphology

"%

studies on Aeolosomatidae, Chaetopteridae, Clitellata, Glyceridae, Hesionidae, Histriobdellidae, Lopadorhynchidae, Magelonidae, Pisionidae, Sabellidae, Serpulidae, Sternaspis or Tomopteridae.

6.3 TERMS TO DESCRIBE ANNELID LARVAE The following terms used to describe the larvae of annelids has largely been adopted and modified from Bhaud and Cazaux (1982; 1987). Further work is needed on refining these terms to more useful for example in properly represent homology statements. Trochophore. The term trochophore has had a variety of definitions and was reviewed in Rouse (1999), who broadly defined it as any larvae with a prototroch. Throughout this chapter, where name trochophore is used, it refers to the general definition. In contrast to this broad definition, trochophore is often used in a strict sense for larvae having an opposed-band method of feeding, involving the ciliary bands prototroch and metatroch (e.g. Hatschek 1878; Nielsen 1995). This classical ‘opposed-band trochophore’ (Figs. 6.2A, B, C, 6.3A) has been proposed to represent the plesiomorphic larval form for a large group of animals (e.g., Annelida, Mollusca, Sipuncula) and this was supported in the analysis by Rouse (1999), but only in the broad meaning of the word. The suggestion that the ‘opposed-band trochophore’, in the sense of Hatschek and Nielsen, is plesiomorphic for Annelida or a more inclusive group of animals has been evaluated a number of times (Ivanova-Kazas 1985a, 1985b, 1985c; Haszprunar et al. 1995; Rouse 1999). These authors have pointed out the wide diversity of annelid larval forms and suggested that the ‘opposed-band trochophore’ is a derived larval type. In a few cases among annelids (e.g., clitellates), it would appear that the prototroch has been lost. This means that the larvae may be referred to as modified trochophores, since in a phylogenetic sense they still are trochophore larvae. Two additional terms based on trochophore are often found in the literature; Protrochophore and metatrochophore. Protrochophore. This is one where the larva is completely or nearly completely ciliated (Figs. 6.4A, 6.5F). These larvae can then develop more restricted ciliary bands (Fig. 6.4C), or maintain a broad area of cilia (e.g., many Eunicida). Metatrochophore. This is the stage after the normal trochophore stage, where there are clear signs of segmentation apparent. If the parapodia are not formed then it is referred to as stage I (Figs. 6.3B, E, 6.4B, C, D, F, 6.5B) if there are parapodia present but not functioning then it is stage 2. Once the parapodia are functioning the larva has left the metatrochophore stage (Bhaud and Cazaux 1982). Aulophore. This is a larval form found only in terebelliforms such as Terebellidae (Fig. 6.5I) and Pectinariidae (Fig. 6.5J), and is a metatrochophore living in a tube (Bhaud and Cazaux 1982). Chaetosphaera. This is a larval form first described by Häcker (1898) and is usually applied to well-developed larvae of some spioniforms (Fig. 6.5G)

"& Reproductive Biology and Phylogeny of Annelida

Fig. 6.4. Other annelid trochophores showing a variety of ciliary band complements. A. Lateral view of 40-hour-old larva of Chaetopterus pergamentaceus (Chaetopteridae) showing apical tuft, possible prototroch and mesotroch. Modified from Wilson, E. B. 1883. Studies from the Biological Laboratory, Johns Hopkins University 2: 271-299, Fig. 82. B. Ventral view of 4-day- old larva of Cirriformia spirabrancha (Cirratulidae) showing akrotroch, prototroch, buccal opening, gastrotroch, neurotroch and telotroch. Modified from Blake, J. A. 1975c. Transactions of the American Microscopical Society 94: 179-188, Fig. 2. C. Ventral view of three-chaetiger larvae of Eunice valens (Eunicidae). Modified from Åkesson, B. 1967a. Acta Zoologica 48: 141-192, Fig. 1, showing akrotroch, prototroch, metatroch, neurotroch and telotroch. D. Lateral view of 5day-old larva of Clymenella torquata (Maldanidae) showing apical tuft, prototroch, neurotroch, telotroch and chaeta on first segment. Modified from Newell, G. E. 1951. Proceedings of the Zoological Society of London 121: 561-586, Fig. 17. E. Ventral view of 3-day-old larva of Ophelia bicornis showing apical tuft, prototroch, neurotroch and telotroch. Modified from Wilson, D. P. 1948. Journal of the Marine Biological Association of the United Kingdom 27: 540-553, Fig. 1F. F. Dorsal view of an early larva of Leitoscoloplos pugettensis (Orbiniidae) showing akrotroch, prototroch, probable metatroch, telotroch and segmental ciliary Fig. 6.4 contd

Annelid Larval Morphology

"'

that can swim with undulating movements (as well as via cilia). They can also roll up in a ball forcing their long spiny chaetae outwards as protective array (Bhaud and Cazaux 1982). Similar behaviour, also arguably defensive (or for flotation), is found in sabellariid, chrysopetalid and myzostome larvae, though these larvae tend not to be long enough to curl into a ball. Erpochaete. This is a creeping larval stage moving on or in the sediment using its chaetae (Bhaud and Cazaux 1982). The term is not commonly used since most authors would refer to these stages as young juveniles. Mitraria. The distinctive larval form found only in Oweniidae (Figs. 6.3D, 6.6C, D). After a ‘normal’ trochophore phase the larva becomes a planktotrophic mitraria. Wilson (1932b) elegantly described the development of the mitraria phase and its ‘catastrophic metamorphosis’ that results in much of the larval body being cast-off and a juvenile worm settling to the bottom (Fig. 6.6E). Larvae similar to the mitraria of Owenia have been described for Myriochele (Thorson 1946). Rostraria. Rostraria larvae almost certainly are those of amphinomids and euphrosinids (Mileikovsky 1960, 1961). The larvae (Figs. 6.3E, 6.5D) unlike the adults, have a pair of distinctive tentacle that are apparently used for feeding via ciliary bands (Jägersten 1972). Excellent micrographs of these larvae can be found in Pernet et al. (2001). Nectochaete. Larval stage with functional parapodia (Bhaud and Cazaux 1982); usually applied to larvae within Phyllodocida (Fig. 6.5E). Nectosoma. Similar to chaetosphaera larvae in that the larvae swim by undulation these larvae cannot roll up into a ball (Bhaud and Cazaux 1982); usually applied to larvae of Poecilochaetus.

6.4 LECITHOTROPHY AND LARVAL FEEDING Across the breadth of annelid diversity the group generally shows larval development where yolk reserves provided in the egg allow development of the larvae through to the juvenile stage. This is referred to as lecithotrophy Fig. 6.4 contd

band that is possibly a nototroch. Modified from Blake, J. A. 1980. Ophelia 19: 1-18, Fig. 1. G. Ventral view of early larva of Anaitides williamsi (Phyllodocidae) showing apical tuft, meniscotroch, prototroch buccal opening and neurotroch. Modified from Blake, J. A. 1975b. Ophelia 14: 23-84, Fig. 8A. The feeding mechanisms of this group have yet to be elucidated. H. Lateral view of an early larva of Harmothoe longosetis (Polynoidae) showing probable apical tuft, meniscotroch, prototroch, buccal opening, oral brush and neurotroch.Modified from Cazaux, C. 1968. Archives de Zoologie Expérimentale et Générale 109: 477-543, Fig. V.1. I. Lateral view of Pectinaria koreni (Pectinariidae) trochophore showing apical tuft, prototroch that has posterior extensions on either side of the buccal opening, neurotroch and telotroch. Modified from Sveshnikov, V. A. 1978. Akademiia Nauk SSSR, Moscow, 1-151, Fig. 39. J. Lateral view of early larval stage of Ridgeia sp. (Vestimentifera, Siboglinidae). Modified from Southward, E. C. 1988. Journal of the Marine Biological Association of the United Kingdom 68: 465-487, Fig. 4. K. Ventral view of early larva of Scolecolepis fuliginosa (Spionidae) showing apical tuft, prototroch and telotroch. Modified from Day, J. H. 1934. Journal of the Marine Biological Association of the United Kingdom 19: 633-654, Fig. 4. Note the lack of a metatroch but this larva does use some, as yet undescribed, form of feeding.

# Reproductive Biology and Phylogeny of Annelida

Fig. 6.5 contd

Annelid Larval Morphology

#

or lecithotrophic development. This means that the eggs have to be relatively large (100 µm or greater) in order for the larva to develop. No clitellates show larval feeding independent of the parent, though many develop from small eggs and require albumin in the cocoon to complete development (Needham 1990). Some leeches are exceptional in that the parents do provide the juveniles with food (Kutschera and Wirtz 1986), but most often larval feeding is thought of in terms of marine larvae feeding in the plankton. This is referred to as planktotrophy, or planktotrophic development. Polychaetes are generally thought of as having small eggs (60-80 µm) that give rise to planktotrophic larvae that must feed in order to become juveniles, but a comprehensive survey of the group reveals that this is in fact not that common. In a review of 306 species for which the life cycle is known Wilson (1991) listed only 79 that exhibited this form of reproduction. A further 44 species show external fertilisation and lecithotrophic or direct developing larvae. The remaining 183 polychaetes have some form of brooding. Of course, this is by no means an indication of the real proportions of the various reproductive modes among polychaetes; it only reflects our present state of knowledge. The life cycles of most species are still unknown. A planktotrophic larval form using an ‘opposed-band’ or ‘downstream’ system is fundamental to the restricted definition of a trochophore, according to Nielsen (1995). This requires the use of the prototroch to generate a current for locomotion and feeding, and the metatroch creates a beat in the opposite direction (Strathmann 1993). The ciliated food groove, made up of short cilia between the prototroch and metatroch, moves captured food to the mouth. The presence of a prototroch and metatroch (ie, opposed-bands) has led Strathmann (1987; 1993) to suggest that among annelids amphinomids, echiurids, oweniids, polygordiids, sabellariids, serpulids and opheliids, may feed in a similar manner. All other polychaete groups do not have opposed-band larval feeding, with the possible exception of Saccocirridae. Fig. 6.5 contd

Fig. 6.5. SEMs of annelid larvae at various stages showing a variety of ciliary bands. All larvae were collected from the plankton off Belize, with the exception of Metabonellia haswelli, Arenicola sp. and Marphysa sp., and so taxon names are difficult to assign. A. Ventro-lateral view of a lecithotrophic trochophore of the Metabonellia haswelli (Echiura) taken from a jelly mass in Sydney Harbour. Note the lack of a metatroch. B. Ventro-lateral view of a metatrochophore taken from a jelly mass of Arenicola sp. (Arenicolidae) from Belize. C. Lateral view of a capitellid metatrochophore larva with a series of thoracic segments; some of them with chaetae. D. Dorsal view of an amphinomid rostraria larva before it has developed the pair of characteristic feeding tentacles. These appear to be developing just behind the nuchal organs. E. Dorsal view of a glycerid nectochaete with 6 chaetigers. F. Ventral view of a Marphysa sp. (Eunicidae) protrochophore with one pair of chaetae appearing behind the prototroch. G. Lateral view of a spionid chaetosphaera larva. This larva has a single gastrotroch at this stage. H. Lateral view of an advanced chaetopterid larva. This larva has a single mesotroch at this stage and it is clearly a segmental ring of cilia. I. Ventrolateral view of a terebellid aulophore larva removed from its tube. J. Dorsal view of a pectinariid aulophore larva removed from its tube. Original.

#

Reproductive Biology and Phylogeny of Annelida

Fig. 6.6. Annelid larvae that show catastrophic metamorphosis. A. Frontal view of a Polygordius exolarva taken from the plankton off South Australia showing prostomium with two eyespots inside the larval episphere. B. Posterior view of the same larva showing the juvenile segmented body emergent from the larval episphere and hyposphere. C. Lateral view of an oweniid mitraria larva taken from plankton off Belize showing long larval chaetae and episphere with ciliated margin. D. Closeup of mitraria episphere showing juvenile inside larval body. E. Juvenile immediately after metamorphosis (taken only a few seconds after the previous micrograph). Larval chaetae have been shed, as has the episphere. Original.

Annelid Larval Morphology

#!

Where planktotrophic larvae have been found in other polychaete groups, such as Capitellidae, Chaetopteridae, Magelonidae (as young larvae), Pectinariidae, Phyllodocida, Protodrilidae, Saccocirridae (some), and Spionidae, they utilise other forms of particle capture. The larvae of chaetopterids, pectinariids, pisionids and some spionids utilise some form of mucous feeding that may involved currents generated by the prototroch (Nozais et al. 1997; Pernet 2004; R.R. Strathmann 1987; Werner 1953; Åkesson 1961). The larvae of protodrilids and some saccocirrids use an eversible pharynx (Jägersten 1952; Sasaki and Brown 1983) and are encounter predators. Where larval feeding has been found in Capitellidae, Magelonidae (young), Phyllodocida and many Spionidae it appears that it involves the use of the prototroch. It cannot be classified as opposed-band downstream larval feeding since members of all these taxa lack a metatroch. This led Rouse (2000a) to use a more general definition of downstream larval feeding that accommodated these other feeding methods. This assumed that any feeding method that involved a downstream current generated by the prototroch for feeding was homologous. By expanding the concept of downstream feeding beyond that of just opposed-band feeding, the global homology of this larval feeding mode is greatest. This allowed a robust assessment of the hypothesis that feeding larvae may be primitive, as proposed by Strathmann (1993). The results of this study will be outlined in a later section. Based on the broad definition by Rouse (2000a), major annelid groups that have been found to only have downstream-feeding larvae are Amphinomidae, Chrysopetalidae, Glyceridae, Nephtyidae, Oweniidae, Pectinariidae, Polynoidae, and Sabellariidae. On the other hand, Capitellidae, Echiura (Fig. 6.5A), Hesionidae, Magelonidae, Opheliidae, Pholoidae, Phyllodocidae, Serpulidae, and Spionidae also have members with larvae that do not feed (or are lecithotrophic until becoming encounter predators) and hence are lecithotrophic (Table 6.1). Rouse (2000a) assumed lecithotrophy to be secondary in each case; i.e., the presence of downstreamfeeding is assumed to be plesiomorphic in each polymorphic group. This assumption that feeding is always plesiomorphic will have to be tested using more detailed cladistic analyses at the appropriate levels and may well be shown to be wrong in some cases. For example, in a preliminary analysis on the evolution of reproduction in Serpulidae, Rouse and Fitzhugh (1994) suggested that feeding larvae in this group may be secondary; a result that has recently been supported by a more detailed analysis by Kupriyanova (2003). What will perhaps be surprising to some is the incidence of groups of annelids where no feeding larvae have been found at all. These are Alvinellidae, Ampharetidae, Arenicolidae (Fig. 6.5B), Cirratulidae, Dorvilleidae, Eunicidae (6.5D), Flabelligeridae, Goniadidae, Histriobdellidae, Lumbrineridae, Maldanidae, Myzostomida, Nereididae, Onuphidae, Orbiniidae (including Questidae), Protodriloididae, Psammodrilidae, Sabellidae, Siboglinidae, Sternaspis, Syllidae,

Aciculata Eunicida

Aciculata Amphinomida

Scolecida

Clitellata

Development studies

Dorvilleidae Eunicidae Hartmaniellidae Histriobdellidae Lumbrineridae

(Richards 1967; Åkesson 1973a, 1973b) (Richards 1967; Åkesson 1967a) Nothing known (Haswell 1916; Shearer 1910) (Bhaud and Cazaux 1987; Cazaux 1972; Okuda 1946; Richards 1967; Sato et al. 1982)

(Needham 1990) (Anderson 1973) (Bergter et al. 2004) Arenicolidae (Guberlet 1933; Newell 1948, 1949) (Bailey-Brock 1984) (Farke and Berghuis 1979; Watson and Bentley 1998; this study) Capitellidae (Eckelbarger and Grassle 1987; Eisig 1899; Hansen 1993; Wilson 1933) Maldanidae (Bookhout and Horn 1949; Cazaux 1972; Newell 1951; Rouse 1992) Cossuridae Nothing known Opheliidae (Dales 1952b; Guérin 1971, 1973; Hermans 1978; Miner et al. 1999; Wilson 1948) Orbiniidae (Anderson 1959, 1961; Blake 1980; Giangrande 1991) Paraonidae (Bhaud 1983) Rouse (1999, 2000a,c) dubiously regarded Fewkes (1883) as showing a paraonid larva Questidae (Giere and Riser 1981) Scalibregmatidae Nothing known Amphinomidae (Jägersten 1972; Kudenov 1974; Kudenov 1977; Marsden 1960; Mileikovsky 1961; Pernet et al. 2001) Euphrosinidae (Mileikovsky 1960)

Less inclusive taxa

None to date None to date

Not known; has tentacles like amphinomid larvae None to date None to date

Sometimes; opposed bands in tentacles

None to date

Some; opposed bands, encounter predation and unknown mechanisms None to date None to date; pelagic juveniles seen

None to date Sometimes; mechanisms unknown see (Rouse 2000a) None to date

None to date

Planktotrophy

Table 6.1 Developmental Studies on Annelida. This is not comprehensive, so as to save space. See other chapters in this volume and major reviews (Giangrande 1997; Pernet et al. 2001; Rouse 1999, 2000a; Wilson 1991). Mechanisms of larval feeding are briefly described. If ‘None’ or ‘Sometimes’, it means that lecithotrophy is occurring in the group.

#" Reproductive Biology and Phylogeny of Annelida

Aciculata Phyllodocida

Aciculata Eunicida

Paralacydonia Pholoidae

Nereididae

Ichthyotomidae Lacydoniidae Lopadorhynchidae Myzostomida (Note this placement is controversial) Nautiliniellidae Nephtyidae

Eulepethidae Glyceridae Goniadidae Hesionidae

Acoetidae Aphroditidae Chrysopetalidae

Oenonidae Onuphidae

None to date Yes; meniscotroch and possibly prototroch

None to date

(Blake 1975b; Dales 1950; Mazurkiewicz 1975; Reish 1957; Wilson 1932a; Zottoli 1999) (Bhaud 1967) (Cazaux 1968; Heffernan and Keegan 1988a, 1988b; Laubier 1975; Phillips and Pernet 1996)

Nothing known (Rasmussen 1973; Thorson 1946; Wilson 1936b) (Yokouchi 1991)

(Kleinenberg 1886; Åkesson 1967b) (Beard 1884; Eeckhaut et al. 2003; Eeckhaut and Jangoux 1993; Jägersten 1939; Kato 1952)

Table 6.1 contd

Yes; mechanism undescribed; encounter predators? None to date; very late larvae may feed Not known Sometimes; oral brush in some, others mechanism not known

Not known None to date

Nothing known (Blake 1975b; Cazaux 1967) (Simpson 1962) Yes; mechanism undescribed (Blake 1975b; Cazaux 1972) Possible; some lecithotrophic (Blake 1975b; Haaland and Schram 1982; Haaland and Schram 1983; Sometimes; mechanism undescribed Rasmussen 1956; Schram and Haaland 1984) Nothing known

Nothing known (Allen 1959; Blake 1975a; Hsieh and Simon 1987; Paxton 1986; Paxton et al. 1995) Nothing known (Drasche 1885) (Blake 1975b; Cazaux 1968; Kisseleva 1992)

Annelid Larval Morphology

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Phyllodocidae

Less inclusive Taxa

Development studies

Planktotrophy

(Blake 1975b; Lacalli 1986; Meyer 1938; Thorson 1946; Tzetlin 1998) Sometimes; mechanism undescribed. Some would appear to be encounter predators Pilargidae (Blake 1975b; Britayev 1981) Not known Aciculata Phyllodocida Pisionidae (Aiyar and Alikunhi 1940; Stecher 1968; Åkesson 1961) Yes; ‘slime net’ Polynoidae (Cazaux 1968; Holborow 1971; Phillips and Pernet 1996) Yes; oral brush (Pernet 2000) Pontodoridae Nothing known Sigalionidae (Cazaux 1968; Phillips and Pernet 1996) Yes; oral brush Sphaerodoridae (Christie 1984; Mileikovsky 1967) Not known Syllidae (Cazaux 1969, 1984; Dales 1951; Garwood 1991) (Pernet 1998) None to date Tomopteridae (Åkesson 1962) None to date (Jouin and Swedmark 1965) (Jouin 1971) None to date Aciculata incertae sedis Nerillidae Aberranta Nothing known Spinther Nothing known Acrocirridae Nothing known Canalipalpata Cirratulidae (incl. (Blake 1975c; Gibson 1977; Gibson and Clark 1976; Monticelli 1910; None to date Cirratuliformia Ctenodrilinae) Okuda 1946; Qian and Chia 1989; Sokolow 1911; Wilson 1936a) Fauveliopsidae Nothing known Flabelligeridae (Amor 1994; Spies 1977) None to date Sternaspis (Child 1900; M. Strathmann 1987; Vejdovsky 1882) None to date None to date Alvinellidae (Zal et al. 1995) None to date Ampharetidae (Clavier 1984; Grehan et al. 1991; Nyholm 1957; Okuda 1947; Zottoli 1999; Zottoli 1974) Canalipalpata Pectinariidae (Lagadeuc and Retière 1993; Pernet 2004; Rasmussen 1973; Yes; filtering ‘house’ Terebelliformia Watson 1928; Wilson 1936b)

Table 6.1 contd

#$ Reproductive Biology and Phylogeny of Annelida

Blake 1969; Daro and Polk 1973; Hannerz 1956; Nozais et al. 1997)

Spionidae (incl. Heterospio, Poecilochaetus, Trochochaeta and Uncispionio) Oweniidae Sabellariidae

Annelida Incertae sedis

Canalipalpata incertae sedis

(Hatschek 1881; Korn 1959; Newby 1940; Jägersten 1972; Miner et al. 1999; this study) (Bunke 1967)

(Jouin 1962; Jägersten 1952; Sasaki and Brown 1983)

Aeolosomatidae and Potamodrilus Nothing known Hrabeiella Parergodrilidae Nothing known Psammodrilidae (Swedmark 1955)

Echiura

Protodrilida

Serpulidae Siboglinidae Polygordiidae

(Emlet and Strathmann 1994; Wilson 1932b) (Bhaud and Fernandez-Alamo 2001; Dales 1952a; Pernet 2003; Smith and Chia 1985; R.R. Strathmann 1987; Wilson 1929) (Gambi et al. 2000; Giangrande et al. 2000; Rouse and Fitzhugh 1994; Rouse and Gambi 1998) (Charles et al. 2003; Kupriyanova et al. 2001) (Bakke 1990; Southward 1999, 2000; Young et al. 1996) (Hatschek 1878; Woltereck 1904)

(Wilson 1982; Johnson and Brink 1998)

Nothing known (Bhaud 1966; Cazaux 1965) (Werner 1953)

(Bhaud and Grémare 1988; Heimler 1981; McHugh 1993; Okuda 1946)

Magelonidae

Terebellidae (including Trichobranchinae) Apistobranchus Chaetopteridae

Canalipalpata Sabellida Sabellidae

Canalipalpata Spionida

Canalipalpata Terebelliformia

None to date

Sometimes: opposed bands, pharyngeal apparatus or oral arms Sometimes; with encounter predation or opposed bands None to date

Yes; probably opposed band method

Sometimes; opposed bands

Yes; opposed bands Yes; mechanisms with opposed bands according to Pernet (2003) None to date

Yes: mechanism involves mucous trails and particle capture with buccal ciliation Yes; possibly ciliary feeding then may be encounter predators once palps appear Some; not well understood; ciliary feeding based on abdominal cilia or mucus string

None to date

Annelid Larval Morphology

#%

#& Reproductive Biology and Phylogeny of Annelida Sphaerodoridae, Terebellidae (incl. Trichobranchidae), Tomopteridae (see Table 6.1). Most of these groups are well studied and feeding larval forms, if they occur, would presumably have turned up in detailed plankton surveys such as those of Thorson (1946). This list of families with no planktotrophic larvae given above disagrees with the findings of Wilson (1991) and Giangrande (1997). For instance, Wilson (1991) erroneously listed the Cirratulidae, Dorvilleidae, Eunicidae, Nereididae, Sabellidae and Syllidae as having planktotrophic members. Some of these mistakes are simple taxonomic errors e.g., the serpulid Pomatoceros triqueter has planktotrophic larvae and is listed under the Sabellidae, but most appear to be misinterpretations of planktonic existence for planktotrophy.

6.5.

METAMORPHOSIS

A developmental transition between the larva and juvenile form is often required and this is referred to as metamorphosis. Among annelids, this transformation can be major and rapid or basically non-existent. If there are significant larval structures that are not needed in the adult then these need to be transformed or lost. In most annelids this minimally means the ciliary bands such as the prototroch are lost or altered. For instance, in the opheliid Armandia brevis it has been shown that the cilia of the prototroch, metatroch and telotroch are shed at metamorphosis and the trochoblasts degenerate (Hermans 1978). It is those taxa that have specialised larval feeding structures that metamorphosis is most conspicuous. This is most dramatic in the larvae of Oweniidae and some Polygordiidae. These larvae have elaborate food gathering devices for use in the planktonic environment that are useless in later life. In Polygordiidae there are two basic larval forms, the endolarva and the exolarva (Woltereck 1902). An exolarva is shown in Fig. 8.6A and B and has the future juvenile body emerging from the episphere and hyposphere. A similar larva is seen in Polygordius neapolitanus and metamorphosis involves a gradual loss of the larval body (Hatschek 1878). In other Polygordius with endolarvae, the future juvenile body is enclosed in the larval body. Here metamorphosis involves rupturing the larval body (Woltereck 1902), with it basically being cast aside or eaten by the young juvenile. A similar situation is seen in the mitraria larva of Oweniidae. Fig. 6.6C, D show a premetamorphic mitraria with a large episphere and an array of larval chaetae. The juvenile is actually coiled up inside the larval body. In only a few seconds the larval body and chaetae can be cast off leaving the small juvenile worm ready for a benthic life (Fig. 6.6E). In most annelids metamorphosis, particularly those with benthic lecithotrophic development, is not as drastic as this. The ciliary bands may be simply resorbed and larval chaeta may be gradually be shed. In groups such as fabriciin sabellids and clitellates there is very little in the way of metamorphosis and as seen in Fig. 6.7A-C. The larvae in this case are

Annelid Larval Morphology

#'

Fig. 6.7 contd

$ Reproductive Biology and Phylogeny of Annelida brooded in the female tube and have no need for ciliary band for feeding or dispersal. The larvae develop directly from an embryo through to a juvenile and there is no particular phase that could be referred to as metamorphosis.

6.6 EVOLUTION OF ANNELID LARVAE 6.6.1

Introduction

At present our understanding of annelid phylogeny is poor (see Chapter 1) and so we cannot state with confidence the plesiomorphic form of larval development. Of major interest for those interested in the evolution of animals is whether planktotrophy or lecithotrophy is the plesiomorphic condition for Annelida. For instance, Nielsen (1995) regarded the ‘opposed-band trochophore’ as plesiomorphic for a large clade of Metazoa and similar views have been proposed by Davidson et al. (1995), Strathmann (1993), Wray (1995) and Peterson et al. (1997). While a robust well supported phylogeny of Annelida is still some way off, there have been attempts to assess the evolution of larvae among annelids (Rouse 1999, 2000a, 2000b, 2000c) and this work will be briefly summarised here. It must be noted though that the phylogenetic hypothesis presented will almost certainly be dramatically altered, and so perhaps the broad inferences on the evolution of larvae. Rouse (2000a) obtained results based on a morphological cladistic analysis that downstream larval-feeding appeared independently at least nine times among annelids: in Echiura, the clade comprised Oweniidae, Polygordiidae, Protodrilidae, Protodriloididae, and Saccocirridae (with subsequent loss in the Protodrilidae and Protodrilidae clade); the clade Arenicolidae, Capitellidae, Maldanidae, Opheliidae, Scalibregmatidae (with subsequent loss in the Arenicolidae, Maldanidae clade); the clade Sabellariidae, Sabellidae and Serpulidae (with subsequent loss in the Sabellidae); Spionida, Pectinariidae, Amphinomida, and twice in Phyllodocida excluding Syllidae (with losses in Nereididae, Pisionidae, Sphaerodoridae). The occurrence of downstream larval-feeding among polychaetes is shown in Figs. 6.8, 6.9, and 6.10 (for illustrated transformations, see Rouse 2000a). The variety of annelid larval forms is also illustrated on these figures. There appears to be several types of downstream feeding when this is considered in relation to the occurrence of ciliary bands and what is known Fig. 6.7 contd

Fig. 6.7. Annelid larvae that show little, if any, metamorphosis A. Embryos taken from the tube of a female fabriciin referred to as Augeneriella cf. dubia (Sabellidae) in Rouse (1990) and is Augeneriella alata of Hartmann-Schröder (1991), but actually belongs in ‘Genus A’ of Fitzhugh (1993). B. Larva a few days later shows clear segmentation and developing radioles C. After 9 days in the tube a young juvenile with a fully developed radiolar crown gut as well as anterior and posterior eyes is ready to crawl away. D. A cocoon with four embryos of Enchytraeus coronatus (Clitellata) early in development. E. A cocoon with two embryos of Enchytraeus coronatus (Clitellata) at a similar stage in development. F. Two already segmented worms one day before emerging from cocoon. G. Juvenile just before emerging from the cocoon with chaetae now emergent. A-C by G.W. Rouse; D-G courtesy of A. Bergter.

Annelid Larval Morphology

$

Fig. 6.8. Basal portion of one of six most-parsimonious trees (after SACW) from the A/Pw analysis of Rouse (2000a). Taxon names shown in bold are represented by a larval diagram. The arrow seen in this figure leads to the clade Palpata and is shown in Figs. 6.9 and 6.10. Taxon names that are underlined have members (at least some) that shown downstream feeding, except for Mollusca, where feeding larvae are derived within the Mollusca. Terminals marked with “?” have larvae that are unknown. Sources of the modified larval diagrams are as follows: Mollusca, Neomenia carinata (modified from Jägersten 1972, Fig. 37A); Sipuncula, Sipunculus vulgaris (modified from Gerould 1907, Fig. 50); Arenicolidae, Arenicola marina (modified from Newell 1948, Fig. 7); Maldanidae Clymenella torquata (modified from Lacalli 1980, Fig. 29); Capitellidae, Mediomastus fragilis (modified from Rasmussen 1956, Fig. 21); Opheliidae Ophelia bicornis (modified from Wilson 1948, Fig. 1F); Orbinidae, Leitoscoloplos pugettensis (modified from Blake 1980, Fig. 1B).

$

Reproductive Biology and Phylogeny of Annelida

Fig. 6.9. Portion of one of six most-parsimonious trees (after SACW) from the A/Pw analysis of Rouse (2000a) showing one of the two major clades of Palpata; Aciculata. Taxon names shown in bold are represented by a larval diagram. The sister group to Aciculata is Canalipalpata and is shown in Fig. 6.10. Taxon names that are underlined have members (at least some) that shown downstream feeding. Terminals Fig. 6.9 contd

Annelid Larval Morphology

$!

about larval feeding. The broad definition, assumption of homology of downstream feeding made in Rouse (2000a), and the fact that polymorphic terminals were scored as having downstream feeding, amount to a biases in favor of this mode being primitive. Close examination of the actual feeding methods actually suggests the initial homology hypothesis is incorrect. This ultimately is supported by the disparate occurrence of downstream larval feeding shown on Figs. 6.8, 6.9, and 6.10. Though little is known about the actual downstream feeding of certain larval forms, it is clear that there are several modes of feeding, and up to 10 separate types can be identified. These are summarised as follows.

6.6.2 Downstream Feeding with Meniscotroch (four kinds?) Several taxa in Phyllodocida with planktotrophic larvae lack a metatroch (and food groove). These are Nephtyidae, Phyllodocidae, Polynoidae and Sigalionidae. The feeding mechanism in the latter two taxa was elucidated by Phillips and Pernet (1996) and involves the prototroch, meniscotroch and the oral brush. This mechanism allows the capture of a much greater range of particle sizes (2-60 µm in diameter) than opposed-band feeding. Phyllodocidae and Nephtyidae lack an oral brush (though for nephtyids, see Wilson 1936b), but have been recorded as ingesting large particles (R.R. Strathmann 1987) and do possess a meniscotroch. Other polychaete families within Phyllodocida also have a meniscotroch and have planktotrophic larvae. This includes Chrysopetalidae, Glyceridae, Hesionidae, and possibly Paralacydonia (see Table 6.1), though the latter taxa seems to lack a meniscotroch. The feeding mechanisms of these taxa have yet to be described, but may involve the use of the prototroch and meniscotroch (where present) and this has been seen in Chrysopetalidae (B. Pernet pers. comm.).

6.6.3 Opposed-band Larval Feeding (six kinds?) Opposed-band larval feeding was scored as present in Rouse (1999) for Echiura, amphinomids, oweniids, polygordiids, serpulids and opheliids. The cladistic placement of these taxa (Figs. 6.8, 6.9, 6.10) suggests that this mode Fig. 6.9 contd

marked with “?” have larvae that are unknown. Sources of the modified larval diagrams are as follows: Polynoidae, Harmothoe longisetis (modified from Cazaux 1968, Fig. V.1); Sigalionidae, Sthenelais boa (modified from Cazaux 1968, Fig. XIV. 2); Pholoidae, Pholoe synopthalmica (modified from Cazaux 1968, Fig. XVI.2); Chrysopetalidae, Paleonotus belli (modified from Blake 1975b, Fig. 4); Glyceridae, Glycera convoluta (modified from Fuchs 1911, Fig.5); Paralacydonia paradoxa (modified from Bhaud 1967, Fig. 3A); Phyllodocidae, Anaitides williamsi (modified from Blake 1975b, Fig. 8A); Pisionidae, Pisione remota (modified from Aiyar and Alikunhi 1940, Fig. 8); Hesionidae, Ophiodromus pugettensis (modified from Blake 1975b, Fig. 13); Nephtyidae, Nephtys hombergi (modified from Fuchs 1911, Fig. 2); Nereididae, Platynereis bicanaliculata (modified from Blake 1975b, Fig. 21); Syllidae, Syllis variegata (modified from Cazaux 1984, Fig. 2); Amphinomidae, Eurythoe complanata (modified from Kudenov 1974, Fig. 35); Dorvilleidae, Dorvillea rudolphi (modified from Blake 1975b, Fig. 28); Lumbrineridae, Lumbrineris impatiens (modified from Cazaux 1972, Fig. VII.2); Eunicidae, Eunice valens (as E. kobiensis) (modified from Åkesson 1967a, Fig. 1); Onuphidae, Nothria elegans (modified from Blake 1975a, Fig. 1B).

$" Reproductive Biology and Phylogeny of Annelida

Fig. 6.10 Portion of one of six most-parsimonious trees (after SACW) from the A/Pw analysis of Rouse (2000a) showing one of the two major clades of Palpata; Canalipalpata. Taxon names shown in bold are represented by a larval diagram. The sister group to the Canalipalpata is Aciculata and is shown in Fig. 6.10 contd

Annelid Larval Morphology

$#

evolved independently in each taxon (with the exception of Oweniidae and Polygordiidae where it may be homologous). When the actual morphology of the structures providing this opposed-band feeding is studied it appears that this convergence may be reasonably explained. For instance the opposedband larval feeding in the oweniid Owenia fusiformis is unique in that it is based on simple cilia (Emlet and Strathmann 1994), whereas other opposedband systems appear to be based on compound cilia. In Amphinomidae, the diagram of particle flow by Jägersten (1972) suggests a ciliated food groove is present and that opposed-band larval feeding is occurring on the tentacles, something unique to this group. Hermans (1978) and Miner et al. (1999) described opposed-band larval feeding in Armandia brevis. While A. brevis does gather small particles in a classical way it also captures large particles with oral ciliature and it is not known yet which of the two mechanisms is the plesiomorphic form for opheliids. Given that other opheliids are lecithotrophic (e.g., Ophelia bicornis see Wilson 1948) and there are planktotrophic larvae that lack a metatroch, such as Armandia cirrosa (Guérin 1973), the opposed band feeding maybe secondary within Opheliidae. Further detailed study on other taxa with opposed band feeding may reveal other evidence for convergence in this feeding mode. Pernet (2003) states that opposed band larval feeding occurs in Sabellariidae, but the details of this have yet to be published. Strathmann (1987) mentions a metatroch is present in Sabellariidae, but this is based on unpublished observations and is not supported by other studies. A metatroch was not shown by Wilson (1929), Dales (1952a) or Smith and Chia (1985) in their detailed studies of sabellariid larvae. Wilson (1929) did refer to posterior extensions of the prototroch that overhang the lateral edges of the mouth as lip folds and it is conceivable that they act in a similar fashion to a metatroch by beating food back towards the mouth.

Fig. 6.10 contd

Fig. 6.9. Taxon names that are underlined have members (at least some) that shown downstream feeding. Terminals marked with “?” have larvae that are unknown. Sources of the modified larval diagrams are as follows: Flabelligeridae Flabelliderma commensalis (modified from Spies 1977, Fig. 8); Cirratulidae Cirriformia spirabrancha (modified from Blake 1975c, Fig. 2); Ampharetidae Melinna palmata (modified from Grehan et al. 1991, Fig. 1B); Pectinariidae Pectinaria koreni (modified from Sveshnikov 1978, Fig. 39); Trichobranchidae Terebellides stroemi (modified from Willemöes-Suhm 1871, Fig. 25); Terebellidae Amphitrite ornate (modified from Mead 1897, Fig. X); Spionidae Scolecolepis fuliginosa (modified from Day 1934, Fig. 4); Magelonidae Magelona alleni (modified from Wilson 1982, Fig. 1); Chaetopteridae, Chaetopterus variopedatus (modified from Cazaux 1965, Fig. 4); Sabellariidae, Sabellaria alveolata (modified from Wilson 1929, Fig. 1.8); Sabellidae, Chone duneri (modified from Yun and Kikuchi 1991, Fig. 1E); Serpulidae, Galeolaria caespitosa (modified from Andrews and Anderson 1962, Fig. 6); Siboglinidae, Siboglinum fiordicum (modified from Bakke 1974, Fig. 5); Polygordiidae, Polygordius sp. (modified from Hatschek 1878, Fig. 28); Protodrilus adhearens (modified from Jägersten 1952, Fig. 30); Saccocirridae Saccocirrus papillocerus (modified from Pierantoni 1906, Fig. 28); Oweniidae Owenia fusiformis (modified from Wilson 1932b, Fig. 29.1).

$$ Reproductive Biology and Phylogeny of Annelida

6.6.4 Other Downstream Feeding Mechanisms (four kinds?) Among other polychaetes, Capitellidae, Magelonidae and Spionidae may also have downstream-feeding larvae, though the actual mechanisms have yet to be described. The unusual feeding mechanism of Pectinariidae that involves a ‘filter house’ has only recently been elucidated (Pernet 2004). Capitellidae have been regarded as having opposed-band feeding (Nielsen 1998). Hansen (1993) suggested that the larvae of the capitellid Mediomastus fragilis feed with a prototroch and metatroch. While this species may be a downstream-feeder, the larvae of M. fragilis clearly lack a metatroch, as shown by Rasmussen (1956) (as H. filiformis) and no other studies of capitellid larvae have described a metatroch.Feeding by larval spionids and magelonids has been described briefly (Daro and Polk 1973; Wilson 1982). Daro and Polk (1973) state that lateral circular currents are produced by the larvae of Polydora ciliata larvae that are 3-8 segments long and that they ingest particles less than 20 µm in diameter. They suggest that ‘abdominal cilia’ are responsible for generating this feeding current. However, since a neurotroch is lacking in spionid larvae, the only abdominal bands that could generate this current are gastrotrochs (Wilson 1928) and it seems from the diagram by Daro and Polk (1973) that the prototroch could also be involved. How food particles are actually captured by young spionid larvae has yet to be explained. Wilson (1982) fed young magelonid larvae on various flagellates and diatoms. Presumably the young larvae fed with the aid of their prototroch, which Wilson (1982) described as being somewhat similar to that of sabellariids in being expanded ventrally over the mouth. With the development of their feeding palps the magelonid larvae became encounter predators on bivalve larvae (though see Johnson and Brink, (1998) for new views on encounter predation by polychaete larvae).

6.7 SUMMARY AND CONCLUSIONS There is clearly a wide diversity of larval development and larval feeding modes in annelids, and opposed-band feeding represents but one of these. It would appear that various larval-feeding modes have evolved independently from lecithotrophic condition. In each case it would appear that the prototroch, which has a primarily locomotory role, has become involved in larval-feeding in association with other ciliary bands such as the meniscotroch, metatroch and/or oral brush. It is possible that larval feeding has evolved separately as many as 10 times among polychaetes. A more robust phylogenetic hypothesis for Annelida is certainly needed. Also there remain many polychaete groups where knowledge of larval development is poorly known (Table 6.1).

6.8

ACKNOWLEDGEMENTS

Thanks to Barrie Jamieson for conceiving this volume. Many thanks to Annette Bergter for generously providing the pictures of developing

Annelid Larval Morphology



%$Enchytraeus coronatus. Thanks to Fredrik Pleijel for his comments on the chapter.

6.9 LITERATURE CITED Aiyar, R. G. and Alikunhi, K. H. 1940. On a new pisionid from the sandy beach, Madras. Records of the Indian Museum, Calcutta 42: 89-107. Åkesson, B. 1961. On the histological differentiation of the larvae of Pisione remota (Pisionidae, Polychaeta). Acta Zoologica 42: 177-225. Åkesson, B. 1962. The embryology of Tomopteris helgolandica (Polychaeta). Acta Zoologica 43: 135-199. Åkesson, B. 1967a. The embryology of the polychaete Eunice kobiensis. Acta Zoologica 48: 141-192. Åkesson, B. 1967b. On the nervous system of the Lopadorynchus larva (Polychaeta). Arkiv för Zoologi 20: 55-78. Åkesson, B. 1973a. Morphology and life history of Ophryotrocha maculata sp. n. (Polychaeta, Dorvilleidae). Zoologica Scripta 2: 141-144. Åkesson, B. 1973b. Reproduction and larval morphology of five Ophryotrocha species (Polychaeta, Dorvilleidae). Zoologica Scripta 2: 145-155. Allen, M. J. 1959. Embryological development of the polychaetous annelid, Diopatra cuprea (Bosc). Biological Bulletin. Marine Biological Laboratory, Woods Hole, Mass. 116: 339-361. Amor, A. 1994. Gametes, fertilization and development of Pherusa sp., an endolithic worm (Polychaeta, Flabelligeridae). Mémoires du Muséum Nationale d’Histoire Naturelle 162: 612. Anderson, D. T. 1959. The embryology of the polychaete Scoloplos armiger. Quarterly Journal of Microscopical Science 100: 89-166. Anderson, D. T. 1961. The development of the polychaete Haploscoloplos fragilis. Quarterly Journal of Microscopical Science, London 102: 257-272. Anderson, D. T. 1966. The comparative early embryology of the Oligochaeta, Hirudinea and Onychophora. Proceedings of the Linnean Society of New South Wales 91: 10-43. Anderson, D. T. 1971. Embryology. Pp. 73-103. In R. O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World (with contributions by D.G. Cook, D.V. Anderson, J. van der Land), Oliver and Boyd, Edinburgh. Anderson, D. T. 1973. Embryology and Phylogeny in Annelids and Arthropods. Pergamon Press, Oxford, 495 pp. Andrews, J. C. and Anderson, D. T. 1962. The development and settling of the polychaete Galeolaria caespitosa Lamarck (Fam. Serpulidae). Proceedings of the Linnean Society of New South Wales 87: 185-188. Bailey-Brock, J. H. 1984. Spawning and development of Arenicola brasiliensis (Nonato) in Hawaii (Polychaeta; Arenicolidae). Pp. 439-449. In P. A. Hutchings (ed.), Proceedings of the First International Polychaete Conference, Sydney, Australia, 1983, The Linnean Society of New South Wales, Sydney. Bakke, T. 1974. Settling of the larvae of Siboglinum fiordicum Webb (Pogonophora) in the laboratory. Sarsia 56: 57-70. Bakke, T. 1990. Pogonophora. Pp. 37-48. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates. Volume IV, Part B Fertilization, Development, and Parental Care. John Wiley and Sons, Chichester. Beard, J. 1884. On the life history and development of the genus Myzostoma (F.S. Leuckart). Mittheilungen aus der Zoologischen Station zu Neapel 5: 544-580.

$& Reproductive Biology and Phylogeny of Annelida Bergter, A., Beck, L. A. and Paululat, A. 2004. Embryonic development of the oligochaete Enchytraeus coronatus: An SEM and histological study of embryogenesis from one-cell stage to hatching. Journal of Morphology 261: 26-42. Bhaud, M. 1966. Étude du devéloppement et l’écologie de quelques larves de Chaetopteridae (Annélides polychètes). Vie Milieu 17: 1087-1120. Bhaud, M. 1967. Étude du développement de quelques larves d’annélides polychètes à Banyuls-sur-Mer. Vie et Milieu 18: 531-558. Bhaud, M. 1983. Premières observations de la larve planctonique récoltee en Haute Mer d’un représentant des Paraonidae (Annélide Polychète). Vie et milieu 33: 41-48. Bhaud, M. and Cazaux, C. 1982. Les larves de polychètes des côtes de France. Oceanis 8: 57-160. Bhaud, M. and Cazaux, C. 1987. Description and identification of polychaete larvae; their implications in current biological problems. Oceanis 13: 596-753. Bhaud, M. and Grémare, A. 1988. Larval development of the terebellid polychaete Eupolymnia nebulosa (Montagu) in the Mediterranean Sea. Zoologica Scripta 17: 347-356. Bhaud, M. R. and Fernandez-Alamo, M. A. 2001. First description of the larvae of Dianthyrsus (Sabellariidae, Polychaeta) from the Gulf of California and Bahia de Banderas, Mexico. Bulletin of Marine Science 68: 221-232. Blake, J. A. 1969. Reproduction and larval development of Polydora from nothern New England (Polychaeta: Spionidae). Ophelia 7: 1-63. Blake, J. A. 1975a. The larval development of Polychaeta from the northern California coast. II. Nothria elegans (Family Onuphidae). Ophelia 13: 43-61. Blake, J. A. 1975b. The larval development of Polychaeta from the northern California coast. III Eighteen species of Errantia. Ophelia 14: 23-84. Blake, J. A. 1975c. The larval development of Polychaeta from the northern California. I. Cirriformia spirabrancha (family Cirratulidae). Transactions of the American Microscopical Society 94: 179-188. Blake, J. A. 1980. The larval development of Polychaeta from the northern California coast. IV. Leitoscoloplos pugettensis and Scoloplos acmeceps (family Orbiniidae). Ophelia 19: 1-18. Bookhout, C. G. and Horn, E. C. 1949. The development of Axiothella mucosa (Andrews). Journal of Morphology 84: 145-183. Britayev, T. A. 1981. Larvae of Cabira cf. bohajensis (Polychaeta, Pilargidae) from Vostok bay in the Sea of Japan. Doklady Akademii Nauk SSSR 260: 1278-1280. Bunke, D. 1967. Zur Morphologie und Systematik der Aeolosomatidae Beddard 1895 und Potamodrilidae nov. fam. (Oligochaeta). Zoologische Jahrbücher, Abteilung für Anatomie und Ontogenie der Tiere 94: 187-368. Cazaux, C. 1965. Développement larvaire de Chaetopterus variopedatus (Renier). Actes de la Société Linnéenne de Bordeaux 102: 1-16. Cazaux, C. 1967. Développement larvaire de Glycera convoluta Keferstein. Vie et Milieu 18: 559-571. Cazaux, C. 1968. Étude morphologique du developpement larvaire d’annelides polychetes (Bassin d’Arcachon). I. Aphroditidae, Chrysopetalidae. Archives de Zoologie Expérimentale et Générale 109: 477-543. Cazaux, C. 1969. Étude morphologique du developpement larvaire d’annelides polychetes (Bassin d’Arcachon). II. Phyllodocidae, Syllidae, Nereidae. Archives de Zoologie Expérimentale et Générale 110: 145-202. Cazaux, C. 1972. Développement larvaire d’annélides polychètes (Bassin d’Arcachon). Archives de Zoologie Expérimentale et Générale 113: 71-108.

Annelid Larval Morphology

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Cazaux, C. 1984. Reproduction et développement larvaire du Syllidae Syllis variegata Grube, 1860 (Annelide, Polychete). Cahiers de Biologie Marine 25: 123139. Charles, F., Jordana, E., Amouroux, J. M., Gremare, A., Desmalades, M. and Zudaire, L. 2003. Reproduction, recruitment and larval metamorphosis in the serpulid polychaete Ditrupa arietina (O. F. Müller). Estuarine Coastal and Shelf Science 57: 435-443. Child, C. M. 1900. The early development of Arenicola and Sternaspis. Wilhelm Roux’ Archiv für Entwicklungsmechanik der Organismen 9: 587-722. Christie, G. 1984. The reproductive biology of a Northumberland population of Sphaerodorum gracilis (Rathke, 1843) (Polychaeta: Sphaerodoridae). Sarsia 69: 117121. Clavier, J. 1984. Description du cycle biologique d’Ampharete acutifrons (Grube, 1860) (Annélide Polychète). Compte rendu des séances de l’Académie des sciences III Sciences de la vie 299: 59-62. Dales, R. P. 1950. The reproduction and larval development of Nereis diversicolor (O. F. Müller). Journal of the Marine Biological Association of the United Kingdom 29: 321-360. Dales, R. P. 1951. Observations of the structure and life history of Autolytus prolifer (O. F. Müller). Journal of the Marine Biological Association of the United Kingdom 30: 119-128. Dales, R. P. 1952a. The development and structure of the anterior region of the body in the Sabellariidae, with special reference to Phragmatopoma californica. Quarterly Journal of Microscopical Science 93: 435-452. Dales, R. P. 1952b. The larval development and ecology of Thoracophelia mucronata (Treadwell). Biological Bulletin. Marine Biological Laboratory, Woods Hole 102: 232-242. Damen, P. and Dictus, W. J. A. G. 1994. Cell lineage of the prototroch of Patella vulgata (Gastropoda, Mollusca). Developmental Biology 162: 364-383. Daro, M. H. and Polk, P. 1973. The autecology of Polydora ciliata along the Belgian coast. Netherlands Journal of Sea Research 6: 130-140. Davidson, E. H., Peterson, K. J. and Cameron, R. A. 1995. Origin of bilaterian body plans: Evolution of developmental regulatory mechanisms. Science 270: 13191325. Dawydoff, C. 1959. Ontogenèse des Annélides. Pp. 594-686. In P.-P. Grassé (ed.), Traité de Zoologie. Anatomie, Systématique, Biologie, vol. 5, Masson et Cie, Paris. Day, J. H. 1934. Development of Scolecolepis fuliginosa (Clpd.). Journal of the Marine Biological Association of the United Kingdom 19: 633-654. Drasche, R. von. 1885. Beiträge zur Entwicklung der Polychaeten. 2. Entwicklung von Sabellaria spinulosa, Hermione hystrix und einer Phyllodocidae. Verlag von Carl Gerold’s Sohn, Wien, 23 pp. Eckelbarger, K. J. and Grassle, J. P. 1987. Interspecific variation in genital spine, sperm and larval mophology in six sibling species of Capitella. Bulletin of the Biological Society of Washington 7: 62-76. Eeckhaut, I., Fievez, L. and Muller, M. C. 2003. Larval development of Myzostoma cirriferum (Myzostomida). Journal of Morphology 258: 269-283. Eeckhaut, I. and Jangoux, M. 1993. Life cycle and mode of infestation of Myzostoma cirriferum (Annelida), a symbiotic myzostomid of the comatulid crinoid Antedon bifida (Echinodermata). Diseases of Aquatic Organisms 15: 207-217. Eisig, H. 1899. Zur Entwicklungsgeschichte der Capitelliden. Mittheilungen aus der Zoologischen Station zu Neapel 13: 1-292.

% Reproductive Biology and Phylogeny of Annelida Emlet, R. B. and Strathmann, R. R. 1994. Functional consequences of simple cilia in the mitraria of oweniids (an anomalous larvae of an anomalous polychaete) and comparisons with other larvae. Pp. 143-157. In W. H. Wilson, S. A. Stricker and G. L. Shin (eds), Reproduction and Development of Marine Invertebrates., Johns Hopkins University Press, Baltimore. Farke, H. and Berghuis, E. M. 1979. Spawning, larval development and migration behaviour of Arenicola marina in the laboratory. Netherlands Journal of Sea Research 13: 512-528. Fewkes, J. W. 1883. On the development of certain worm larvae. Bulletin of the Museum of Comparative Zoology at Harvard College 11: 167-208. Fitzhugh, K. 1993. Novafabricia brunnea (Hartman, 1969), new combination, with an update on relationships among Fabriciinae taxa (Polychaeta: Sabellidae). Contributions in Science, Natural History Museum of Los Angeles County 438: 1-12. Fuchs, H. M. 1911. Note on the early larvae of Nephthys and Glycera. Journal of the Marine Biological Association of the United Kingdom 9: 164-170. Gambi, M. C., Giangrande, A. and Patti, F. P. 2000. Comparative observations on reproductive biology of four species of Perkinsiana (Polychaeta: Sabellidae: Sabellinae). Bulletin of Marine Science 67: 299-309. Garwood, P. R. 1991. Reproduction and the classification of the family Syllidae (Polychaeta). Ophelia Supplement 5: 81-87. Gerould, J. H. 1907. The development of Phasocolosoma (Studies on the embryology of the Sipunculidae, II). Zoologische Jahrbücher, Abteilung für Anatomie und Ontogenie der Tiere 23: 77-162. Giangrande, A. 1991. Reproduction, larval development and post-larval growth of Naineris laevigata (Polychaeta, Orbiniidae) in the Mediterranean Sea. Marine Biology 111: 129-137. Giangrande, A. 1997. Polychaete reproduction patterns, life-cycles and life-histories: an overview. Oceanography and Marine Biology, An Annual Review 35: 323386. Giangrande, A., Licciano, M., Pagliara, P. and Gambi, M. C. 2000. Gametogenesis and larval development in Sabella spallanzanii (Polychaeta: Sabellidae) from the Mediterranean Sea. Marine Biology 136: 847-861. Gibson, P. H. 1977. Reproduction in the cirratulid polychaetes Dodecaceria concharum and D. pulchra. Journal of Zoology 182: 89-102. Gibson, P. H. and Clark, R. B. 1976. Reproduction of Dodecaceria caulleryi (Polychaeta: Cirratulidae). Journal of the Marine Biological Association of the United Kingdom 56: 649-674. Giere, O. W. and Riser, N. W. 1981. Questidae — Polychaetes with oligochaetoid morphology and development. Zoologica Scripta 10: 95-103. Gravely, F. H. 1909. Studies on polychaete larvae. Quarterly Journal of Microscopical Science 53: 597-627. Grehan, A., Retière, C. and Keegan, B. 1991. Larval development in the ampharetid Melinna palmata Grube (Polychaeta). Ophelia Supplement 5: 321-332. Guberlet, J. E. 1933. Observations on the spawning and development of some Pacific annelids. Proceedings of the Fifth Pacific Science Congress 5: 4213-4220. Guérin, J.-P. 1971. Modalites d’elevage et description des stades larvaires de Polyophthalmus pictus Dujardin (Annelide Polychete). Vie Milieu 22: 143-152. Guérin, J.-P. 1973. Le developpement larvaire d’Armandia cirrosa Filippi (Annelide Polychete). Tethys 4: 969-974. Haaland, B. and Schram, T. A. 1982. Larval development and metamorphosis of Gyptis rosea (Malm) (Hesionidae, Polychaeta). Sarsia 67: 107-118.

Annelid Larval Morphology

%

Haaland, B. and Schram, T. A. 1983. Larval development and metamorphosis of Ophiodromus flexuosus (Delle Chiaje) (Hesionidae, Polychaeta). Sarsia 68: 8596. Hannerz, L. 1956. Larval development of the polychaete families Spionidae Sars, Disomidae Mesnil, and Poecilochaetidae N. Fam. in the Gullmar Fjord (Sweden). Zoologiska Bidrag från Uppsala 31: 1-204. Hansen, B. 1993. Aspects of feeding, growth and stage development by trochophora larvae of the boreal polychaete Mediomastus fragile (Rasmussen) (Capitellidae). Journal of Experimental Marine Biology and Ecology 166: 273-288. Hartmann-Schröder, G. 1991. Teil 16. Die Polychaeten der subtropisch-tropische bis tropische Ostküste zwischen Maclean (New South Wales) und Gladstone (Queensland) sowie von Heron Island (Grosses Barriere-Riff). Mitteilungen aus dem Hamburgischen Zoologischen Museum und Institut 88: 17-71. Haswell, W. A. 1916. On the embryology of Stratiodrilus (Histriobdellidae). Quarterly Journal of Microscopical Science 61: 301-312. Haszprunar, G., Salvini-Plawen, L. von and Rieger, R. M. 1995. Larval planktotrophy— A primitive trait in the Bilateria? Acta Zoologica 76: 141-154. Hatschek, B. 1878. Studien über die Entwicklungsgeschichte der Anneliden. Ein Beitrag zur Morphologie der Bilaterien. Arbeiten aus dem Zoologischen Institute der Universität Wien und der Zoologischen Station in Triest 1: 277-404. Hatschek, B. 1881. Über Entwicklung von Echiurus und die systematische Stellung der Echiuridae (Gephyrei chaetiferi). Arbeiten aus dem Zoologischen Institute der Universität Wien und der Zoologischen Station in Triest 3: 1-34. Heffernan, P. and Keegan, B. F. 1988a. The larval development of Pholoe minuta (Polychaeta: Sigalionidae) in Galway Bay, Ireland. Journal of the Marine Biological Association of the United Kingdom 68: 339-350. Heffernan, P. and Keegan, B. F. 1988b. Quantitative and ultrastructural studies on the reproductive biology of the polychaete Pholoe minuta in Galway Bay. Marine Biology 99: 203-214. Heimler, W. 1981. Untersuchungen zur Larvalentwicklung von Lanice conchilega (Pallas) 1766 (Polychaeta, Terebellomorpha) Teil II: Bau und Ultrastruktur der Trochophora-larve. Zoologische Jahrbücher (Anatomie und Ontogenie der Tiere) 106: 12-45. Hermans, C. O. 1978. Metamorphosis in the opheliid polychaete Armandia brevis. Pp. 113-126. In F.-S. Chia and M. E. Rice (eds), Settlement and Metamorphosis of Marine Invertebrate Larvae, Elsevier, New York. Hickman, C. S. 1999. Larvae in invertebrate development and evolution. Pp. 21–59. In B.K. Hall and M. H. Wake (eds), The Origin and Evolution of Larval Forms, Academic Press, San Diego, CA. Holborow, P. L. 1971. The fine structure of the trochophore of Harmothoë imbricata. Pp. 237-246. In D. J. Crisp (ed.), Fourth European Marine Biology Symposium, Cambridge University Press, Cambridge. Hsieh, H.-L. and Simon, J. L. 1987. Larval development of Kinbergonuphis simoni, with a summary of development patterns in the family Onuphidae (Polychaeta). Bulletin of the Biological Society of Washington 7: 194-210. Häcker, V. 1896. Pelagische Polychäten-Larven. Zur Kenntnis der Neapler Frühjahrs-Auftriebs. Zeitschrift für Wissenschaftliche Zoologie 62: 74-168. Häcker, V. 1898. Pelagische Polychätenlarven. 2. Zur Biologie der atlantischen Hochseeformen. Biologische Zentralblatt 18: 39-54. Irvine, S. Q., Chaga, O. and Martindale, M. Q. 1999. Larval ontogenetic stages of Chaetopterus: Developmental heterochrony in the evolution of chaetopterid polychaetes. 197: 319-331.

%

Reproductive Biology and Phylogeny of Annelida

Ivanova-Kazas, O. M. 1985a. The evolution of the polychaetous larvae. Explorations of the Fauna of the Seas 34: 40-45. Ivanova-Kazas, O. M. 1985b. The origin and phylogenetic significance of the trochophoran larvae. I The larvae of coelomate worms and molluscs. Zoologicheskii Zhurnal 64: 485-497. Ivanova-Kazas, O. M. 1985c. The origin and phylogenetic significance of the trochophoran larvae. II. Evolutionary significance of the larvae of coelomate worms and molluscs. Zoologicheskii Zhurnal 64: 650-660. Johnson, K. B. and Brink, L. A. 1998. Predation on bivalve veligers by polychaete larvae. Biological Bulletin 194: 297-303. Jouin, C. 1962. Le développement larvaire de Protodrilus chaetifer Remane (Archiannélides). Comptes Rendus de l’Académie des Science, Paris 255: 3065-3067. Jouin, C. 1971. Status of the knowledge of the systematics and ecology of Archiannelida. Smithsonian Contributions to Zoology 76: 47-56. Jouin, C. and Swedmark, B. 1965. Paranerilla limicola n.g., n.sp., Archiannélide Nerillidae du benthos vaseux marin. Cahiers de Biologie Marine 6: 201-218. Jägersten, G. 1939. Zur Kenntnis der Larvenentwicklung bei Myzostomum. Ark. Zool. 31A: 1-21. Jägersten, G. 1952. Studies on the morphology, larval development and biology of Protodrilus. Zoologiska Bidrag från Uppsala 29: 426-512. Jägersten, G. 1972. Evolution of the Metazoan Life Cycle., Academic Press, New York, 282 pp. Kato, K. 1952. On the development of Myzostome. Science Reports of the Saitama University Series B (Biology and Earth Sciences) 1: 1-16. Kisseleva, M. I. 1992. New genus and species of a polychaete of the Chrysopetalidae from the Black Sea. Zoologicheskii Zhurnal 72: 128 - 132. Kleinenberg, N. 1886. Die Entstehung des Annelides aus der Larve von Lopadorhynchus. Zeitschrift für Wissenschaftliche Zoologie 44: 1-227. Korn, H. 1959. Ergänzende Beobachtungen zur Struktur der Larve von Echiurus abyssalis Skor. Zeitschrift für Wissenschaftliche Zoologie 164: 199-237. Kudenov, J. D. 1974. The Reproductive Biology of Eurythoe Complanata (Pallas, 1766), (Polychaeta: Amphinomidae). University of Arizona, Ph.D. Thesis, 124 pp. Kudenov, J. D. 1977. Brooding behavior and protrandry in Hipponoe gaudichaudi (Polychaeta: Amphinomidae). Bulletin of the Southern California Academy of Sciences 76: 85-90. Kupriyanova, E. K. 2003. Live history evolution in Serpulimorph polychaetes: a phylogenetic analysis. Hydrobiologia 496: 105-114. Kupriyanova, E. K., Nishi, E., ten Hove, H. A. and Rzhavsky, A. V. 2001. A review of life history in serpulimorph polychaetes: ecological and evolutionary perspectives. Oceanography and Marine Biology: An Annual Review 39: 1-101. Kutschera, U. and Wirtz, P. 1986. Reproductive behaviour and parental care of Helobdella striata (Hirudinea, Glossiphoniidae): a leech that feeds its young. Ecology 72: 132-142. Lacalli, T. C. 1980. A guide to the marine flora and fauna of the Bay of Fundy: polychaete larvae from Passamaquoddy Bay. Canadian Technical Report of Fisheries and Aquatic Sciences 940: 1-27. Lacalli, T. C. 1986. Prototroch structure and innervation in the trochophore larva of Phyllodoce (Polychaeta). Canadian Journal of Zoology 64: 176-184. Lagadeuc, Y. and Retière, C. 1993. Critères d’identification rapide des stades de développement des larves de Pectinaria koreni (Malmgren) (Annélide; Polychète) de la Baie de Seine (Manche). Vie et Milieu 43: 217-224.

Annelid Larval Morphology

%!

Laubier, L. 1975. Adaptations morphologiques et biologiques chez un Aphroditien interstitiel: Pholoe swedmarki sp. n. Cahiers de Biologie Marine 16: 671-683. Marsden, J. R. 1960. Polychaetous annelids from the shallow waters around Barbados and other islands of the West Indies, with notes on their larval forms. Canadian Journal of Zoology 38: 989-1020. Mazurkiewicz, M. 1975. Larval development and habits of Laeonereis culveri (Webster) (Polychaeta: Nereidae). Biological Bulletin. Marine Biological Laboratory, Woods Hole 149: 186-204. McHugh, D. 1993. A comparative study of reproduction and development in the polychaete family Terebellidae. Biological Bulletin. Marine Biological Laboratory, Woods Hole, Mass. 185: 153-168. Mead, A. D. 1897. The early development of marine annelids. Journal of Morphology 13: 227-326. Meyer, A. 1938. Der Rogen (Spawn) und die Entwicklung der Trocophora von Eulalia viridis (Phyllodocidae). Biologia Generalis 14: 334-389. Mileikovsky, S. A. 1960. Appurtenance of a polychaete larva of the rostraria type from plankton of the Norwegian and Barents Seas of the species Euphrosyne borealis Oersted 1843 and of all larvae of this type to the families Euphrosynidae and Amphinomidae (Polychaeta, Errantia, Amphinomimorpha). (in Russian). Doklady Akademii Nauk SSSR 134: 731-734. Mileikovsky, S. A. 1961. Assignment of two Rostraria-type polychaete larvae from the plankton of the northwest Atlantic to species Amphinome passasi Quatrefages 1865 and Chloea atlantica McIntosh 1885 (Polychaeta, Errantia, Amphinomorpha). Doklady Akademii Nauk SSSR 141: 1109-1112. Mileikovsky, S. A. 1967. Larval development of polychaetes of the family Sphaerodoridae and some considerations of its systematics. Doklady Akademii Nauk SSSR 177: 851-854. Miner, B. G., Sanford, E., Strathmann, R. R., Pernet, B. and Emlet, R. E. 1999. Functional and evolutionary implications of opposed bands, big mouths, and extensive oral ciliation in larval opheliids and echiurids (Annelida). Biological Bulletin 197: 14-25. Monticelli, F. S. 1910. Raphidrilus nemasoma Montic. nuovo Ctenodrilide del Golfo di Napoli (Revisione diCtenodrilidi). Archivio Zoologico Unione Zoologica Italiana Napoli 4: 401-436. Needham, A. E. 1990. Annelida-Clitellata. Pp. 1-36. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates, Volume IV, Part B, Fertilization, development, and parental care. Wiley, Chichester. Newby, W. W. 1940. The embryology of the echiuroid worm Urechis caupo. Memoirs of the American Philosophical Society 16: 1-219. Newell, G. E. 1948. A contribution to the knowledge of the life history of Arenicola marina. Journal of the Marine Biological Association of the United Kingdom 27: 554-580. Newell, G. E. 1949. The later larval life of Arenicola marina. Journal of the Marine Biological Association of the United Kingdom 28: 635-639. Newell, G. E. 1951. The life history of Clymenella torquata (Leidy) (Polychaeta). Proceedings of the Zoological Society of London 121: 561-586. Nielsen, C. 1995. Animal evolution. Oxford University Press, Oxford. Nielsen, C. 1998. Origin and evolution of animal life cycles. Biological Reviews of the Cambridge Philosophical Society 73: 125-155. Nozais, C., Duchene, J. C. and Bhaud, M. 1997. Control of position in the water column by the larvae of Poecilochaetus serpens (Polychaeta) — the importance of mucus secretion. Journal of Experimental Marine Biology and Ecology 210: 91-106.

%" Reproductive Biology and Phylogeny of Annelida Nyholm, K.-G. 1957. Contributions to the life-history of the ampharetid, Melinna cristata. Zoologiska Bidrag från Uppsala 29: 79-91. Okuda, S. 1946. Studies on the development of Annelida Polychaeta I. Journal of the Faculty of Science, Hokkaido Imperial University, Ser. VI, Zoology 9: 115-219. Okuda, S. 1947. On an ampharetid worm, Schistocomus sovjecticus Annenkova, with some notes on its larval development. Journal of the Faculty of Science, Hokkaido University Series 6, Zoology 9: 321-329. Paxton, H. 1986. Generic revision and relationships of the family Onuphidae (Annelida: Polychaeta). Records of the Australian Museum 38: 1-74. Paxton, H., Fadlaoui, S. and Lechapt, J.-P. 1995. Diopatra marocensis, a new brooding species of Onuphidae (Annelida: Polychaeta). Journal of the Marine Biological Association of the United Kingdom 75: 949-955. Pernet, B. 1998. Benthic egg masses and larval development of Amblyosyllis speciosa (Polychaeta: Syllidae). Journal of the Marine Biological Association of the United Kingdom 78: 1369-1372. Pernet, B. 2000. Reproduction and development of three symbiotic scaleworms (Polychaeta: Polynoidae). Invertebrate Biology 119: 45-57. Pernet, B. 2003. Persistent ancestral feeding structures in nonfeeding annelid larvae. Biological Bulletin 205: 295-307. Pernet, B. 2004. The cryptic filtering house of an invertebrate larva. Science 306: 17571757. Pernet, B., P. Y. Qian, G.W. Rouse, C. M. Young and Eckelbarger, K. J. 2001. Phylum Annelida. Chapter 12. Pp. 209-236. In C. M. Young, M. Sewell and M. E. Rice (eds), Atlas of Marine Invertebrate Larvae. Academic Press, London. Peterson, K. J., Cameron, R. A. and Davidson, E. H. 1997. Set-aside cells in maximal indirect development: Evolutionary and developmental significance. Bioessays 19: 623-631. Phillips, N. E. and Pernet, B. 1996. Capture of large particles by suspension-feeding scaleworm larvae (Polychaeta, Polynoidae). Biological Bulletin 191: 199-208. Pierantoni, U. 1906. Osservazioni sullo sviluppo embrionale e larval del Saccocirrus papillocercus Bobr. Mittheilungen aus der Zoologischen Station zu Neapel 18: 4672. Qian, P.-Y. and Chia, F.-S. 1989. Sexual reproduction and larval development of Rhaphidrilus nemasoma Monticcelli, 1910 (Polychaeta: Ctenodrilidae). Canadian Journal of Zoology 67: 2345-2351. Rasmussen, E. 1956. Faunistic and biological notes on marine invertebrates III. Biologiske Meddelelser det Kongelige Dansk Videnskabernes Selskab. 23: 1-84. Rasmussen, E. 1973. Systematics and ecology of the Isefjord marine fauna (Denmark). Ophelia 11: 1-507. Reish, D. J. 1957. The life history of the polychaetous annelid Neanthes caudata (delle Chiaje), including a summary of development in the family Nereidae. Pacific Science 11: 216-228. Richards, T. L. 1967. Reproduction and development of the polychaete Stauronereis rudolphi, including a summary of development in the superfamily Eunicea. Marine Biology 1: 124-133. Rouse, G. W. 1990. New species of Oriopsis (Sabellidae: Polychaeta) and a new record for Augeneriella cf. dubia Hartmann-Schröder 1965 (Sabellidae: Polychaeta) from eastern Australia. Records of the Australian Museum 42: 221-235. Rouse, G. W. 1992. Oogenesis and larval development in Micromaldane spp. (Polychaeta: Capitellida: Maldanidae). Invertebrate Reproduction and Development 21: 215-230.

Annelid Larval Morphology

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Rouse, G. W. 1999. Trochophore concepts: ciliary bands and the evolution of larvae in spiralian Metazoa. Biological Journal of the Linnean Society 66: 411-464. Rouse, G. W. 2000a. Bias? What bias? Gain and loss of downstream larval-feeding in animals. Zoologica Scripta 29: 213-236. Rouse, G. W. 2000b. The epitome of hand waving? Larval feeding and hypotheses of metazoan phylogeny. Evolution and Development 2: 222-233. Rouse, G. W. 2000c. Polychaetes have evolved feeding larvae several times. Bulletin of Marine Science 67: 391-409. Rouse, G. W. and Fitzhugh, K. 1994. Broadcasting fables: Is external fertilization really primitive? Sex, size and larvae in sabellid polychaetes. Zoologica Scripta 23: 271-312. Rouse, G. W. and Gambi, M. C. 1998. Evolution of reproductive features and larval development in the genus Amphiglena Claparède (Polychaeta: Sabellidae). Marine Biology 131: 743-753. Sasaki, S. and Brown, R. 1983. Larval development of Saccocirrus uchidai from Hokkaido, Japan and Saccocirrus krusadensis from New South Wales, Australia (Archiannelida, Saccocirridae). Annotationes Zoologicae Japonenses 56: 299-314. Sato, M., Tsuchiya, M. and Nishihira, M. 1982. Ecological aspects of the development of the polychaete, Lumbrineris latreilli (Audouin et Milne-Edwards): significance of direct development and non-simultaneous emergence of the young from the jelly mass. Bulletin of the Marine Biological Station Asamushi, Tôhoku University 17: 71-85. Schram, T. A. and Haaland, B. 1984. Larval development and metamorphosis of Nereimyra punctata (O.F. Müller) (Hesionidae, Polychaeta). Sarsia 69: 169181. Schroeder, P. C. and Hermans, C. O. 1975. Annelida: Polychaeta. Pp. 1-213. In A. C. Giese and J. S. Pearse (eds), Reproduction of Marine Invertebrates. III. Annelids and Echiurans., vol. 3, Academic Press, New York. Shearer, C. 1910. On the anatomy of Histriobdella homari. Quarterly Journal of Microscopical Science 55: 287-359. Simpson, M. 1962. Gametogenesis and early development of the polychaete Glycera dibranchiata. Biological Bulletin. Marine Biological Laboratory, Woods Hole 123: 412-423. Smith, P. R. and Chia, F. S. 1985. Larval development and metamorphosis of Sabellaria cementarium Moore, 1906 (Polychaeta: Sabellariidae). Canadian Journal of Zoology 63: 1037-1049. Sokolow, I. 1911. Über eine neue Ctenodrilusart und ihre Vermehrung. Zeitschrift für wissenschaftliche Zoologie 97: 547-603. Southward, E. C. 1988. Development of the gut and segmentation of newly settled stages of Ridgeia (Vestimentifera): implications for relationship between Vestimentifera and Pogonophora. Journal of the Marine Biological Association of the United Kingdom 68: 465-487. Southward, E. C. 1999. Development of Perviata and Vestimentifera (Pogonophora). Hydrobiologia 402. Southward, E. C. 2000. Pogonophora. Pp. 331-351. In P. Beesely, G. J. B. Ross and C. J. Glasby (eds), Polychaeta and Allies: The Southern Synthesis. Fauna of Australia Volume 4A. Polychaeta, Myzostomida, Pogonophora, Echiura, Sipuncula. CSIRO Publishing, Melbourne. Spies, R. B. 1977. Reproduction and larval development of Flabelliderma commensalis (Moore). Pp. 323-345. In D. J. Reish and K. Fauchald (eds), Essays on Polychaetous Annelids in Memory of Dr. Olga Hartman, The Allan Hancock Foundation, University of Southern California, Los Angeles.

%$ Reproductive Biology and Phylogeny of Annelida Stecher, H. J. 1968. Zur Organisation und Fortplanzung von Pisione remota (Southern) (Polychaeta, Pisionidae). Zeitschrift für Morphologie und Ökologie der Tiere 61: 347-410. Strathmann, M. 1987. Reproduction and Development of Marine Invertebrates of the Northern Pacific Coast. Data and Methods for the Study of Eggs, Embryos, and Larvae. University of Washington Press, Seattle, 670 pp. Strathmann, R. R. 1978. The evolution and loss of feeding larval stages of marine invertebrates. Evolution 32: 894-906. Strathmann, R. R. 1987. Larval feeding. Pp. 465-550. In A. C. Giese, J. S. Pearse and V. B. Pearse (eds), Reproduction of Marine Invertebrates. Vol. IX. General Aspects: Seeking Unity in Diversity., Blackwell Scientific Publications and The Boxwood Press., California. Strathmann, R. R. 1993. Hypotheses on the origins of marine larvae. Annual Reviews in Ecology and Systematics 24: 89-117. Sveshnikov, V. A. 1978. Morphology of Larval Polychaetes. Akademiia Nauk SSSR, Moscow, 151 pp. Swedmark, B. 1955. Recherches sur la morphologie, le développement et la biologie de Psammodrilus balanoglossoides. Polychète Sédentaire de la microfaune des sables. Archives de zoologie expérimentale et générale 92: 141-220. Thorson, G. 1946. Reproduction and larval development of Danish marine bottom invertebrates with special reference to the planktonic larvae in the Sound (Øresund). Meddelelser fra Kommissionen for Danmarks Fiskeri-Og Havundersøgelser, Serie: Plankton 4: 1-523. Tzetlin, A. B. 1998. Giant pelagic larvae of Phyllodocidae (Polychaeta, Annelida). Journal of Morphology 238: 93-107. Vejdovsky, F. 1882. Untersuchungen über die Anatomie, Physiologie und Entwicklung von Sternaspis. Denkschriften der Akademie der Wissenschaften, Wien. 43: 33-90. Watson, A. T. 1928. Observations on the habits and life-history of Pectinaria (Lagis) koreni, Mgr. Proceedings and Transactions of the Liverpool Biological Society 42: 25-60. Watson, G. J. and Bentley, M. G. 1998. Oocyte maturation and post-fertilization development of Arenicola marina (L.) (Annelida: Polychaeta). Invertebrate Reproduction and Development 33: 35-46. Werner, B. 1953. Beobachtungen ueber den Nahrungserweib und die Metamorphose der Metatrochophora von Chaetopterus variopedatus Renier und Claparède (Polychaeta Sedentaria). Helgoländer wissenschaftliche Meeresuntersuchungen 4: 225-238. Willemöes-Suhm, R. 1871. Biologische Beobachtungen ueber niedere Meeresthiere. Zeitschrift für wissenschaftliche Zoologie 21: 380-396. Wilson, D. P. 1982. The larval development of three species of Magelona (Polychaeta) from localities near Plymouth. Journal of the Marine Biological Association of the United Kingdom 62: 385-401. Wilson, D. P. 1928. The larvae of Polydora ciliata Johnston and Polydora hoplura Claparède. Journal of the Marine Biological Association of the United Kingdom 15: 567-603. Wilson, D. P. 1929. The larvae of the British sabellarians. Journal of the Marine Biological Association of the United Kingdom 16: 221-268. Wilson, D. P. 1932a. The development of Nereis pelagica Linnaeus. J. Mar. Biol. Ass. U.K. 18: 203-217. Wilson, D. P. 1932b. On the Mitraria larva of Owenia fusiformis Delle Chiaje. Philosophical Transactions of the Royal Society of London. Series B 221: 231-334.

Annelid Larval Morphology

%%

Wilson, D. P. 1933. The larval stages of Notomastus latericeus Sars. Journal of the Marine Biological Association of the United Kingdom 18: 511-518. Wilson, D. P. 1936a. The development of Audouinia tentaculata (Montagu). Journal of the Marine Biological Association of the United Kingdom 20: 567-579. Wilson, D. P. 1936b. Notes on the early stages of two polychaete Nephtys hombergi Lamarck and Pectinaria koreni Malmgren. Journal of the Marine Biological Association of the United Kingdom 21: 305-310. Wilson, D. P. 1948. The larval development of Ophelia bicornis Savigny. Journal of the Marine Biological Association of the United Kingdom 27: 540-553. Wilson, D. P. 1982. The larval development of three species of Magelona (Polychaeta) from localities near Plymouth. Journal of the Marine Biological Association of the United Kingdom 62: 385-401. Wilson, E. B. 1883. Observations on the early developmental stages of some polychaetous annelides. Studies from the Biological Laboratory, Johns Hopkins University 2: 271-299. Wilson, W. H. 1991. Sexual reproductive modes in polychaetes: classification and diversity. Bulletin of Marine Science 48: 500-516. Woltereck, R. 1902. Trochophora-Studien I. Über die Histologie der Larve und die Entstehungdes Annelids bei den Polygordius-Arten der Nordsee. Zoologica, Stuttgart 34: 1-71. Woltereck, R. 1904. Beiträge zur praktischen Analyse der Polygordius- Entwicklung, nach dem Nordsee- und Mittelmeertypus. Archiv für Entwicklungsmechanik der Organismen, Berlin 18: 377-403. Wray, G. A. 1995. Evolution of larvae and developmental modes. Pp. 413-447. In L. McEdward (ed.), Ecology of Marine Invertebrate Larvae., CRC Press, Boca Raton. Yokouchi, K. 1991. Seasonal distribution and food habits of planktonic larvae of benthic polychaetes in Volcano Bay, southern Hokkaido, Japan. Ophelia Supplement 5: 401-410. Young, C. M., Vásquez, E., Metaxas, A. and Tyler, P. A. 1996. Embryology of vestimentiferan tube worms from deep-sea methane/sulphide seeps. Nature 381: 514-516. Yun, S. G. and Kikuchi, T. 1991. Larval development and settlement of Chone duneri Malmgren (Polychaeta: Sabellidae). Publications from the Amakusa Marine Biological Laboratory, Kyushu 11: 31-42. Zal, F., Jollivet, D., Chevaldonné, P. and Desbruyères, D. 1995. Reproductive biology and population structure of the deep sea hydrothermal vent worm Paralvinella grasslei (Polychaeta: Alvinellidae) at 13°N on the east Pacific rise. Marine Biology 122: 637-648. Zottoli, R. A. 1974. Reproduction and larval development of the ampharetid polychaete Amphicteis floridus. Transactions of the American Microscopical Society 93: 78-89. Zottoli, R.A. 1999. Early development of the deep-sea ampharetid (Polychaeta: Ampharetidae) Decemunciger apalea Zottoli. Proceedings of the Biological Society of Washington 112: 199-209.

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CHAPTER

7

Larval Ecology of the Annelida Pei-Yuan Qian and Hans-Uwe Dahms

7.1 INTRODUCTION Through the course of evolution by natural selection, annelid larvae have well adapted to their particular niches and may little resemble the adults of their own species, neither morphologically nor physiologically. Larvae and adults — although they share the same genome — show distinct biological and ecological differences — often using different habitats and food sources. Despite their ecological dissimilarity, the success of the one form as the early phase of the life cycle determines the existence and reproduction of a later phase (Hadfield 1998). These sorts of reproductive patterns comprising two or more ecologically distinct phases are termed complex life cycles (Strathmann 1990). The transition between phases of a complex life cycle often consists of an abrupt morphological, physiological, structural, functional, and ecological change termed metamorphosis (Giangrande 1997). The larger the difference between adult and larval lifestyles, the more drastic the metamorphosis will be. Jägersten (1972) argued that complex life cycles are the original condition for marine invertebrates because the occurrence of free-spawning small eggs, external fertilization and subsequent indirect, planktotrophic development represents a uniformly plesiomorphic form of life cycle. This way of interpreting life history evolution still represents a guiding scheme for the evolution of invertebrate reproductive modes for the fact that it provides the most reasonable scenario for the ancestral mode of reproduction in marine invertebrates (but see Olive 1985; Rouse and Fitzhugh 1994; Haszprunar et al. 1995; Rouse 1999 for alternative views). Among the Annelida the life cycle containing a larval stage is not restricted to the Polychaeta. The second largest group, the Clitellata (Oligochaeta and Hirudinida) can show indications of larval embryonic features in their development as well (Rouse 2000a). Within the annelid taxon Hirudinida (=leeches), members of the Glossiphonidae, brood eggs and feed their young after hatching, with Department of Biology and Coastal Marine Laboratory, Hong Kong University of Science and Technology, Clear Water Bay, Hong Kong SAR, China

& Reproductive Biology and Phylogeny of Annelida variation in the duration and degree of care among individual species (Paez et al. 2004). The basis for understanding the distribution and variation of parental care is the selective balance between the improvement in offspring fitness and the decrement to future parental fitness that result from the provision of care. A principal advantage of care in glossiphoniid leeches is thought to be protection of offspring from predation (Kutschera and Wirtz 1986). Helobdella parents take care of both eggs and young. Following fertilization, H. papillornata lays 20 to 60 eggs that are attached directly to the ventral surface of the parent. After approximately 14 days, the eggs hatch and the newly emerged young remain attached to the parent’s ventral surface for up to 60 days until they are capable of hunting on their own. Throughout the care period, the parent provides protection from predators, ventilates the eggs/young to ensure that they receive sufficient oxygen and provides prey to the juveniles after hatching (Paez et al. 2004). Polychaete annelids, on the other hand, develop through free living larval stages in most cases (Qian 1999, Fig. 7.1). To make our task manageable, we therefore have chosen to emphasize patterns associated with the ecology of polychaete development. However, by omitting patterns specific to asexual reproduction, and to a large extent, taxa that are parasitic, holoplanktonic, or have a direct mode of development, we have let aside a considerable portion of the Annelida (see Giese and Pearse 1975, Giese and Kanatani 1987). The diversity and complexity of polychaete larvae in form, developmental patterns, behavior, nutritional characteristics, and ecology (cf. Giangrande 1997) have attracted considerable attention from marine biologists. This will be dealt with elsewhere (Chapter 6). Larval diversity of the polychaetes offers opportunities and constraints for functional morphology, behavior, ecology, and life cycle evolution (cf. Chia 1989). However, although the early stages of a number of polychaetes have been known since the late nineteenth century, their behavioral and ecological diversity is still poorly understood. The point at which metamorphosis to a benthic life form begins is somewhat controversial, and descriptions of various species differ depending on the perspective of the authors. Some authors consider the appearance of the first true metameric somite as marking the beginning of the juvenile stage, regardless of the larval or juvenile life-style or ecological niche occupation for any particular species. This system has considerable advantage, because it designates a specific developmental event that can be identified in every species. This transitional point is also more easily comparable with the definition of metamorphosis or larva-juvenile transformation in other animal phyla. However, most annelid embryologists have tended over the years to differentiate larval versus juvenile stages more on the basis of successive developmental shifts in life style than on the specific chronology of morphogenetic changes. Any system used, is complicated by the tremendous variety of metamorphosis patterns found among polychaetes. Some species may go through three or

Larval Ecology of the Annelida

&

Fig. 7.1 Generalized scheme of possible life histories of the Annelida. From Qian, P.Y. 1999. Hydrobiologia 402: 239-253, Fig. 1.

four distinguishable phases during the transition from a trochophore larva to a definitive juvenile (Qian and Chia 1989). Special terms are used to identify some of these phases, e.g. to identify specialized larvae or early juveniles of certain species, but these are beyond the scope of this chapter. Useful guides to larval polychaete development include Lacalli (1980), Bhaud and Cazaux (1987), Rouse and Fauchald (1997), Rouse (1999, 2000a, b), and Young (2002). Comparative larval ultrastructure has recently been reviewed by Heimler (1988). Levin and Bridges (1995) presented a classification scheme for invertebrate larval development that can readily be applied for annelids as well. It consists of four categories: A. dispersal potential and recruitment, B. mode of larval nutrition, C. location or site of development, and D. the morphogenesis involved in development. These categories allow highlighting the remarkable range of diversity in annelid larval development from a biological and an ecological perspective.

7.2 LARVAL ECOLOGY 7.2.1 Dispersal Potential and Abiotic Factors Most marine invertebrate taxa as the Annelida have a dispersal stage at some point in their life histories. The main likely adaptive benefits of freeliving larvae for polychaete species that are slow moving or sessile as adults are as follows (see also Pechenik 1999): - dispersal and genetic exchange between geographically separated populations of the same species,

&

Reproductive Biology and Phylogeny of Annelida

- connectivity of various populations of a given species, - rapid colonization of areas following local extinctions and, - lack of direct competition with adults for food or space during development due to habitat segregation. How far a larva can disperse depends on how long the larval form can be maintained and on the velocity and direction of water currents in which the larvae stay. Pronounced dispersal generally comes along with larger genetic homogeneity between geographically disjunct adult populations, greater ability to recolonize areas after local extinctions, reduced rates of speciation, and greater species longevity (Levin and Bridges 1995). Scheltema (1971) has introduced several terms that explicitly address larval dispersal capabilities of marine invertebrates. Teleplanic (tel = far, planos = wanderer) larvae have planktonic periods exceeding two months and possibly lasting a year or more. Teleplanic forms beside other invertebrates are common among spionid and chaetopterid polychaetes (Scheltema 1992). This author also suggests that teleplanic larvae are capable of transoceanic dispersal in both the Atlantic and the Pacific Oceans from one continent to the other in east-west direction following the major oceanic currents. Until now there is no agreement on descriptive terms for larvae with more limited dispersal potential. Scheltema (cf. Levin and Bridges 1995) has offered the term anchiplanic (anchi = near, planos = wanderer) for larvae that remain in the plankton only for a few hours to a few days. Abbreviated dispersal ability can be found in some representatives of almost every animal phylum, and for the Annelida it is particularly common among certain polychaetes such as the Spirorbinae. At the other end of a motilityspectrum there are non-planktonic larvae, which are considered aplanic. Examples of aplanic larvae include forms that are competent to settle as soon as they emerge from adults (as in the exogoniin syllid and fabriciin sabellid polychaetes). These will also include lecithotrophic larvae that develop fully during encapsulation (Scheltema 1971). Larval behavior and locomotion. Swimming behavior of polychaete larvae is important for active horizontal and particularly vertical larval locomotion, for actively selecting settlement sites and delaying metamorphosis until a suitable substratum is found (Young 1990). Considering the role of oceanographic processes for recruitment larval behavior exhibited during the planktonic dispersal phase is also of resurrected interest for dispersal processes (Dahms and Qian 2004). Generally, marine invertebrate larvae control their horizontal distribution and dispersal by navigating vertically in the water column (Young 1995). Most polychaete larvae and planktonic juveniles swim slowly relative to horizontal currents. They may be able to exert some control over their vertical distribution by constantly swimming or by adjusting sinking rates (for instance, by erecting larval chaetae) (see Bhaud 1990, Bhaud et al. 1990). Bhaud and Cazaux (1990) investigated the buoyancy of the aulophora larvae of terebellid polychaetes. Aulophora larvae build a gelatinous tube open at both ends. It has previously been assumed that the tube is a floating

Larval Ecology of the Annelida

&!

device, but these authors demonstrated that the isolated tubes actually sink faster than tubes containing larvae. This apparent paradox was resolved by the discovery of a long strand of mucus secreted by the primary tentacle and extending from the upper opening of the tube. This mucus strands slow down the sinking by increasing drag. Salinity and temperature. Effects of abiotic parameters such as salinity and temperature on larval development are of prime importance for marine polychaetes and therefore have been studied for a number of species from different habitats. As for Hydroides elegans, these parameters, in combination with food, were investigated in 4 laboratory experiments by Qiu and Qian (1997). In these studies three 2-factor experiments tested the effects of salinity (15 to 35‰ ) and temperature (15 to 30°C) on the survival and duration of development from newly-released oocytes to 2-cell, 2-cell to blastula, and blastula to the trochophore stage respectively (Fig. 7.2). A fourth 3-factor experiment tested the effects of salinity and temperature, and the concentration of the single-cell alga Isochrysis galbana (0 to 106 cells ml –1 ) on survival, settlement, and duration of development from trochophore to newly settled juvenile. Within the experimental range, temperature had no effect on survivorship, but low temperature led to a longer duration of development. Low salinity reduced survivorship and settlement, and lengthened the duration of development. Low food concentration reduced survivorship and settlement, and lengthened the duration of development from trochophore to newly-settled juvenile. At concentrations 35% larvae survived through the 10 d experiment but lost their ability to become competent (Fig. 7.3). Percentages of trochophores reaching settlement were similar at 104, 105, and 106 cells ml–3. Duration of development was shortest at concentrations of 105 cells ml–1, while trochophores at 104 and 106 cells ml-1 had similar but longer durations of development. The results of Qiu and Qian (1997) suggest that in Hong Kong waters, the decrease in salinity during the summer overrides the benefits of high temperature and is responsible for the decline in H. elegans settlement. The increase in phytoplankton concentration from early spring to early summer may contribute to the formation of settlement peaks. Temperature, however, does not seem to be a limiting factor for early development and settlement of H. elegans.

7.2.2 Larval Trophic Ecology Feeding mechanisms and rates vary among types and stages of polychaete larvae. This variation may affect the growth rate from even a small egg to a large juvenile, with consequences for adult fitness. Many taxa develop feeding larvae with large and elaborate structures that are absent in nonfeeding larvae of related species and in postlarval stages (see Chapter 6). The structures peculiar to feeding larvae suggest that much of the larval body has been shaped by the functional requirements of acquiring food. Field ecologists have very much avoided the study of larval stages in their natural environment due to difficulties in accessing them in the field — and

Fig. 7.2. Effects of salinity and temperature on fertilization and embryonic development of Hydroides elegans. Expt. I: Newly-released oocyte to 2-cell. A. survivorship. B. duration of development. Oocytes all died at salinities 95% of eggs were fertilised within 15 minutes after the gametes were mixed. (Pechenik and Qian 1998). In contrast, sperm concentrations > 107 sperm ml–1 are required to achieve fertilisations in serpulin Galeolaria caespitosa (Kupriyanova, unpubl (Fig. 12.3A)). Fertilisation rates in this species are also influenced by gamete age, male-female

#!$ Reproductive Biology and Phylogeny of Annelida compatibility, and ambient water temperature. The gamete traits of G. caespitosa apparently enable this gregarious serpulid to perform under conditions of high population density (Kupriyanova 2002). Little is known about the fertilisation biology in the many brooding serpulins and spirorbins. Gee (1965) reported that in Spirorbis spirorbis gametes shed through the segmental organs and fertilisation occurs inside the tube. Broadcasting was previously assumed to be a common fertilisation mechanism for all brooding tube-dwellers. However, discovery of a spermatheca in Spirorbis spirorbis (Daly and Golding 1977; Picard 1980) and paired ones in Salmacina sp. (Rouse 1996c) suggests more complex fertilisation biology in brooding species. No information is available on efficiency of fertilisation in brooding species and the assumption that the fertilisation rate for incubating species is high may be not substantiated. Spirorbins are capable of self-fertilisation, although it does not occur as readily as cross-fertilisation. Self-fertilisation could not be demonstrated in the hermaphrodite serpulid Salmacina (Nishi and Nishihira 1993). Brooding of larvae is quite common among serpulids. Tube incubation is known for species of Filograna and for Paraprotula apomatoides. Brooding in ovicells on the tube occurs in a variety of ways. Chitinopoma serrula produces pouches with twin chambers at the tube orifice, each containing 10-20 larvae. Ovicells of Microprotula ovicellata resemble swellings encircling the distal part of the tube. The ovicells in Rhodopsis pusilla are wide inverted pouches arranged one by one along the length of the tube. Pseudovermilia pacifica has a cup- to dome-shaped ovicells over the entrance of the tube. Paraprotis dendrova broods embryos inside the branchial crown on an appendage growing from the mouth(Fig. 12.3C). Metavermilia ovata holds developing embryos inside the base of its branchiae, whereas Floriprotis sabiuraensis broods in pockets of the thoracic membranes. Brooding in gelatinous masses near the tube mouth is found in Protula tubularia. The list above may not be exhaustive (for references see Table 12.1) and further studies may reveal additional incubating methods in the group. In all these cases, the larvae are likely to be lecithotrophic, though whether some species release planktotrophic larvae from brooding structures is yet unknown. Spirorbinae all brood their lecithotrophic larvae either in the parental tube or in the opercular brood-chambers. Tube incubation types vary according to the methods of embryo anchorage within the tube. Embryos lie free in the tube (Paralaeospira), they form an egg string attached to the tube by a posterior filament (Spirorbis), adhere to each other and to the tube wall in Circeis (Fig. 12.3B) and Paradexiospira. They also may be attached anteriorly to a thoracic funnel-like stalk or epithelial oviducal funnel (Protolaeospira, Helicosiphon, Romanchella, Metalaeospira and Eulaeospira). Opercular incubation is found in more than a half of spirorbin species. The brood chambers of the some spirorbins (Amplicaria, Pileolaria, Nidificaria, Vinearia, Simplaria, Protoleodora and Bushiella) are formed by invagination of the opercular ampulla itself. Such brood chambers are used for a number of broods. The primary non-brooding operculum is either

Sabellida

#!%

Fig. 12.5 Evolution of reproduction. Cladogram showing feeding (planktotrophic) larvae that result from broadcast spawning as a derived condition within Serpulidae. The plesiomorphic condition is to have lecithotrophic larvae that are retained by the parent for at least part of development. From Kupriyanova, E. K. (2003). Hydrobiologia 496: 105-114, Fig. 4.

shed after the chamber is formed (e.g. Pileolaria) or is fused to the chamber for additional embryo protection (Bushiella). When breeding ceases, the brooding chamber may be replaced by a non-brooding operculum, which may again be later replaced by a new brood chamber. Brood chambers of other spirorbins (Neodexiospira, Janua, Pillaiospira, Leodora) are formed distally by the calcified opercular plate outside of the opercular ampulla. Every brood chamber is used for only one brood and is shed to liberate larvae. The cytological processes during fertilisation in serpulids have been studied for Pomatoceros triqueter, Hydroides elegans and H. norvegicus, Ficopomatus enigmaticus, Galeolaria caespitosa, and Spirorbis spirorbis (reviewed in Kupriyanova et al. 2001). A classic series of studies on the ultrastructure of sperm-egg interaction in Hydroides dianthus by (Colwin and Colwin 1961; Colwin and Colwin 1961a, 1961b) addressed the functional significance of the acrosome reaction, and the sequence of events during the fusion of the gamete membrane. Members of Frenulata almost all shed spermatophores with long filaments into the surrounding seawater (Bakke 1990). These are gathered by females and fertilization is thought to be internal, at least in Siboglinum. In this genus, larvae are brooded in the tube of the female, at least in the species for which there is currently information. In vestimentiferan siboglinids there are no spermatophores. Rather they form masses that

Spherical long acrosome Sabellaria alveolata Spherical long acrosome Sabellaria cementarium

Lygdamis indicus Lygdamis muratus Phalacrostemma cidariophilus Phragmatopoma californica Phragmatopoma lapidosa

Idanthyrsus

Sabellariidae

Spherical long acrosome

Owenia ‘fusiformis’ Spherical

Oweniidae

Sperm head

Minor taxon

Major taxon

No

No

Free-spawning

Free-spawning

Free-spawning

Brooding or freespawning Free-spawning

No

70 000

Fecundity (eggs per female)

Free-spawning Free-spawning

85

70

Egg µm

No No

No

Sperm storage

planktotrophic

planktotrophic

planktotrophic

planktotrophic planktotrophic

planktotrophic

planktotrophic

Larval development

Smith and Chia 1985

Pasteels 1965; Wilson 1970

Amieva et al. 1987; Dales 1952; Thomas 1994 Eckelbarger 1976, 1984; Mauro 1975

Bhaud and Fernandez-Alamo 2001; Jamieson and Rouse 1989 Bhaud 1975 Wilson 1977 Bhaud 1969

Dauvin and Gillet 1991; Gentil et al. 1990; Ménard et al. 1989; Thiébaut and Dauvin 1992; Wilson 1932

References

Table 12.1 Reproduction in Sabellida Not comprehensive; major reviews should also be consulted (Gardiner and Jones 1993; Giangrande 1997; Kupriyanova et al. 2001; Rouse and Fitzhugh 1994; Southward 1993; 2000)

#!& Reproductive Biology and Phylogeny of Annelida

Cylindrical ?

No ? Cylindrical ? Spherical

Spherical

Branchiomma lucullana

Chone duneri Chone ecaudata Demonax medius Demonax micropthalmus Euchone analis

Spherical

300400

Spherical

Branchiomma luctuosum

Laonome albicingillum Megalomma vesiculosum

240 200 200 250

Elongate

No

No

Yes

150

250

150200 200600 120

Amphiglena

Yes Yes

Elongate Elongate

Caobangia Amphicorina

Sabellidae Sabellinae

Sabellidae

125200

Fabriciinae

Sabellariidae

Yes

Elongate

Sabellaria floridensis Sabellaria spinulosa Sabellaria vulgaris

1100-1300

1000-4000

5-10

10 or less

2-7

planktotrophic

Free-spawning

Free-spawning

Intratubular brooding

Free-spawning Extratubular brooding Extratubular brooding Free-spawning

Extratubular brooding

Free-spawning

Intratubular brooding

ovoviviparous Intratubular brooding

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic lecithotrophic lecithotrophic lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic lecithotrophic

lecithotrophic

Free-spawning Intratubular brooding

planktotrophic

planktotrophic

Free-spawning

Free-spawning

Table 12.1 contd

Wilson 1936

Hsieh 1995, 1997

Rouse 1995a, 1995b, 1996c; Rouse and Fitzhugh 1994 Jones 1974 Rouse 1992a, 1992b; Rouse and Fitzhugh 1994 Rouse and Gambi 1997, 1998a, 1998b Sordino and Gambi 1992; Sordino and Gambi 1994 Dragesco-Kernéis 1980; Rouse and Fitzhugh 1994 Yun and Kikuchi 1991a, b Okuda 1946 McEuen et al. 1983 Kerby 1972; Rouse and Fitzhugh 1994 Curtis 1977

Pasteels 1965; Douglas P. Wilson 1970 Eckelbarger 1975

Eckelbarger 1977

Sabellida

#!'

Serpulidae Filograninae

Sabellidae Sabellinae

Major taxon

Table 12.1 contd

Spherical

Myxicola infundibulum Perkinsiana antarctica

Rhodopsis pusilla

Protula tubularia

Elongate

Microprotula ovicellata Protula cf. tubularia Protula globifera Spherical Protula palliata Spherical

Yes

Elongate

Elongate

Yes

Spherical

Sabella spallanzanii Terebrasabella heterouncinata

Filograna implexa

No

Spherical

No

Sperm storage

Perkinsiana riwo

Oval

Sperm head

Minor taxon

10,000 – 13, 000

4000080000 54-374

Fecundity (eggs per female)

78-90 1

80

85

180200 80

250

8

235

142

Egg µm

Brooding in gelatinous mass outside tube Brood chambers on tube

Free-spawning

Brood chambers on tube Free-spawning

Brooding in tube

Intratubular brooding

Brooding on branchial crown Free-spawning

Brooding on branchial crown

Free-spawning

Brooding or freespawning

lecithotrophic

lecithotrophic

planktotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

Larval development

Ben-Eliahu and ten Hove 1989; Nishi and Yamasu 1992a

Salensky 1882

Tampi 1960 Franzén 1956 Kupriyanova et al. 2001

Nelson-Smith 1971; Nishi 1993; Rouse 1996c Uchida 1978

Fitzhugh and Rouse 1999; Simon and Rouse 2005

Giangrande et al. 2000

Dean et al. 1987; Rouse and Fitzhugh 1994 Gambi and Patti 1999; Rouse and Fitzhugh 1994 Rouse 1996b

References

#" Reproductive Biology and Phylogeny of Annelida

Serpulidae Serpulinae

Serpulidae Filograninae

Spherical

Spherical

Elongate

Elongate

Elongate

Galeolaria Spherical caespitosa Galeolaria hystrix Spherical Hydroides dianthus Spherical

Chitinopoma serrula Crucigera irregularis Crucigera zygophora Ditrupa arietina Ficopomatus enigmaticus Ficopomatus miamiensis Ficopomatus uschakovi Floriprotis sabiuraensis

Chitinopoma arndti

Salmacina sp.

Salmacina cf. dysteri

60-62 40000 45 600-80000

No No

4000

60-64 500-20000

50

80 60

20

26

No

No

70

180200 90

120150

Free-spawning Free-spawning

planktotrophic planktotrophic

planktotrophic

planktotrophic

Free-spawning Brooding in pockets of the thoracic membrane Free-spawning

planktotrophic

planktotrophic planktotrophic

planktotrophic

planktotrophic

lecithotrophic

lecithotrophic

lecithotrophic

Free-spawning

Free-spawning Free-spawning

Free-spawning

Brood chambers on tube Brood chambers on tube Free-spawning

Brooding in tube

Brooding in tube

#"

Andrews and Anderson 1962; Grant 1981 Kupriyanova et al. 2001 Colwin and Colwin 1961a, b Scheltema et al. 1981 Table 12.1 contd

Bailey-Brock 1985; Uchida 1978

Hill 1967

Charles et al. 2003 Dixon 1981; Morris et al. 1980 Lacalli 1976

Strathmann 1987

Dons 1933; Franzén 1982; Thorson 1946 Strathmann 1987

Zibrowius 1983

Franzén 1956, 1958; Nishi and Nishihira 1993; Nishi and Yamasu 1992b; Rullier 1960 Rouse 1996c

Sabellida

Serpulidae Serpulinae

Major taxon

Table 12.1 contd

Pomatoceros terranovae Pomatoleios Spherical kraussii Pseudochitinopoma occidentalis Pseudovermilia cf. pacifica

Spherical

60

No

60

60-80

80

67

45

No

Hydroides fusicola Marifugia cavatica Metavermilia cf. ovata Paraprotis dendrova Paraprotula apomatoides Placostegus tridentatus Pomatoceros triqueter

45-63

45-53

Egg µm

No

No

Hydroides ezoensis Spherical

Spherical

No

Spherical

Hydroides elegans

Sperm storage

Sperm head

Minor taxon

2500

40

Fecundity (eggs per female)

Brood chambers on tube

Free-spawning (?)

Free-spawning

Free-spawning

Free-spawning

Free-spawning

Free-spawning Free-spawning Brooding inside the base of branchiae Brooding on branchial crown Brooding in tube

Free-spawning

Brooding or freespawning Free-spawning

planktotrophic

planktotrophic

planktotrophic

planktotrophic

lecithotrophic

planktotrophic

planktotrophic

planktotrophic

Larval development

Kupriyanova et al. 2001

Hess 1993

Crisp 1977; Sawada 1984

Dorresteijn and Luetjens 1994; Føyn and Gjøen 1954; Segrove 1941 Kupriyanova et al. 2001

Franzén 1956

Nishi 1992; Nishi and Yamasu 1992c Uchida 1978

Carpizo-Ituarte and Hadfield 1998; Franzén 1956; Matsuo and Yoshioshi 1983 Matuso and Ko 1981; Miura and Kajihara 1981 Matsuo and Yoshioshi 1983 Matjasic and Sket 1966 Kupriyanova et al. 2001

References

#" Reproductive Biology and Phylogeny of Annelida

Serpulidae Spirorbinae

Serpulidae Serpulinae

Yes

Yes

Spirorbis cuneatus Spirorbis (Velorbis) gesae Spirorbis inornatus

Spirorbis rothlisbergi Spirorbis rupestris 110180

150230

Up to 35

Up to 70

About 15

20

100150

Spirorbis corallinae

Eggs in tube 1

Eggs in tube 1

Eggs in tube 1

Eggs in tube 1 Eggs in tube 1

Eggs in tube 1

Eggs in tube 1

Free-spawning

60 Up to 24

Free-spawning

65

Spirorbis bifurcatus

Free-spawning

83

No

Free-spawning

80

Spherical

No

Free-spawning

Spherical

Free-spawning

Serpula columbiana Serpula vermicularis Spirobranchus corniculatus Spirobranchus giganteus Spirobranchus polycerus Spirobranchus tetraceros 65

Brooding in tube

Semivermilia cf. uchidai

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic lecithotrophic

lecithotrophic

lecithotrophic

planktotrophic

planktotrophic

planktotrophic

planktotrophic

planktotrophic

planktotrophic

Table 12.1 contd

Gee and Knight-Jones 1962; Picard 1980

Knight-Jones and KnightJones 1977; Silva and Knight-Jones 1962 Gee 1964 Knight-Jones and KnightJones 1995 Gee 1967; L’Hardy and Quiévreux 1964; Picard 1980 Knight-Jones 1978

Knight-Jones 1978

Gaikwad 1988

Lacalli 1976

Allen 1957

Smith 1984

Strathmann 1987; Young and Chia 1982 Franzén 1956

Kupriyanova et al. 2001

Sabellida

#"!

Serpulidae Spirorbinae

Major taxon

Table 12.1 contd

Fecundity (eggs per female)

Janua pagenstecheri

Elongate

Circeis sp. Elongate Paradexiospira Elongate (Spirorbides) vitrea Yes

Yes

Yes

Circeis armoricana

Circeis oshurkovi Circeis paguri

Yes

Spirorbis strigatus Spirorbis tridentatus Elongate

120150

7-25

4-65

10-12

Up to 9 110- Up to 50 180 140 by 6-295 95

1-90 (usually 10-60)

Yes

Elongate

110190

Egg µm

Spirorbis spirorbis

Sperm storage

Up to 50

Sperm head

Spirorbis spatulatus

Minor taxon

Brood chambers 1

Eggs in tube 2 Eggs in tube 2

Eggs in tube 2 Eggs in tube 2

Eggs in tube 2

Eggs in tube 1 Eggs in tube 1

Eggs in tube 1

Eggs in tube 1

Brooding or freespawning

lecithotrophic

lecithotrophic lecithotrophic

lecithotrophic lecithotrophic

lecithotrophic

lecithotrophic lecithotrophic

lecithotrophic

lecithotrophic

Larval development

Daly 1978a, 1978b; Daly and Golding 1977; Franzén 1956; Knight-Jones 1951; Picard 1980 Knight-Jones 1978 Franzén 1956; Picard 1980; Silva 1962 Al-Ogily and Knight-Jones 1981; Picard 1980; Rzhavsky and Britayev 1988 Rzhavsky 1998 Al-Ogily and Knight-Jones 1981; Knight-Jones and Knight-Jones 1977 Franzén 1956 Franzén 1956; Hess 1993; Picard 1980; Quievreux 1962 Franzén 1956; Knight-Jones et al. 1974; Picard 1980

Knight-Jones 1978

References

#"" Reproductive Biology and Phylogeny of Annelida

Serpulidae Spirorbinae

Yes

60

Eggs in tube 3

5-40

Eggs in tube 4 Eggs in tube 4 Eggs in tube 4

About 200 >200

Eggs in tube 3

Up to 20 About 90

Eggs in tube 3

About 10

Eggs in tube 3

Brood chambers 1

Brood chambers 1

8 1-14

Brood chambers 1

8

Brood chambers 1

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

Brood chambers 1

Brood chambers 1

80

Neodexiospira foraminosa Neodexiospira formosa Neodexiospira lamellosa Neodexiospira pseudocorrugata Neodexiospira steueri Paralaeospira levinseni Paralaeospira malardi Paralaeospira parallela Eulaeospira convexis Helicosiphon platyspira Helicosiphon biscoensis Metalaeospira pixelli

lecithotrophic

Brood chambers 1

8-12

140 by 40 160 2-60

Neodexiospira alveolata Neodexiospira brasiliensis

Table 12.1 contd

Knight-Jones and Knight-

Knight-Jones et al. 1973

Knight-Jones 1978

Knight-Jones et al. 1974

Harris 1968; Knight-Jones et al. 1974 Knight-Jones et al. 1974; Knight-Jones 1972 Knight-Jones and Walker 1972; Vine 1977 Picard 1980; Quievreux 1962 Vine 1977

Knight-Jones et al. 1974; Knight-Jones 1972 Knight-Jones et al. 1974

Fauchald 1983; Knight-Jones et al. 1975; Rzhavsky and Britayev 1984 Nishi and Yamasu 1992d

Okuda 1946

Sabellida

#"#

Serpulidae Spirorbinae

Major taxon

Table 12.1 contd

Metalaeospira tenuis Protolaeospira (P.) eximia Protolaeospira (P.) pedalis Protolaeospira (P.) striata Protolaeospira (P.) tricostalis Protolaeospira (Dextralia) stalagmia Romanchella pustulata Romanchella quadricostalis Romanchella solea Bushiella (B.) abnormis Bushiella (Jugaria) atlantica Bushiella sp. Nidificaria nidica Nidificaria palliata

Minor taxon

Elongate

Sperm head

Yes

Sperm storage

100160

Egg µm

Eggs in tube 4 Eggs in tube 4

20-106 About 50

Eggs in tube 4 Brood chambers 2 Brood chambers 2

Up to 20 20-107 Up to 3

Brood chambers 2 Brood chambers 2 Brood chambers 2

Eggs in tube 4

Up to 25

2-4 20

Eggs in tube 4

165-270

Eggs in tube 4

Eggs in tube 4

Up to 17 > 200

Eggs in tube 4

About 30

Up to 13

Brooding or freespawning Eggs in tube 4

Fecundity (eggs per female)

lecithotrophic lecithotrophic lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic

Larval development

Franzén 1956 Knight-Jones 1978 Knight-Jones 1978

Knight-Jones 1978

Vine 1977 Hess 1993

Canete and Ambler 1990; Knight-Jones 1978 Knight-Jones, 1973

Knight-Jones and Walker 1972

Knight-Jones 1973

Knight-Jones and KnightJones 1994 Picard 1980

Hess 1993

Knight-Jones 1973

References

#"$ Reproductive Biology and Phylogeny of Annelida

Elongate

Elongate

Elongate

Frenulata

Vestimentifera

Osedax ?

Yes

Yes?

115150

100+

130600

230

Eggs in tube 1. Egg string attached to tube by a filament. Eggs in tube 2. Adhered directly to wall. Eggs in tube 3. Egg string free in tube Eggs in tube 4. Attached anteriorly to a thoracic stalk Brood chambers 1. Formed by calcified plate outside opercular ampulla Brood chambers 2. Formed by invagination of opercular ampulla

Siboglinidae

Serpulidae Spirorbinae

Pileolaria berkeleyana Pileolaria daijonesi Pileolaria dakarensis Pileolaria lateralis Pileolaria marginata Pileolaria militaris Elongate Pileaolaria spinifer Pileolaria tiarata Protoleodora uschakovi Simplaria potswaldi Simplaria pseudomilitaris Vinearia zibrowii

Free-swimming larvae

Brood in tube, some may have free swimming larvae Free-swimming larvae

Brood chambers 2

2-4 5-100

Brood chambers 2 Brood chambers 2

About 30 8-20

Brood chambers 2 Brood chambers 2

About 15 Up to 16 Brood chambers 2 Brood chambers 2 Brood chambers 2 Brood chambers 2

Brood chambers 2 Brood chambers 2

About 10 About 5

9-14 Up to 12 Up to 10 Up to 150

Brood chambers 2

4-56

lecithotrophic?

lecithotrophic

lecithotrophic

lecithotrophic

lecithotrophic lecithotrophic

lecithotrophic lecithotrophic lecithotrophic lecithotrophic

lecithotrophic lecithotrophic

lecithotrophic lecithotrophic

Gardiner and Jones 1993; Hilário et al. 2005; Marsh et al. 2001; Young et al. 1996 Rouse et al. 2004

Bakke 1990; Southward 1975, 1993; 2000

Knight-Jones 1978

Knight-Jones 1978 Knight-Jones et al. 1974

Franzén 1958 Knight-Jones 1978 Knight-Jones 1978 Knight-Jones 1984

Knight-Jones 1978 Knight-Jones 1978

Knight-Jones 1972 Knight-Jones 1978

Hess 1993

Sabellida

#"%

#"& Reproductive Biology and Phylogeny of Annelida appear to be spermatozeugmata, with the sperm embedded in a sticky matrix (Southward and Coates 1989). The spermatozeugmata have elongate tails in Ridgeia piscesae, as seen in the spermatophores of the other siboglinids (Southward and Coates 1989). It has been suggested that Riftia pachyptila is a free spawner (Carey et al. 1989), but Southward and Coates (1989) argue that this was an artefact because the observations were based on a shocked animal and the material emitted seen by Carey et al. (1989) was bundles of late spermatids. Observations of in situ spawning of Riftia pachyptila by Van Dover (1994) indicates that Southward and Coates (1989) were correct and that spermatozeugmata are spawned into the water and attach to females. The females are then triggered to spawn and after fertilization the eggs are expelled by the female into the surrounding water where development occurs (Van Dover 1994). Van Dover (1994) argued that fertilization was probably internal, based on observations by Jones (1981) that sperm were found in the genital tracts of females. This has now been confirmed in a study by Hilário et al. (2005) showing sperm storage and internal fertilization in five different vestimentiferans from both hydrothermal vents and cold seeps. They found that sperm are stored in a spermatheca at the posterior end of each of the pair of oviducts. Hilário et al. (2005) also showed experimentally that most vestimentiferan eggs are fertilized internally with a rate typically lower than 100%. Meiosis is completed after eggs are released from the female, and the dispersal phase includes the entire embryonic period.

12.7 LARVAL DEVELOPMENT Wilson (1932) elegantly described the development of the ‘mitraria’ larva of Owenia and its ‘catastrophic metamorphosis’ that results in much of the larval body being cast-off and a juvenile worm settling to the bottom. The larvae pass through a normal trochophore phase before becoming a planktotrophic mitraria. Larvae similar to the mitraria of Owenia have been described for Myriochele (Thorson 1946). The developmental pattern is similar in all Sabellariidae studied to date and is reviewed by Eckelbarger (1978). How the larvae feed has yet to be elucidated. There is a distinct extension of the prototroch in the buccal region that may help to capture food (Rouse 2000). Larvae can stay in the plankton for extended periods before settling and they have long bundles of provisional chaetae that appear to be anti-predation devices. These are held in place along the body by special grasping cilia in the telotroch (Wilson 1929, 1977). The ultrastructure of the pair of peristomial palps has been described in Phragmatopoma larvae by Amieva and Reed (1987). Larval development is planktonic with feeding trochophore larvae in well known and commercially important fouling serpulins such as the speciose group Hydroides, as well as common and widely distributed taxa Crucigera, Ficopomatus, Galeolaria, Pomatoceros, Serpula and Spirobranchus (Fig. 12.4A). In total, 26 species from 10 genera are shown to have planktotrophic

Sabellida

#"'

larvae (reviewed by Kupriyanova et al. 2001). Developmental events in planktotrophic larvae are very similar. The zygotes undergo synchronous holoblastic cleavages up to the blastula stage. The uniformly ciliated blastula develops into an early trochophore with a single equatorial ciliary band, the prototroch that separates a rounded episphere from a conical hyposphere. Later, a second ciliary ring, the metatroch, develops below the prototroch and a band of short feeding cilia forms between the prototroch and metatroch. Suspension feeding by serpulid larvae is achieved by use of the opposed band system (the prototroch and the metatroch), as described for Serpula columbiana by Strathmann et al. (1972). On the right side of the episphere, an ocellus forms. Next, the larva develops the left ocellus identical to the right one. After this stage the growth is mostly confined to the hyposphere and the larva elongates and develops three chaetigerous segments. Before the settlement a small fourth trunk segment is delineated and paired branchial rudiments appear posterior to the metatroch. Non-feeding planktonic development reported for Serpulidae is known in Protula sp. by Tampi (1960). The development is very similar to that of feeding larvae but the active gut is still not formed by the 3-chaetiger stage. Non-feeding development in Protula sp. from Florida observed by Pernet (pers. comm.) was similar to that described by Tampi (1960). Although Serpulinae are best known for their planktonic, feeding, trochophore larvae, they show a surprising range of brooding mechanisms (reviewed by Kupriyanova et al. 2001). Development of brooded serpulin larvae has been studied in less detail than planktotrophic larvae. Studies of development of non-feeding larvae (Fig. 12.4B) of Salmacina dysteri, Paraprotis dendrova, and Rhodopsis pusilla (Nishi and Yamasu 1992a, 1992b, 1992c) suggest that the developmental events and general larval morphology are very similar for brooded and planktonic serpulin larvae. The studies of lecithotrophic development inside the spirorbin brooding structures are fragmentary. Development from the early trochophore to swimming competent larvae is described for Pileolaria cf. militaris, Spirorbis sp., Circeis cf. armoricana, Neodexiospira alveolata, N. pseudocorrugata, Circeis cf. armoricana and Spirorbis spirorbis (see Kupriyanova et al. 2001 for details). Like serpulin trochophores, early spirorbin trochophores are subdivided by a prototroch into an episphere and a hyposphere. The prototroch of the early spirorbins consists of two bands of cilia. Eye spots may be present or absent at the early stage. A functional mouth and anus are absent. In metatrochophores (Fig. 12.4C), the collar forms ventrally under the prototroch; the eyes spots are present. In the late metatrochophore the mouth opens and branchial and opercular buds develop. A competent spirorbin larva released from the brooding chamber has three chaetigers and a terminal segment, three bands of cilia (prototroch, metatroch, and neurotroch), apical cilia, eyespots, branchial and opercular buds and a large collar. The stomach is not functional and is filled with yolk. In Siboglinidae, the larvae of all taxa described to date are lecithotrophic and many are brooded for some period (e.g., Crassibrachia,

## Reproductive Biology and Phylogeny of Annelida Oligobrachia, Nereilinum and Siboglinum), but others probably have freeswimming larvae (Southward 1993). In vestimentiferans, small, yolky, and slightly buoyant eggs develop into nonfeeding trochophore larvae. Marsh et al. (2001) found that the larvae of Riftia pachyptila can live for up to 38 days. The palps of siboglinids arise behind the larval prototroch, and hence are peristomial structures (Rouse and Fauchald 1997). Adult Siboglinidae have no mouth and the gut lumen is nearly completely occluded by the endoderm, though a small lumen does appear to be present (Southward 1982). According to Southward (1982) this was previously referred to as the medial coelomic cavity by Ivanov (1963). A transitory mouth or anus has been shown in Siboglinum poseidoni (Callsen-Cencic and Flügel 1995) and Ridgeia (Jones and Gardiner 1988; Southward 1988). This appears to be the pathway for bacteria to occupy the trophosome.

12.8 ASEXUAL REPRODUCTION Asexual reproduction is not known in Oweniidae or Sabellariidae, In Sabellidae it occurs via paratomy and has been reviewed by Knight-Jones and Bowden (1984). It only occurs in Sabellinae, with examples such as Sabella variabilis and taxa in Bispira and Branchiomma. Asexual reproduction via paratomy also occurs in Serpulinae and was reviewed in Kupriyanova et al. (2001). Asexual reproduction is best known for Filograna and Salmacina, but has also been described for Filogranula gracilis, Josephella marenzelleri, Rhodopsis pusilla, three species of Spiraserpula, and Filogranella elatensis. In asexually reproducing serpulins the parental animal divides into two by transverse fission in the middle of the abdomen. Before the separation takes place, the new cephalic region forms in the middle part of parental specimen by transformation of abdominal segments into thoracic ones (morphallaxis). Asexual reproduction typically leads to formation of “colonies” comprising a network of branching tubes. There has only been one report of asexual reproduction in Siboglinidae and this was for Sclerolinum brattstromi, which fragments readily (Southward 1975) and each fragment can regenerate anterior and posterior ends.

12.9 EVOLUTION OF REPRODUCTIVE MECHANISMS Little has been done on the evolution of reproductive mechanisms in annelids, but much of what work has been done is concentrated on Sabellida. Rouse and Fitzhugh (1994) assessed the evolution of various reproductive features in Sabellidae and examined the influence of body size on these characters. They suggest that the ancestral Sabellidae was gonochoric, had sperm with elongate heads and was a brooder of directdeveloping larvae. The general covariation of small body size with these reproductive traits suggested that small body size is also plesiomorphic for the family. Within Sabellinae, sperm with spherical nuclei and mitochondria, external fertilization and swimming larvae are secondarily derived. Brooding has subsequently re-appeared in apomorphic taxa such as

Sabellida

##

Amphiglena, Perkinsiana and Potamilla. A phylogenetic study by Kupriyanova (2003) suggests that benthic and non-feeding larvae are plesiomorphic in serpulid polychaetes, while planktonic and feeding larvae are apomorphic (Fig. 12.5). Both of these studies challenge the traditional view that external fertilization and planktotrophic larvae can be viewed as uniformly primitive traits.

12.10 ACKNOWLEDGEMENTS Thanks to Barrie Jamieson for conceiving this volume and for his comments on this chapter. This work was supported by the Australian Research Council, the South Australian Museum and the Kanagawa Academy of Science and Technology, Japan.

12.11 LITERATURE CITED Al-Ogily, S. M. and Knight-Jones, E. W. 1981. Circeis paguri, the spirorbid polychaete associated with the hermit-crab Eupagurus bernhardus. Journal of the Marine Biological Association of the United Kingdom 61: 821-826. Allen, M. J. 1957. The breeding of polychaetous annelids near Parguera, Puerto-Rico. Biological Bulletin 113: 49-57. Amieva, M. R. and Reed, C. G. 1987. Functional morphology of the larval tentacles of Phragmatopoma californica (Polychaeta: Sabellariidae): Composite larval and adult organs of multifunctional significance. Marine Biology 95: 243-258. Amieva, M. R., Reed, C. G. and Pawlik, J. R. 1987. Ultrastructure and behavior of the larva of Phragmatopoma californica (Polychaeta: Sabellariidae): Identification of sensory organs potentially involved in substrate selection. Marine Biology 95: 259-266. Andrews, J. C. and Anderson, D. T. 1962. The development and settling of the polychaete Galeolaria caespitosa Lamarck (Fam. Serpulidae). Proceedings of the Linnean Society of New South Wales 87: 185-188. Bailey-Brock, J. H. 1985. Polychaetes from Fijian coral reefs. Pacific Science 39: 195220. Bakke, T. 1983. Pogonophora. Pp. 377-385. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates, Volume II: Spermatogenesis and Sperm Function., John Wiley and Sons, Chichester. Bakke, T. 1990. Pogonophora. Pp. 37-48. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates. Volume IV, Part B Fertilization, Development, and Parental Care, John Wiley and Sons, Chichester. Banse, K. 1972. Redescription of some species of Chone Kröyer and Euchone Malmgren, and three new species (Sabellidae, Polychaeta). Fishery Bulletin. Fisheries and Wildlife Service. United States Department of Interior 70: 459-495. Bartolomaeus, T. 1995. Structure and formation of the uncini in Pectinaria koreni, Pectinaria auricoma (Terebellida) and Spirorbis spirorbis (Sabellida): implications for annelid phylogeny and the position of the Pogonophora. Zoomorphology 115: 161-177. Bartolomaeus, T. 1999. Structure, function and development of segmental organs in the Annelida. Hydrobiologia 402: 21-37. Beklemishev, V. N. 1944. Osnovy sravintel’noi anatomii bespozvonochnykh [Principles of Comparative Anatomy of Invertebrates]. Akademia Nauk, Moscow.

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Reproductive Biology and Phylogeny of Annelida

Ben-Eliahu, M. N. and ten Hove, H. A. 1989. Redescription of Rhodopsis pusilla Bush, a little known but widely distributed species of Serpulidae (Polychaeta). Zoologica Scripta 18: 381-395. Bhaud, M. R. 1969. Developpement larvaire de Phalacrostemma cidariophilus Marenzeller, 1895. Vie et Milieu 20: 543-558. Bhaud, M. R. 1975. Nouvelle observations de Sabellariidae (Annelides Polychaetes) dans la region malgache. Cahiers O.R.S.T.O.M. (Office de la Recherche Scientifique et Technique Outre-Mer) serie Océanographie 13: 69-78. Bhaud, M. R. and Fernandez-Alamo, M. A. 2001. First description of the larvae of Dianthyrsus (Sabellariidae, Polychaeta) from the Gulf of California and Bahia de Banderas, Mexico. Bulletin of Marine Science 68: 221-232. Callsen-Cencic, P. and Flügel, H. J. 1995. Larval development and the formation of the gut of Siboglinum poseidoni Flügel and Langhof (Pogonophora, Perviata). Evidence of protostomian affinity. Sarsia 80: 73-89. Canete, J. I. and Ambler, R. P. 1990. Growth and age determination in the spirorbid polychaete Romanchiella pustulata. Revista de Biologia Marina 25: 147-164. Carey, S. C., Felbeck, H. and Holland, N. D. 1989. Observations of the reproductive biology of the hydrothermal vent tube worm Riftia pachyptila. Marine Ecology Progress Series 52: 89-94. Carpizo-Ituarte, E. and Hadfield, M. G. 1998. Stimulation of metamorphosis in the polychaete Hydroides elegans Haswell (Serpulidae). Biological Bulletin 194: 14-24. Caullery, M. 1914. Sur les Siboglinidae, type nouveau d’invertébrés receuillis par l’expédition du Siboga. Bulletin de la Société Zoologique de France (Evolution et Zoologie) 39: 350-353. Caullery, M. and Mesnil, F. 1897. Études sur la morphologie comparée et la phylogenie des espèces chez les Spirorbes. Bulletin Scientifique de la France et de la Belgique 30: 185-233. Charles, F., Jordana, E., Amouroux, J.-M., Gremare, A., Desmalades, M. and Zudaire, L. 2003. Reproduction, recruitment and larval metamorphosis in the serpulid polychaete Ditrupa arietina (O.F. Muller). Estuarine Coastal and Shelf Science 57: 435-443. Chlebovitsch, V. V. 1959. Species of Polychaeta from the Kuril Islands, which are new or recorded for the first time in U.S.S.R. fauna. Zoologicheskii Zhurnal 38: 167181. (in Russian). Chughtai, I. 1986. Fine structure of spermatozoa in Perkinsiana rubra and Pseudopotamilla reniformis (Sabellidae: Polychaeta). Acta Zoologica 67: 165-171. Clark, R. B. and Olive, P. J. W. 1973. Recent advances in polychaete endocrinology and reproductive biology. Oceanography and Marine Biology: An Annual Review 11: 176-223. Colwin, A. L. and Colwin, L. H. 1961. Fine structure of the spermatozoon of Hydroides hexagonus (Annelida), with special reference to the acrosomal region. Journal of Biophysical and Biochemical Cytology 10: 211-230. Colwin, L. H. and Colwin, A. L. 1961a. Changes in spermatozoa during fertilization in Hydroides hexagonus (Annelida). I. Passage of the acrosomal region through the vitelline membrane. Journal of Biophysical and Biochemical Cytology 10: 231254. Colwin, L. H. and Colwin, A. L. 1961b. Changes in spermatozoa during fertilization in Hydroides hexagonus (Annelida). II. Incorporation with the egg. Journal of Biophysical and Biochemical Cytology 10: 255-274. Crisp, M. 1977. The development of the serpulid Pomatoleios kraussi (Annelida, Polychaeta). Journal of Zoology, London 183: 147-160.

Sabellida

##!

Curtis, M. 1977. Life cycles and population dynamics of marine benthic polychaetes from Disko Bay area of West Greenland. Ophelia 16: 9-58. Dales, R. P. 1952. The development and structure of the anterior region of the body in the Sabellariidae, with special reference to Phragmatopoma californica. Quarterly Journal of Microscopical Science 93: 435-452. Dales, R. P. 1961. The coelomic and peritoneal cell systems of some sabellid polychaetes. Quarterly Journal of Microscopical Science 102: 327-346. Dales, R. P. 1962. The polychaete stomodeum and the interrelationships of the families of the Polychaeta. Proceedings of the Zoological Society of London 139: 289-328. Daly, J. M. 1978a. The annual cycle and the short term periodicity of breeding in a Northumberland population of Spirorbis spirorbis (Polychaeta: Serpulidae). Journal of the Marine Biological Association of the United Kingdom 58: 161-176. Daly, J. M. 1978b. Growth and fecundity in a Northumberland population of Spirorbis spirorbis (Polychaeta: Serpulidae). Journal of the Marine Biological Association of the United Kingdom 58: 177-190. Daly, J. M. and Golding, D. W. 1977. A description of the spermatheca of Spirorbis spirorbis (L.) (Polychaeta: Serpulidae) and evidence for a novel mode of sperm transmission. Journal of the Marine Biological Association of the United Kingdom 57: 219-227. Dauvin, J.-C. and Gillet, P. 1991. Spatio-temporal variability in population structure of Owenia fusiformis Delle Chiaje (Annelida: Polychaeta) from the Bay of Seine (eastern English Channel). Journal of Experimental Marine Biology and Ecology 152: 105-122. Dean, D., Chapman, S. R. and Chapman, C. S. 1987. Reproduction and development of the sabellid polychaete Myxicola infundibulum. Journal of the Marine Biological Association of the United Kingdom 67: 431-439. Dixon, D. R. 1981. Reproductive biology of the serpulid Ficopomatus (Mercierella) enigmaticus in the Thames estuary, S. E. England. Journal of the Marine Biological Association of the United Kingdom 61: 805-815. Dons, C. 1933. Om vekst og forplantning hos Miroserpula inflata. Det Kongelige Norske Vidensabers Selskab Forhandlinger 6: 35-37. Dorresteijn, A. W. C. and Luetjens, C. M. 1994. Morphometric analysis of cellular specification in Platynereis and Pomatoceros embryogenesis (Annelida, Polychaeta). Mémoires du Museum National d‘Histoire Naturelle 162: 45-50. Dragesco-Kernéis, A. 1980. Phototaxie chez Dasychone lucullana (Delle Chiaje). Cahiers de Biologie Marine 21: 467-478. Eckelbarger, K. J. 1975. Developmental studies of the post-settling stages of Sabellaria vulgaris (Polychaeta: Sabellariidae). Marine Biology 30: 137-149. Eckelbarger, K. J. 1976. Larval development and population aspects of the reef- building polychaete Phragmatopoma lapidosa from the east coast of Florida. Bulletin of Marine Science 26: 117-132. Eckelbarger, K. J. 1977. Larval development of Sabellaria floridensis from Florida and Phragmatopomna californica from Southern California (Polychaeta: Sabellariidae), with a key to the sabellariid larvae of Florida and with a review of development in the family. Bulletin of Marine Science 27: 241-255. Eckelbarger, K. J. 1978. Metamorphosis and settlement in the Sabellariidae. Pp. 145164. In F.-S. Chia and M. E. Rice (eds), Settlement and Metamorphosis of Marine Invertebrate Larvae. Elsevier, New York. Eckelbarger, K. J. 1979. Ultrastructural evidence for both autosynthetic and heterosynthetic yolk formation in the oocytes of an annelid (Phragmatopoma lapidosa: Polychaeta). Tissue and Cell 11: 425-443.

##" Reproductive Biology and Phylogeny of Annelida Eckelbarger, K. J. 1984. Ultrastructure of spermatogenesis in the reef-building polychaete Phragmatopoma lapidosa (Sabellariidae) with special reference to acrosome morphogenesis. Journal of Ultrastructure Research 89: 146-164. Emlet, R. B. and Strathmann, R. R. 1994. Functional consequences of simple cilia in the mitraria of oweniids (an anomalous larvae of an anomalous polychaete) and comparisons with other larvae. Pp. 143-157. In W. H. Wilson, S. A. Stricker and G. L. Shin (eds), Reproduction and Development of Marine Invertebrates. Johns Hopkins University Press, Baltimore. Fauchald, K. 1977. The polychaete worms. Definitions and keys to the orders, families and genera. Natural History Museum of Los Angeles County. Science Series 28: 1-188. Fauchald, K. 1983. Life diagram patterns in benthic polychaetes. Proceedings of the Biological Society of Washington 96: 160-177. Faulkner, G. H. 1929. The anatomy and the histology of bud-formation in the serpulid Filograna implexa together with some cytological observations on the nuclei of the neoblasts. Journal of the Linnean Society (Zoology) 37: 109-190. Fitzhugh, K. 1989. A systematic revision of the Sabellidae-CaobangiidaeSabellongidae complex (Annelida: Polychaeta). Bulletin of the American Museum of Natural History 192: 1-104. Fitzhugh, K. and Rouse, G. W. 1999. A remarkable new genus and species of fan worm (Polychaeta: Sabellidae: Sabellinae) associated with marine gastropods. Invertebrate Biology 118: 357-390. Franzén, A. 1956. On spermatogenesis, morphology of spermatozoan, and biology of fertilization among invertebrates. Zoologiska Bidrag från Uppsala 31: 355-482. Franzén, A. 1958. On sperm morphology and acrosome filament formation in some Annellida, Echiuroidea, and Tunicata. Zoologiska Bidrag från Uppsala 33: 1-28. Franzén, A. 1982. Ultrastructure of spermatids and spermatozoa in three polychaetes with modified biology of reproduction: Autolytus sp., Chitinopoma serrula and Capitella capitata. International Journal of Invertebrate Reproduction 5: 185-200. Franzén, Å. 1973. The spermatozoon of Siboglinum. Acta Zoologica 54: 179-192. Franzén, Å. and Rice, S. A. 1988. Spermatogenesis, male gametes and gamete interaction. Pp. 309-333. In W. Westheide and C. O. Hermans (eds), The Ultrastructure of Polychaeta, Microfauna Marina 4, Gustav Fischer Verlag, Stuttgart. Føyn, B. and Gjøen, I. 1954. Studies on the serpulid Pomatoceros triqueter (L.) I. Observations on the life history. Nytt Magasin for Zoologi 2: 73-81. Gaikwad, U. D. 1988. Artificial fertilization in serpulid worm Spirobranchus tetraceros (Schmarda) (Polychaeta, Annelida) and some larval stages. Pp. 280-284. In S. Palanichamy (ed.), Recent Advances in Invertebrate Reproduction and Aquaculture, Arulmugi Palaniandavar College of Art and Culture, Palani, India. Gambi, M. C., Giangrande, A. and Patti, F. P. 2000. Comparative observations on reproductive biology of four species of Perkinsiana (Polychaeta: Sabellidae: Sabellinae). Bulletin of Marine Science 67: 299-309. Gambi, M. C. and Patti, F. P. 1999. Reproductive biology of Perkinsiana antarctica (Kinberg) (Polychaeta, Sabellidae) in the Straits of Magellan (South America): Systematic and ecological implications. Scientia Marina 63: 253-259. Gardiner, S. L. and Jones, M. L. 1985. Ultrastructure of spermiogenesis in the vestimentiferan tube worm Riftia pachyptila (Pogonophora: Obturata). Transactions of the American Microscopical Society 104: 19-44. Gardiner, S. L. and Jones, M. L. 1993. Vestimentifera. Pp. 371-460. In F. W. Harrison and M. E. Rice (eds), Microscopic Anatomy of Invertebrates, Volume 12: Onychophora, Chilopoda and Lesser Protostomata. Wiley-Liss, New York.

Sabellida

###

Gee, J. M. 1964. The British Spirorbinae with a description of S. cuneatus sp. n. and a review of the genus Spirorbis. Proceedings of the Zoological Society of London 143: 405-441. Gee, J. M. 1967. Growth and breeding of Spirorbis rupestris (Polychaeta: Serpulidae). Journal of Zoology, London 152: 235-244. Gee, J. M. and Knight-Jones, E. W. 1962. The morphology and larval behaviour of a new species of Spirorbis (Serpulidae). Journal of the Marine Biological Association of the United Kingdom 42: 641-654. Gee, J. M. and Williams, G. B. B. 1965. Self- and cross ferilization in Spirorbis borealis and S. pagenstecheri. Journal of the Marine Biological Association of the United Kingdom 45: 275-285. Gentil, F., Dauvin, J.-C. and Ménard, F. 1990. Reproductive biology of the polychaete Owenia fusiformis Delle Chiaje in the Bay of Seine (eastern English Channel). Journal of Experimental Marine Biology and Ecology 142: 13-23. Giangrande, A. 1997. Polychaete reproduction patterns, life-cycles and life-histories: an overview. Oceanography and Marine Biology: An Annual Review 35: 323-386. Giangrande, A., Licciano, M., Pagliara, P. and Gambi, M. C. 2000. Gametogenesis and larval development in Sabella spallanzanii (Polychaeta: Sabellidae) from the Mediterranean Sea. Marine Biology 136: 847-861. Gilson, G. 1895. The nephridial duct of Owenia. Anatomischer Anzeiger 10: 191-194. Grant, N. J. 1981. A scanning electron microscopic study of larva; development in the marine polychaete, Galeolaria caespitosa. Cell and Tissue Research 215: 171-179. Gravier, C. 1909. Contributions a l’étude de la morphologie et de l’évolution des Sabellariens St-Joseph [Hermelliens (Quatrefages)]. Annales des Sciences Naturelles (Zoologie comprenant L’anatomie, La physiologie, La classification et L’histoire naturelle des animaux) (Sér. 9) 9: 287-304. Harris, T. 1968. Spirorbis species (Polychaeta: Serpulidae) from the Isles of Scilly, including descriptions of two new species. Journal of the Marine Biological Association of the United Kingdom 48: 593-602. Haswell, W. A. 1884. The marine annelids of the order Serpulea. Some observations on their anatomy, with the characteristics of the Australian species. Proceedings of the Linnean Society of New South Wales 9: 649-675. Hess, H. C. 1993. The evolution of parental care in brooding spirorbid polychaetes: The effect of scaling constraints. American Naturalist 141: 577-596. Hill, M. B. 1967. The life cycles and salinity tolerance of the serpulid Mercierella enigmatica (Fauvel) and Hydroides uncinata (Philippi) at Lagos, Nigeria. Journal of Animal Ecology 36: 302-332. Hilário, A., Young, C. M. and Tyler, P. A. 2005. Sperm storage, internal fertilization, and embryonic dispersal in vent and seep tubeworms (Polychaeta: Siboglinidae: Vestimentifera). Biological Bulletin 208: 20-28. Hsieh, H. L. 1995. Laonome albicingillum, a new fan worm species (Polychaeta, Sabellidae, Sabellinae) from Taiwan. Proceedings of the Biological Society of Washington 108: 130-135. Hsieh, H. L. 1997. Self-fertilization — a potential fertilization mode in an estuarine sabellid polychaete. Marine Ecology Progress Series 147: 143-148. Ivanov, A. V. 1963. Pogonophora. Academic Press, London, Ivanov, A. V. 1994. On the systematic position of Vestimentifera. Zoologische Jahrbücher Abteilung für Systematik, Geographie, und Biologie der Tiere 121: 409-456. Jamieson, B. G. M. and Rouse, G. W. 1989. The spermatozoa of the Polychaeta (Annelida): An ultrastructural review. Biological Reviews 64: 93-157.

##$ Reproductive Biology and Phylogeny of Annelida Johansson, K. E. 1939. Lamellisabella zachsi Uschakow, ein Vertreter eine neuen Tierklasse Pogonophora. Zoologiska Bidrag från Uppsala 18: 253-268. Jones, M. L. 1974. On the Caobangiidae, a new family of the Polychaeta, with a redescription of Caobangia billeti Girard. Smithsonian Contributions to Zoology 175: 1-55. Jones, M. L. 1985. On the Vestimentifera, new phylum: Six new species, and other taxa, from hydrothermal vents and elsewhere. Bulletin of the Biological Society of Washington 6: 117-158. Jones, M. L. and Gardiner, S. L. 1988. Evidence for a transient digestive tract in Vestimentifera. Proceedings of the Biological Society of Washington 101: 423-433. Jyssum, S. 1957. Investigations of the neoblasts and oogenesis in the serpulid, Pomatoceros triqueter L. Nytt Magasin for Zoologi 5: 5-10. Kerby, C. J. 1972. The Biology of Sabella microphthalma (Polychaeta). Ph.D. Dissertation, George Washington University. King, P. E., Bailey, J. H. and Babbage, P. C. 1969. Vitellogenesis and formation of the egg chain in Spirorbis borealis (Serpulidae). Journal of the Marine Biological Association of the United Kingdom 49: 141-150. Kirtley, D. W. 1994. A Review and Taxonomic Revision of the Family Sabellariidae Johnston, 1865 (Annelida; Polychaeta), Sabecon Press, Vero Beach, FL, 223 pp. Knight-Jones, E. W. 1951. Gregariousness and some other aspects of the setting behaviour of Spirorbis. Journal of the Marine Biological Association of the United Kingdom 30: 201-222. Knight-Jones, E. W., Knight-Jones, P. and Bregazzi, P. K. 1973. Helicosiphon biscoensis Gravier (Polychaeta: Serpulidae) and its relationship with other Spirorbinae. Zoological Journal of the Linnean Society, London 52: 9-22. Knight-Jones, E. W., Knight-Jones, P. and Llewellyn, L. C. 1974. Spirorbinae (Polychaeta: Serpulidae) from southeastern Australia. Notes on their taxonomy, ecology and distribution. Records of the Australian Museum 29: 107-151. Knight-Jones, P. 1972. New species and a new subgenus of Spirorbinae (Serpulidae: Polychaeta) from Kenya. Journal of Zoology, London 166: 1-18. Knight-Jones, P. 1973. Spirorbinae (Serpulidae: Polychaeta) from southeastern Australia. Pt. 1. A new genus, four new subgenera and seven new species. Bulletin of the British Museum (Natural History) 24: 229-259. Knight-Jones, P. 1978. New Spirorbidae (Polychaeta: Sedentaria) from the east Pacific, Atlantic, Indian and southern oceans. Zoological Journal of the Linnean Society 64: 201-240. Knight-Jones, P. 1981. Behaviour, setal inversion and phylogeny of Sabellida (Polychaeta). Zoologica Scripta 10: 183-202. Knight-Jones, P. 1984. A new species of Protoleodora (Spirorbidae: Polychaeta) from eastern U.S.S.R., with a brief revision of related genera. Zoological Journal of the Linnean Society 80: 109-120. Knight-Jones, P. and Bowden, N. 1984. Incubation and scissiparity in Sabellidae (Polychaeta). Journal of the Marine Biological Association of the United Kingdom 64: 809-818. Knight-Jones, P. and Knight-Jones, E. W. 1977. Taxonomy and ecology of British Spirorbidae (Polychaeta). Journal of the Marine Biological Association of the United Kingdom 57: 453-500. Knight-Jones, P. and Knight-Jones, E. W. 1994. Spirorbidae (Polychaeta) from Signy Island, South Orkneys, including 3 new species. Ophelia 40: 75-94. Knight-Jones, P. and Knight-Jones, E. W. 1995. Spirorbidae (Polychaeta) from Madeira including a new species and subgenus of Spirorbis. Mitteilungen aus dem Hamburgischen Zoologischen Museum und Institut 92: 89-101.

Sabellida

##%

Knight-Jones, P., Knight-Jones, E. W. and Kawahara, T. 1975. A review of the genus Janua, including Dexiospira (Polychaeta: Spirorbinae). Zoological Journal of the Linnean Society 56: 91-129. Knight-Jones, P. and Walker, A. J. M. 1972. Spirorbinae (Serpulidae: Polychaeta) on limpets from the South Orkney Islands. Bulletin of British Antarctic Survey 31: 33-40. Kopp, J. C. 1985. A preliminary ultrastructural study of Phragmatopoma (Polychaeta) gametes. Invertebrate Reproduction and Development 8: 297-302. Kryvi, H. and Graebner, I. 1975. Acrosome formation and the centriolar complex in the spermatozoa of Sabella penicillum (Polychaeta). An electron microscopical study. Cell and Tissue Research 161: 47-53. Kupriyanova, E. K. 2003. Life history evolution in Serpulimorph polychaetes: a phylogenetic analysis. Hydrobiologia 496: 105-114. Kupriyanova, E. K., Nishi, E., ten Hove, H. A. and Rzhavsky, A. V. 2001. A review of life history patterns in serpulimorph polychaetes: ecological and evolutionary perspectives. Oceanography and Marine Biology: An Annual Review 39: 1-101. Kupriyanova, E. K. and Havenhand, J. N. 2002. Variation in sperm swimming behaviour and its effect on fertilization success in the serpulid polychaete Galeolaria caespitosa. Invertebrate Reproduction and Development 41: 1-6. L’Hardy, J.-P. and Quiévreux, C. 1964. Observations sur Spirorbis (Laeospira) inornatus (Polychète Serpulidae) et la systematique des Spirorbinae. Cahiers de Biologie Marine 5: 287-294. Lacalli, T. 1976. Remarks on the larvae of two serpulids. Canadian Journal of Zoology 55: 300-303. Liwanow, N. A. and Porfirjewa, N. A. 1967. Die Organisation der Pogonophoren und deren Beziehungen zu den Polychäten. Biologische Zentralblatt 86: 177-204. Marsh, A. G., Mullineaux, L. S., Young, C. M. and Manahan, D. T. 2001. Larval dispersal potential of the tubeworm Riftia pachyptila at deep-sea hydrothermal vents. Nature 411: 77-80. Matjasic, J. and Sket, B. 1966. Développement larvaire du serpulien cavernicole Marifugia cavatica Absalon et Hrabe (Polychaeta, Sedentaria). International Journal of Speleology 2: 9-16. Matsuo, R. and Yoshioshi, K. 1983. A scanning electron microscopical observation on the spermatozoa of three species of Hydroides (Polychaeta, Serpulidae). Marine Fouling 4: 23-25. Matuso, R. and Ko, Y. 1981. Preliminary notes on the development, growth and maturation of laboratory reared three species of Hydroides (Annelida: Polychaeta). Bulletin of the Faculty of Fisheries, Nagasaki University 51: 23-28. Mauro, N. A. 1975. The premetamorphic developmental rate of Phragmatopoma lapidosa Kinberg, 1867, compared with that in temperate sabellariids (Polychaeta: Sabellariidae). Bulletin of Marine Science 25: 387-392. McEuen, F. S., Wu, B. L. and Chia, F. S. 1983. Reproduction and development of Sabella media, a polychaete with extratubular brooding. Marine Biology 76: 301-309. McHugh, D. 1997. Molecular evidence that echiurans and pogonophorans are derived annelids. Proceedings of the National Academy of Sciences of the United States of America 94: 8006-8009. McIntosh, W. C. 1917. Notes from the Gatty Marine Laboratory, St. Andrews. No. 40. On the nervous system and other points in the structure of Owenia and Myriochele. Annals and Magazine of Natural History 19: 233-265. Meyer, K. and Bartolomaeus, T. 1996. Ultrastructure and formation of the hooked setae in Owenia fusiformis delle Chiaje, 1842 — implications for annelid phylogeny. Canadian Journal of Zoology 74: 2143-2153.

##& Reproductive Biology and Phylogeny of Annelida Minichev, Y. S. and Bubko, O. V. 1992. [Are the Oweniidae polychaetes?]. Explorations of the Fauna of the Seas 43: 47-51. (In Russian) Miura, T. and Kajihara, T. 1981. The development of a serpulid worm, Hydroides ezoensis (Annelida, Polychaeta). Proceedings of the Japanese Society of Systematic Zoology 20: 7-12. Mona, M. H., Eissa, S. H. H., Abdel-Gawad, A. M. and Barbary, M. S. 1994. Ultrastructural investigation of spermatogenesis in the tubeworm Hydroides dirampha (Polychaeta, Serpulidae). Journal of the Egyptian German Society of Zoology 13: 115-128. Morris, R. H., Abbot, D. P. and Haderlie, E. C. 1980. Intertidal Invertebrates of California, Stanford University Press, Stanford. Ménard, F., Gentil, F. and Dauvin, J.-C. 1989. Population dynamics and secondary production of Owenia fusiformis Delle Chiaje. Polychaeta from the Bay of Seine eastern English Channel France. Journal of Experimental Marine Biology and Ecology 133: 151-168. Nelson-Smith, A. 1971. Annelids as fouling organisms. Pp. 171-184. In E. B. Gareth Jones and S. K. Eltingham (eds), Marine Borers, Fungi and Fouling Organisms, Organisation for Economic Co-operation and Development, Paris. Nishi, E. 1992. Occurrence of the boring serpulid Floriprotis sabiuraensis Uchida and the brooding serpulid Paraproti dendrova Uchida (Polychaeta: Sedentaria) in Okinawa. Galaxea 11: 15-20. Nishi, E. 1993. Notes of reproductive biology of some serpulids polychaetes at Sesoko Island, Okinawa, with brief accounts of setal morphology of three species of Salmacina and Filograna implexa. Marine Fouling 10: 11-16. Nishi, E. and Nishihira, M. 1993. Hermaphroditism, brooding and gamete production in the serpulid polychaete Salmacina dysteri. Publications from the Amakusa Marine Biological Laboratory, Kyushu 12: 1-11. Nishi, E. and Yamasu, T. 1992a. Brooding and development of Rhodopsis pusilla Bush (Sedentaria, Polychaeta). Bulletin of the College of Science, University of the Ryukyus 54: 93-100. Nishi, E. and Yamasu, T. 1992b. Brooding and larval development of a serpulid tube worm Salmacina dysteri (Huxley) (Sedentaria; Polychaeta). Bulletin of the College of Science, University of the Ryukyus 54: 107-121. Nishi, E. and Yamasu, T. 1992c. Brooding habit and development of a serpulid worm Paraprotis dendrova Uchida (Annelida, Polychaeta, Sedentaria). Bulletin of the College of Science, University of the Ryukyus 54: 83-92. Nishi, E. and Yamasu, T. 1992d. Observation on the reproductive behaviour and the development of a common fouling spirorbid, Dexiospira foraminosa Bush (Sedentaria, Polychaeta). Bulletin of the College of Science, University of the Ryukyus 54: 101-106. Nordback, K. 1956. On the oogenesis and fertilization of the serpulid Hydroides norvegica (Gunnerus). Nytt Magasin for Zoologi 4: 121-123. Okuda, S. 1946. Studies on the development of Annelida Polychaeta I. Journal of the Faculty of Science, Hokkaido Imperial University, Ser. 6, Zoology 9: 115-219. Pasteels, J. 1965. La fécondation étudiée au microscope électronique étude comparative. Bulletin de la Société Zoologique de France (Evolution et Zoologie) 90: 195224. Pawlik, J. R. and Mense, D. J. 1994. Larval transport, food limitation, ontogenetic plasticity, and the recruitment of sabellariid polychaetes. Pp. 275-286. In W. H. Wilson, S. A. Stricker and G. L. Shinn (eds), Reproduction and Development of Marine Invertebrates. Johns Hopkins University Press, Baltimore.

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Pechenik, J. A. and Qian, P.-Y. 1998. Onset and maintenance of metamorphic competence in the marine polychaete Hydroides elegans Haswell in response to three chemical cues. Journal of Experimental Marine Biology and Ecology 226: 51-74. Pettibone, M. H. 1982. Annelida. Pp. 1-43. In S. P. Parker (ed.), Synopsis and Classification of Living Organisms, vol. 2. McGraw-Hill Book Co, New York. Picard, A. 1980. Spermatogenesis and sperm-spermatheca relations in Spirorbis spirorbis (L.). International Journal of Invertebrate Reproduction 2: 73-83. Potswald, H. 1967. Observations on the genital segments of Spirorbis (Polychaeta). Biological Bulletin 132: 91-107. Potswald, H. E. 1967a. An electron microscope study of spermiogenesis in Spirorbis (Laeospira) moerchi Levinsen (Polychaeta). Zeitschrift für Zellforschung und Mikroskopische Anatomie 83: 231-248. Potswald, H. E. 1967b. Observations on the genital segments of Spirorbis (Polychaeta). Biological Bulletin 132: 91-107. Potswald, H. E. 1969. Cytological observations on the so-called neoblasts in the serpulid Spirorbis. Journal of Morphology 128: 241-259. Quievreux, C. 1962. Morphologie et anatomie des larves de Spirorbis vitreus (Fabricius) et Spirorbis malardi (Caullery et Mesnil) (Annelides Polychetes). Cahiers de Biologie Marine 3: 1-12. Rioja, E. 1923. Estudio sistemático de las especies Ibéricas del suborden Sabelliformia. Trabajos del Museo Nacional de Ciencias Naturales Serie Zoológica 48: 1-144. Rouse, G. W. 1988. An ultrastructural study of the spermatozoa of Eulalia sp. (Phyllodocidae), Lepidonotus (Polynoidae), Lumbrinereis sp. (Lumbrinereidae) and Owenia fusiformis (Oweniidae). Helgoländer Meeresuntersuchungen 42: 67-78. Rouse, G. W. 1992a. Ultrastructure of spermiogenesis and spermatozoa of four Oriopsis species (Sabellinae; Sabellidae; Polychaeta). Zoologica Scripta 21: 363379. Rouse, G. W. 1992b. Ultrastructure of the spermathecae of Parafabricia ventricingulata and three species of Oriopsis (Polychaeta: Sabellidae). Acta Zoologica 73: 141-151. Rouse, G. W. 1995a. Is sperm ultrastructure useful in polychaete systematics? An example using 20 species of the Fabriciinae (Sabellidae, Polychaeta). Acta Zoologica 76: 57-74. Rouse, G. W. 1995b. Spermathecae of Fabricia and Manayunkia (Sabellidae, Polychaeta). Invertebrate Biology 114: 248-255. Rouse, G. W. 1996a. New Fabriciola and Manayunkia species (Fabriciinae, Sabellidae, Polychaeta) from Papua New Guinea. Journal of Natural History 30: 1761-1778. Rouse, G. W. 1996b. A new species of Perkinsiana (Sabellidae, Polychaeta) from Papua New Guinea; with a description of larval development. Ophelia 45: 101-114. Rouse, G. W. 1996c. Variability of sperm storage by females in the Sabellidae and Serpulidae (Polychaeta). Zoomorphology 116: 179-193. Rouse, G. W. 1999a. Polychaeta, including Pogonophora and Myzostomida. Pp. 81124. In B. G. M. Jamieson (ed.), Reproductive Biology of Invertebrates, Volume IXB. Progress in Male Gamete Ultrastructure and Phylogeny. Oxford and IBH Publishing Co., New Delhi. Rouse, G. W. 1999b. Polychaete sperm: phylogenetic and functional considerations. Hydrobiologia 402: 215-224. Rouse, G. W. 1999c. Trochophore concepts: ciliary bands and the evolution of larvae in spiralian Metazoa. Biological Journal of the Linnean Society 66: 411-464. Rouse, G. W. 2000. Bias? What bias? Gain and loss of downstream larval-feeding in animals. Zoologica Scripta 29: 213-236.

#$ Reproductive Biology and Phylogeny of Annelida Rouse, G. W. 2001. A cladistic analysis of Siboglinidae Caullery, 1914 (Polychaeta, Annelida): formerly the phyla Pogonophora and Vestimentifera. Zoological Journal of the Linnean Society 132: 55-80. Rouse, G. W. 2005. Annelid sperm and fertilization biology. Hydrobiologia 535: 167178. Rouse, G. W. and Fauchald, K. 1995. The articulation of annelids. Zoologica Scripta 24: 269-301. Rouse, G. W. and Fauchald, K. 1997. Cladistics and polychaetes. Zoologica Scripta 26: 139-204. Rouse, G. W. and Fitzhugh, K. 1994. Broadcasting fables: Is external fertilization really primitive? Sex, size and larvae in sabellid polychaetes. Zoologica Scripta 23: 271-312. Rouse, G. W. and Gambi, M. C. 1997. Cladistic relationships within Amphiglena Claparède (Polychaeta: Sabellidae) with a new species and a redescription of A. mediterranea (Leydig). Journal of Natural History 31: 999-1018. Rouse, G. W. and Gambi, M. C. 1998a. Evolution of reproductive features and larval development in the genus Amphiglena Claparède (Polychaeta: Sabellidae). Marine Biology 131: 743-753. Rouse, G. W. and Gambi, M. C. 1998b. Sperm ultrastructure and spermathecal structure in Amphiglena spp. (Polychaeta: Sabellidae). Invertebrate Biology 117: 114-122. Rouse, G. W., Goffredi, S. K. and Vrijenhoek, R. C. 2004. Osedax: Bone-eating marine worms with dwarf males. Science 305: 668-671. Rouse, G. W. and Pleijel, F. 2001. Polychaetes. Oxford University Press, London, 354 pp. Rousset, V., Rouse, G. W., Siddall, M. E., Tillier, A. and Pleijel, F. 2004. The phylogenetic position of Siboglinidae (Annelida), inferred from 18S rRNA, 28S rRNA, and morphological data. Zoologica Scripta 20: 518-533. Rullier, F. 1960. Développement de Salmacina dysteri (Huxley). Cahiers de Biologie Marine 1: 37-46. Rzhavsky, A. V. 1998. Circeis oshurkovi sp.n. (Polychaeta, Spirorbidae) from the North Pacific. Ophelia 48: 207-210. Rzhavsky, A. V. and Britayev, T. A. 1984. The ecology of Janua (Dexiospira) nipponica and J. (D.) alveolata (Polychaeta, Spirorbidae) near the southern shore of the Soviet Far East and the morphology of their tubes. Zoologichesky Zhurnal 63: 1305-1316 (In Russian). Rzhavsky, A. V. and Britayev, T. A. 1988. Specific features of populations of Circeis armoricana on hermit crabs on the East Kamchatka coast. Zoologichesky Zhurnal 67: 17-22. (In Russian). Salensky, W. 1882. Études sur le développement des annélides. Pt. I. 1. Psygmobranchus protensus. Archives de biologie, Liège 3: 345-378. Sawada, N. 1984. Electron microscopical studies of spermatogenesis in polychaetes. Fortschritte der Zoologie 29: 99-114. Scheltema, R. S., Williams, I. P., Shaw, M. A. and Loudon, C. 1981. Gregarious settlement by the larvae of Hydroides dianthus (Polychaeta: Serpulidae). Marine Ecology Progress Series 5: 69-74. Segrove, F. 1941. The development of the serpulid Pomatoceros triqueter L. Quarterly Journal of Microscopical Science 82: 467-540. Silva, de, P. D. H. 1962. Experiments on choice of substrata by Spirorbis larvae (Serpulidae). Journal of Experimental Biology 39: 483-490. Silva, de, P. D. H. and Knight-Jones, E. W. 1962. Spirorbis corallinae n.sp. and some other Spirorbinae (Serpulidae) common on British shores. Journal of the Marine Biological Association of the United Kingdom 42: 601-608.

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Simon, C. A. and Rouse, G. W. 2005. Sperm ultrastructure, spermiogenesis and spermathecal structure in Terebrasabella heterouncinata (Polychaeta: Sabellidae: Sabellinae). Invertebrate Biology 125: 39-49. Smith, P. R. and Chia, F. S. 1985. Larval development and metamorphosis of Sabellaria cementarium Moore, 1906 (Polychaeta: Sabellariidae). Canadian Journal of Zoology 63: 1037-1049. Smith, R. S. 1984. Development and settling of Spirobranchus giganteus (Polychaeta: Serpulidae). Pp. 461-483. In P. A. Hutchings (ed.), First International Polychaete Conference. Linnean Society of New South Wales, Sydney. Smith, R. S. 1991. Relationships within the order Sabellida (Polychaeta). Ophelia Supplement 5: 249-260. Sordino, P. and Gambi, M. C. 1992. Prime osservazioni sulla biologia riproduttiva e sul ciclo vitale di Branchiomma luctuosum (Grube 1869) (Polychaeta Sabellidae). Oebalia Supplement 17: 425-427. Sordino, P. and Gambi, M. C. 1994. Reproductive biology and life cycle of Branchiomma luctuosum (Grube 1869) (Polychaeta Sabellidae) in the Mediterranean Sea. Mémoires du Muséum National d’Histoire Naturelle 162: 640. Southward, E. C. 1975. Pogonophora. Pp. 129-156. In A. C. Giese and J. S. Pearse (eds), Reproduction of Marine Invertebrates, vol. II: Entoprocts and Lesser Coelomates, Academic Press, California. Southward, E. C. 1982. Bacterial symbionts in Pogonophora. Journal of the Marine Biological Association of the United Kingdom 62: 889-906. Southward, E. C. 1988. Development of the gut and segmentation of newly settled stages of Ridgeia (Vestimentifera): implications for relationship between Vestimentifera and Pogonophora. Journal of the Marine Biological Association of the United Kingdom 68: 465-487. Southward, E. C. 1993. Pogonophora. Pp. 327-369. In F. W. Harrison and M. E. Rice (eds), Microscopic Anatomy of Invertebrates, Volume 12, Onychophora, Chilopoda and Lesser Protostomata., Wiley-Liss, New York. Southward, E. C. 2000. Pogonophora. Pp. 331-351. In P. Beesely, G. J. B. Ross and C. J. Glasby (eds), Polychaeta and Allies: The Southern Synthesis. Fauna of Australia, Volume 4A. Polychaeta, Myzostomida, Pogonophora, Echiura, Sipuncula, CSIRO Publishing, Melbourne. Southward, E. C. and Coates, K. A. 1989. Sperm masses and sperm transfer in a Vestimentiferan, Ridgeia piscesae Jones 1985 (Pogonophora Obturata). Canadian Journal of Zoology 67: 2776-2781. Strathmann, M. F. 1987. Phylum Annelida, Class Polychaeta. Pp. 138-195. Reproduction and Development of Marine Invertebrates of Northern Pacific Coast. Data and Methods for the Study of Eggs, Embryos, and Larvae. University of Washington Press, Seattle. Strathmann, R. R., Jahn, T. L. and Fonseca, J. R. C. 1972. Suspension feeding by marine invertebrate larvae: clearance of particles by ciliated bands of a rotifer, pluteus, and trochophore. Biological Bulletin 142: 505-519. Tampi, P. R. S. 1960. On the early development of Protula tubularia (Montagu). Journal of the Marine Biological Association of India 2: 53-56. Ten Hove, H. A. 1984. Towards a phylogeny in serpulids (Annelida; Polychaeta), pp. 181196. In P. A. Hutchings (ed.), First International Polychaete Conference. Linnean Society of New South Wales, Sydney. Thiébaut, E. and Dauvin, J.-C. 1992. Développement morphologique et croissance des juvéniles de l’Owenia fusiformis Delle Chiaje (Polychaeta, Oweniidae). Canadian Journal of Zoology 70: 1701-1711.

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Thomas, F. I. M. 1994. Transport and mixing of gametes in three free-spawning annelids, Phragmatopoma californica (Fewkes), Sabellaria cementarium (Moore), and Schizobranchia insignis (Bush). Journal of Experimental Marine Biology and Ecology 179: 11-27. Thorson, G. 1946. Reproduction and larval development of Danish marine bottom invertebrates with special reference to the planktonic larvae in the Sound (Øresund). Meddelelser fra Kommissionen for Danmarks Fiskeri- Og Havundersøgelser, Serie: Plankton 4: 1-523. Uchida, H. 1978. Serpulid tube worms (Polychaeta, Sedentaria) from Japan with the systematic review of the group. Bulletin of the Marine Park Research Stations 2: 1-98. Van Dover, C. L. 1994. In situ spawning of hydrothermal vent tubeworms (Riftia pachyptila). Biological Bulletin. Marine Biological Laboratory, Woods Hole, Mass. 186: 134-135. Vannini, E. 1950. Studi sulla sessualita e sui poteri riginerativi nel polichete ermafrodita Salmacina incrustans Clap. 1.— Osservazioni sul ciclo riproduttivo sessuale e asessuale. Pubblicazioni della Stazione Zoologica di Napoli 22: 211-256. Vine, P. J. 1977. The marine fauna of New Zealand. Spirorbinae (Polychaeta: Serpulidae). Memoirs of the New Zealand Oceanographic Institute 68: 1-66. Watson, A. T. 1901. On the structure and habits of the Polychaeta of the family Ammocharidae. Journal of the Linnean Society of London (Zoology) 28: 230-260. Wilson, D. P. 1929. The larvae of the British sabellarians. Journal of the Marine Biological Association of the United Kingdom 16: 221-268. Wilson, D. P. 1932. On the Mitraria larva of Owenia fusiformis Delle Chiaje. Philosophical Transactions of the Royal Society of London. Series B 221: 231-334. Wilson, D. P. 1936. The development of the sabellid Branchiomma vesiculosum. Quarterly Journal of Microscopical Science 78: 534-603. Wilson, D. P. 1968a. The settlement behaviour of the larvae of Sabellaria alveolata (L.). Journal of the Marine Biological Association of the United Kingdom 48: 387-435. Wilson, D. P. 1968b. Some aspects of the development of eggs and larvae of Sabellaria alveolata. Journal of the Marine Biological Association of the United Kingdom 48: 367-386. Wilson, D. P. 1970. Additional observations on the larval growth and settlement of Sabellaria alveolata. Journal of the Marine Biological Association of the United Kingdom 50: 1-31. Wilson, D. P. 1970. The larvae of Sabellaria spinulosa and their settlement behaviour. Journal of the Marine Biological Association of the United Kingdom 50: 33-52. Wilson, D. P. 1977. The distribution, development and settlement of the sabellarian polychaete L. muratus (Allen) near Plymouth. Journal of the Marine Biological Association of the United Kingdom 57: 761-792. Wilson, W. H. 1991. Sexual reproductive modes in polychaetes: classification and diversity. Bulletin of Marine Science 48: 500-516. Young, C. M. and Chia, F.-S. 1982. Ontogeny of phototaxis during larval development of the sedentary polychaete, Serpula vermicularis (L.). Biological Bulletin 162: 457-468. Young, C. M., Vásquez, E., Metaxas, A. and Tyler, P. A. 1996. Embryology of vestimentiferan tube worms from deep-sea methane/sulphide seeps. Nature 381: 514-516. Yun, S. G. and Kikuchi, T. 1991. Larval development and settlement of Chone duneri Malmgren (Polychaeta: Sabellidae). Publications from the Amakusa Marine Biological Laboratory, Kyushu 11: 31-42.

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Yun, S. G. and Kikuchi, T. 1991. Reproduction of Chone duneri Malmgren (Polychaeta: Sabellidae). Publications from the Amakusa Marine Biological Laboratory, Kyushu 11: 19-30. Zenkevitsch, L. A. 1925. Biologie, Anatomie und Systematik der Süsswasserpolychaeten des Baikalsees. Zoologisches Jahrbücher Abteilung für Systematik, Geographie, und Biologie der Tiere 50: 1-60. Zibrowius, H. 1983. Chitinopoma arndti n.sp., an incubating bathyal serpulid polychaete from Saint-Paul Island, southern Indian Ocean. Tethys 11: 21-24.

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13

CHAPTER

Spionida James A. Blake

13.1 INTRODUCTION Spioniform polychaetes are sedentary, tubicolous worms that feed and build tubes using a pair of prehensile palps, usually grooved, that arise dorsolaterally from the peristomial segment. Adult morphology varies widely between the spioniform families and often within them. Seven families are classified within Spionida as delineated by Rouse and Fauchald (1997): Apistobranchidae, Chaetopteridae, Magelonidae, Poecilochaetidae, Spionidae, Trochochaetidae, and Uncispionidae. These polychaetes, especially those genera and species normally included in the Spionidae, are among the most familiar invertebrates of coastal benthic communities. Spioniforms have been the subject of numerous studies including comprehensive reviews of reproduction (Söderström 1920; Franzén 1956; Blake and Arnofsky 1999), development (Hannerz 1956; Blake 1969), anatomy (Orrhage 1964), and systematics (Foster 1971; Blake and Kudenov 1978; Blake 1996). The Chaetopteridae and Magelonidae are also well-known polychaete families. Most species in these families occur in shallow-water habitats although a few chaetopterids are known from deep water. The remaining smaller families are less familiar, largely because they occur in deeper water and are not readily encountered in the near-coastal habitats where spionids dominate. Exceptions include a few shallow-water species of Trochochaeta, Poecilochaetus, and Apistobranchus. The most recent review of spioniform reproduction was by Blake and Arnofsky (1999). Spioniforms have a strong capacity to regenerate, two types of asexual reproduction, two distinct types of eggs and patterns of oogenesis, two distinct types of sperm and spermiogenesis, widely varying methods of spawning and larval development, elegant planktic larvae, and the capacity to utilize these diverse processes in establishing a dominant position in benthic assemblages. The review by Blake and Arnofsky (1999)

ENSR Marine and Coastal Center, 89 Water Street, Woods Hole, MA 02543

#$$ Reproductive Biology and Phylogeny of Annelida provided tables of all known studies on reproduction and development of spionids. No effort is made here to repeat those summations and readers are referred to that paper for data and references. For this review, important processes and patterns in spioniform reproduction and development are summarized together with the presentation of previously unpublished data on development of selected species from California and elsewhere studied by the author. The sections that follow begin with a review of the phylogeny and systematics of Spionida and a partial revision of the traditional classification. The basis for this review is the inclusion of reproductive and developmental morphology and biology with traditional morphology. Subsequent sections deal with gametogenesis, biology of fertilization, larval development, and asexual reproduction, with some previously unpublished observations on egg and larval morphology included. Chaetopterids and magelonids are provided only cursory coverage.

13.2 PHYLOGENY AND SYSTEMATICS 13.2.1 Systematic History and Current Classification of Spionida Over the past 110 years there have been several important efforts directed toward establishing a systematic arrangement of the spioniform families and genera. The first noteworthy effort was by Mesnil (1896) who used external morphology of the prostomium, occurrence of the branchiae, and some aspects of chaetae to establish two large groups of spionid genera, but made no effort to establish subfamilies. The monograph by Söderström (1920) established for the first time that reproductive morphology was important for understanding spionid systematics. Söderström’s classification of the Spionidae included the following subfamilies and genera: Nerininae1: Nerine (now = Scolelepis), Scolecolepis (now = Malacoceros), and Aonides Laonicinae: Laonice, Prionospio, and Spiophanes Spioninae: Spio, Microspio, Pygospio, and the Polydora-complex The first two subfamilies consisted of species having thickened egg envelopes and short-headed sperm, whereas the third had species with thin egg envelopes and long-headed sperm. The Laonicinae was separated from the Nerininae based on nephridial structure and the occurrence of interparapodial genital pouches.

1

Söderström used Malacoceros and Scolelepis in an entirely different manner than is used today. The spionids presently referred to Malacoceros were his Scolecolepis, what is now called Scolelepis was Söderström’s Nerine.

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%$Hannerz (1956) analyzed larval morphology in addition to reassessing gametes and reproductive morphology and effectively combined Söderström’s Nerininae and Laonicinae into a single group, but still recognized the genera comprising Söderström’s Spioninae as a distinct entity. Hannerz noticed that the genus Scolelepis (now = Malacoceros) exhibited larval characteristics that were intermediate between his two main groups, including the presence of three pairs of black larval eyes and alternating gastrotrochs posterior to the ciliated pit, both characteristics found on larvae of the Spioninae rather than the combined Nerininae/ Laonicinae. Orrhage (1964) considered spionids from the standpoint of anatomy and morphology. His attempt at defining a systematic arrangement of the spionids resulted in a scheme that followed both Söderström and Hannerz in recognizing the Spioninae as a distinct subfamily with Spio as the primitive genus. Orrhage divided the remaining genera into subfamilies that approximated Söderström’s Nerininae and Laonicinae but with the former restricted to Nerine (now = Scolelepis). In agreement with Hannerz, Orrhage referred Scolecolepis (now = Malacoceros) to a new subfamily Scolecolepidinae. Orrhage’s spionid classification thus included: Nerininae: Nerine (now = Scolelepis) Laonicinae: Aonides, Laonice, Prionospio, and Spiophanes Scolecolepidinae: Scolecolepis (now = Malacoceros) Spioninae: Spio, Pygospio, and the Polydora-complex Pettibone (1963a) revised these genera and in so doing resolved the confusion with the use of Nerine, Scolelepis, Scolecolepis, and Malacoceros. With regard to non-spionid spioniform families, Mesnil (1897) established the Disomidae (= Trochochaetidae) as distinct and regarded it as intermediate between the Spionidae and Chaetopteridae. Söderström (1920) referred the Disomidae to subfamily status (=Disominae) within Spionidae. Mesnil (1925) supported his earlier contention that Disomidae should constitute a separate family. Allen (1904) noticed that Poecilochaetus had similarities with Spionidae. Hannerz (1956) later established the family Poecilochaetidae, regarded the Disomidae as distinct, and considered these two families and the Spionidae as forming “a well-defined group among the spiomorphic polychaetes.” Pettibone (1963b) determined that Disoma Oersted, 1843 was a homonym of Disoma Ehrenberg, 1831 in the Protozoa and resurrected Trochochaeta Levinsen to replace the junior homonym. She also recognized the genus as having family status and established the family Trochochaetidae. The Longosomidae was established by Hartman (1944) for an unusual spionid-like polychaete having distinct body regions and elongated middle body segments. Longosoma was later referred to Heterospio by Hartman (1965) and to the family Heterospionidae. The earlier family name was resurrected and modified to Longosomatidae by Borowski (1995). Green (1982) established the family Uncispionidae for a new genus, Uncispio, taken from shelf depths off southern California. She also referred

#$& Reproductive Biology and Phylogeny of Annelida the flabelligerid genus Uncopherusa to her new family. The uncispionids are also known from fragments in the western North Atlantic and are characterized by having a highly modified anterior body region with cephalic cage, noto- and neuropodial hooded hooks, and giant posterior neuropodial spines. A new species of Uncispio from the Western North Atlantic has enlarged neuropodial spines on chaetiger 3 suggesting a strong relationship to Trochochaeta (Blake and Maciolek, unpublished). According to pre-cladistic systematic efforts, the spioniform polychaetes are currently classified into the following family-level categories: Apistobranchidae Chaetopteridae Longosomatidae Magelonidae Poecilochaetidae Spionidae Subfamily Spioninae Subfamily Laonicinae Trochochaetidae Uncispionidae Systematists have more or less followed this classification for the past 20–30 years. These family-level taxa are those that comprise the so-called spioniform polychaetes or the Order Spionida. In faunal guides that provide keys to their identification, these taxa are distinguished largely by adult morphology. This classification is, however, based on empirical observations rather than quantitative analysis using phylogenetic methods.

13.2.2 Review of Phylogenetic Approaches and Suggested Classification The first effort to prepare a phylogenetic analysis was by Sigvaldadóttir et al. (1997). These authors used 25 adult morphological characteristics of the type species of 28 spionid genera as part of a parsimony analysis. Poecilochaetus, Trochochaeta, and Uncispio were used as outgroups to root the analysis. The results bore little relationship to the earlier arrangements of Spionidae suggested by Hannerz and others. Instead, four clades of spionid genera were indicated: (1) Aonidella and Xandaros2; (2) Prionospio-complex, Laonice, Spiophanes, and Aonides; (3) a large unresolved assemblage of genera including Polydora, Scolelepis, Malacoceros, and Spio; and (4) Atherospio, Pseudatherospio, and Pygospiopsis. The support for these clades was weak, and it is now apparent that the selection of outgroups was unfortunate because of the strong homology of egg and larval morphology of Poecilochaetus, Trochochaeta, and other spionid genera.

2

Xandaros Maciolek exhibits some characteristics in common with the Paraonidae and may not belong to the Spionidae.

Spionida

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Blake and Arnofsky (1999), as part of a review of the reproduction and larval development of spioniform polychaetes, developed a preliminary phylogenetic analysis of 36 genera of Spionidae, Apistobranchidae, Trochochaetidae, Poecilochaetidae, Heterospionidae (=Longosomatidae), and Uncispionidae using 38 characters. Cossura and Cirrophorus were used as outgroups; Chaetopteridae and Magelonidae were excluded. Among the 38 characters, 14 were reproductive and developmental in nature. The results of this analysis clearly showed that the classification of the Spionidae as currently defined was paraphyletic in that there were two major clades consisting of the subfamily Spioninae and a larger clade consisting of all remaining spionid genera and the genera Heterospio, Poecilochaetus, Trochochaeta, and Uncispio. A minor third clade consisting of the enigmatic Pygospiopsis (includes Atherospio) was distinct. Figure 13.1 represents four mapped character states from Blake and Arnofsky (1999) that demonstrate the importance of reproductive and larval characters in spioniform systematics. Figure 13.2 includes unweighted and weighted consensus trees illustrating the relationship of spionid genera to one another from the analysis of Blake and Arnofsky (1999). An expanded phylogenetic analysis using additional characters and taxa that included the magelonids and chaetopterids was later developed as part of a presentation at the Sixth International Polychaete Conference in Curitiba, Brazil in August 1998 (Blake and Arnofsky 2000: Abstract). This analysis added further support to the preliminary results of Blake and Arnofsky (1999) that reproductive and developmental data, when used together with adult morphology, provide a robust suite of characters to better understand the interrelationships of the spioniform polychaetes. Publication of these results is planned for the near future. However, based on Blake and Arnofsky (1999), the Spionidae with its subfamilies and the families Heterospionidae, Uncispionidae, Poecilochaetidae, and Trochochaetidae may be reduced to three clades or subfamily-level categories grouped within a more broadly defined Spionidae (Fig. 13.2B). The first clade is represented by the enigmatic and rare genus Pygospiopsis of which only 4 or 5 species are known. The second clade is restricted to the subfamily Spioninae, including Microspio, Pygospio, Spio, and the Polydora complex. The remaining spionids constitute a third family-level clade, here referred to the Nerininae, a family-level taxon established by Söderström (1920). This subfamily includes the former spionid subfamily Laonicinae and the genera Heterospio, Uncispio, Poecilochaetus, and Trochochaeta, all four of which are currently referred to family-level categories. Within the Nerininae several taxa show close relationships. For example, among the trees generated by Blake and Arnofsky (1999), a distinct subclade always includes Prionospio and its relatives. Dispio, Aonides, and Aonidella typically group together sometimes with Heterospio, Uncispio, and Spiophanes. Poecilochaetus and Trochochaeta invariably exhibit the most derived position in the majority of trees. Spiophanes does not appear to be well resolved among the Nerininae,

#% Reproductive Biology and Phylogeny of Annelida

Fig. 13.1 contd

Spionida

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possibly because its lack of branchiae is negative among the several branchial characters that were used in the analysis. The following suggested reclassification of the Spionidae should not, therefore, be taken as definitive, but rather as one step in understanding the phylogenetic relationships of a large and complex assemblage of polychaetes. Clade 1: Genus Pygospiopsis Blake (includes Atherospio Mackie and Pseudatherospio Lovell)3 Clade 2: Subfamily Spioninae Sars Microspio, Pygospio, Spio, Amphipolydora, Boccardia, Boccardiella, Carazziella, Dipolydora, Tripolydora, Polydora, Pseudopolydora, Polydorella Clade 3: Subfamily Nerininae Söderström Paraprionospio, Prionospio, Streblospio, Dispio, Aonides, Aonidella, Lindaspio, Spiophanes, Rhynchospio, Scolecolepides, Malacoceros, Marenzelleria, Scolelepis, Parascolelepis, Laonice, Heterospio, Uncispio, Poecilochaetus, Trochochaeta This classification provides a means of understanding the reproductive and developmental characteristics that have been so extensively studied for the spionids in terms of phylogeny and evolution. For example, the thickened and sometimes honeycombed eggs of the Nerininae are shown to be apomorphic and derived from spioniforms having eggs with thin membranes. Further, the chaetal and parapodial modifications of Heterospio, Uncispio, Poecilochaetus, and Trochochaeta, sometimes including loss of hooded hooks, are shown to be highly derived with respect to other Nerininae. However, the morphology of these genera is no more unusual among spioniforms than that of, Spiophanes, Lindaspio, Scolelepis, Dispio, or Aonidella, all of which have characters or suites of characters unique unto themselves. The elaborate development of both dorsal and ventral branchiae in Lindaspio is unique not only among spioniforms, but in all the Polychaeta. The very evident commonality in reproductive and Fig. 13.1 contd

Fig. 13.1. A representative tree generated with PAUP* Beta that closely approximates the consensus tree, here used to demonstrate the importance of four reproductive and larval characters in understanding the phylogeny of spionid polychaetes. After Blake, J.A. and Arnofsky, P.L. 1999. Hydrobiologia 402: 57–106, Fig. 14. Clade 1 is Pygospiopsis, Clade 2 is the subfamily Spioninae, and Clade 3 is the subfamily Nerininae. A. Character states for the egg envelope. B. Type of sperm including Ect-aquasperm for species that are broadcast spawners and Introsperm for species that produced egg capsules or are viviparous. C. Presence or absence of egg capsules or egg masses in the tubes of females. D. Presence or absence of larval nototrochs. 3

Among four described species of Pygospiopsis, are three named genera: Pygospiopsis Blake, Atherospio Mackie, and Pseudatherospio Lovell. Blake (1996) synonymized Atherospio with Pygospiopsis. Newly discovered data on postlarval and juvenile morphology for two species suggests that characters previously used at the generic level, including distribution of anterior branchiae, develop secondarily or not at all. These observations suggest that generic characters overlap and only a single genus, Pygospiopsis, is valid (Blake, unpublished).

Fig. 13.2. Two trees generated with Hennig86. From Blake, J.A. and Arnofsky, P.L. 1999. Hydrobiologia 402: 57–106, Fig. 13 that demonstrate the presence of three distinct clades among genera typically referred to five families: Spionidae, Longosomatidae (Heterospio), Uncispionidae (Uncispio), Poecilochaetidae (Poecilochaetus), and Trochochaetidae (Trochochaeta). Clade 1 is Pygospiopsis, Clade 2 is the subfamily Spioninae, and Clade 3 is the subfamily Nerininae. A. Nelson consensus tree generated from 98 most parsimonious trees resulting form equal weighting using mh* and bb*. B. Nelson consensus tree generated from three most parsimonious trees resulting from successive weighting.

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developmental morphology clearly links all of these taxa in a manner that adult morphology does not when used alone. When these results for reproductive, developmental, and adult morphology are eventually combined with molecular data, a consensus phylogeny may lead to further refinement of this classification.

13.3 MORPHOLOGY OF THE FEMALE REPRODUCTIVE SYSTEM, OOGENESIS, AND MORPHOLOGY OF THE EGGS All spionids have paired ovaries. However, spionids have two patterns of ovary structure and subsequent oogenesis (intraovarian and extraovarian) and are one of the few polychaete families to have more than one type of oogenesis (Eckelbarger 1983, 1988, 1992). In species with intraovarian oogenesis, the oocytes are retained in the ovary where they are closely associated with blood vessels from which they derive nutrition. In species with extraovarian oogenesis, the oocytes enter the coelom where they develop in association with coelomocytes. Intraovarian oogenesis has been reported for Poecilochaetus serpens, Streblospio benedicti, and Marenzelleria viridis (Eckelbarger 1980; Allen 1904; Bochert 1996a). In each middle body segment, these species have a single pair of ovaries that is located on nephridial blood vessels and covered by a thin layer of peritoneal cells. Extraovarian oogenesis has been described for Polydora and Spio (Dorsett 1961; Eckelbarger 1992). The ovaries are attached to muscles near the ventral midline; oocytes are released into the coelom where they continue to develop. This pattern of gamete production and spawning was described by Dorsett (1961) for Polydora ciliata and is here illustrated for the first time for P. cornuta (Fig. 13.3A–C). The gonads of P. ciliata and P. cornuta arise from the medial border of the ventral longitudinal muscle in the middle of a few anterior chaetigers. The ovaries appear as a pair of club-shaped sacs that project into the coelom (Fig. 13.3A). In P. ciliata, the oocytes remain in the ovaries until they reach a diameter of 25–30 µm at which time they are released into the coelomic cavity; this pattern appears to be the same for P. cornuta (Fig. 13.3B). After release from the ovaries, the oocytes move posteriorly and accumulate in the parapodial cavities of middle body segments where they continue to grow and mature to a maximal size of about 130 µm (Dorsett 1961). This same pattern has been observed in other polydorids including Boccardia proboscidea (Woodwick 1977) (Fig. 13.3D–E). Vitellogenesis has not been well documented in spionids, although two different types have been reported (Eckelbarger 1992). Streblospio benedicti has intraovarian oogenesis and accumulates yolk by the production of yolk by cells outside the oocyte, a process called heterosynthesis. In Polydora cornuta, which has extraovarian oogenesis, yolk is produced by the oocyte, a process called autosynthesis (Eckelbarger 1992).

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Fig. 13.3. Extraovarian oogenesis in Polydora and Boccardia. A–C.Polydora cornuta: A. One of two paired ovaries arising along ventral longitudinal muscles of anterior setigers of adult females. Original from Morro Bay, California. B. Another specimen showing oocytes being released into coelom and others that are developing. Original from Morro Bay, California C. Nephridium of a third specimen showing nephrostome into which eggs will move and eventually be carried to egg capsules. Original from Morro Bay, California. D–E. Boccardia proboscidea: D. Paired ovaries attached to ventral coelomic epithelium at base of intersegmental epithelium arising from ventral longitudinal musculature (vlm). Origin and development of oocytes (e) within the ovaries is apparent. After McEuen, F.S. 1979. M.S. Thesis, University of the Pacific, Stockton, California, Fig. 65. E. Cross section of mature female showing coelom packed with eggs. After Woodwick, K.H. 1977. pp. 347–371. In Reish, D.J and Fauchald, K., (eds) Essays on Polychaetous Annelids in Memory of Dr. Olga Hartman. Allan Hancock Foundation, University of Southern California, Los Angeles, Fig. 1.

According to Blake and Arnofsky (1999), three different types of eggs occur in spionids: (1) eggs with complex thick, often highly ornamented egg envelopes (= membranes) that may appear to be honeycombed and containing prominent and numerous cortical alveoli (= membrane vesicles); (2) eggs with thick egg envelopes, probably formed of several glossy layers that have a reticulated, but not honeycombed surface and lack cortical alveoli; and (3) eggs with thin envelopes consisting of a single layer that is never ornamented and lacks cortical alveoli.

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The first type of egg occurs in the genera Aonidella, Aonides, Dispio, Laonice, Lindaspio, Malacoceros, Marenzelleria, Parascolelepis, Rhynchospio, Scolelepis, Scolecolepides, Spiophanes, Poecilochaetus, Heterospio, Trochochaeta, and Uncispio (Fig. 13.4A–B). In cases where eggs become large and support direct or lecithotrophic larvae, the sculpturing of the egg envelope is usually altered or lost (e.g., Trochochaeta carica). The second egg type occurs in the Prionospio-complex, including Streblospio, which has a 3-layered egg envelope and appears to be intermediate between the highly ornamented types and those with thin, single-layered egg envelopes. The third egg type occurs in chaetopterids, Magelona, Apistobranchus, Pygospiopsis, Microspio, Spio, the Polydora-complex, and Pygospio (Fig. 13.4C). In Pygospio anucleate eggs that eventually become nurse cells for developing embryos (Fig.

Fig. 13.4. Egg morphology in Spionidae. A–B. Fertilized eggs showing thick egg membranes and honeycombed surface. Originals: A. Spiophanes duplex fertilized egg from Tomales Bay, California. B. Scolelepis sp., recently fertilized eggs from Tomales Bay, California. C–D. C. Spionid eggs with thin egg membranes: Pygospio elegans, normal oocyte from coelom surrounded by non-developing nurse eggs (After Rasmussen, E. 1973. Ophelia, 1973: 1–495, Fig. 29A). D. Gonadal smear of ripe female of Polydora cornuta showing oocytes. Original.

#%$ Reproductive Biology and Phylogeny of Annelida 13.4C) sometimes accompany the normal oocytes in the coelom. Figure 13.4D shows developing oocytes in Polydora cornuta. Eggs with thickened envelopes may be flattened and elliptical in outline or spherical. When observed with light microscopy, the honeycombed egg envelope appears to be perforated by pores that connect cytoplasmically to the cortical alveoli (Allen 1904; George 1966). The number of pores and alveoli varies among genera. The alveoli of Aonides and Dispio are few but large and arranged in two rows. Hannerz (1956) suggested that species having eggs with thin envelopes were derived from genera having eggs with thick envelopes in connection with a change from demersal spawning to brood protection. However, eggs with thin envelopes occur in several other non-spioniform polychaetes, suggesting that it is the less common thick-enveloped eggs that are apomorphic. If so, then spionids having thin-enveloped eggs are plesiomorphic. This hypothesis has been tested and supported as part of a phylogenetic analysis (Blake and Arnofsky 1999; this study). In eggs with thick envelopes, the cytoplasm pulls away from the envelope after fertilization and concentrates in the middle (Allen 1904; Hannerz 1956; George 1966; Blake and Arnofsky 1999). Hannerz (1956) speculated that the pores in the envelope allow water to enter and exert a constant pressure on the cytoplasm. As the embryo grows, the original egg envelope is stretched and smoothed out, becoming incorporated into the larval cuticle. Cilia and chaetae protrude, probably through pores. This process was clearly demonstrated by George (1966) for Marenzelleria viridis. Hannerz (1956) found this type of development in all spionids with thick membraned eggs. Detailed observations of the ultrastructure of spionid eggs and oogenesis are available for only four species: Polydora cornuta, Spio setosa, Streblospio benedicti, and M. viridis (Eckelbarger 1980, 1984, 1992, 1994; Bochert 1996a). All three types of egg envelope are represented. Spio setosa and P. cornuta have thin, single-layered egg envelopes containing simple paired and individual microvilli, respectively (Eckelbarger 1984, 1992, 1994). In S. setosa the microvilli are elongate, thin, double V-shaped structures (Fig. 13.5A); whereas, in P. cornuta the individual microvilli are shorter, solitary, and bulbous structures (Fig. 13.5B). In both cases, the tips of the microvilli project through the egg envelope where they are in direct contact with coelomic or egg capsule fluid. Cortical alveoli are absent in both species. The structure of this spionid egg envelope and their microvilli is virtually identical to those of several Capitella sibling species described by Eckelbarger and Grassle (1983). The egg envelope of Streblospio benedicti consists of three layers having unusual digitiform microvilli that bifurcate basally and lie nearly parallel to the surface (Eckelbarger 1980). The inner and middle layers of the egg envelope consist of filamentous, electron-dense material. The microvilli produce glycocalyx strands that form the outer layer of the egg envelope (Fig. 13.5C). Cortical alveoli are absent.

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Fig. 13.5. Spionid egg envelopes. A. Spio setosa. After Eckelbarger, K. 1992, pp.109–127. In Harrison, F.W. and Gardiner, S.L. (eds), Microscopic Anatomy of Invertebrates, Vol. 7: Annelida. Wiley-Liss, Inc., Fig. 23. B. Streblospio benedicti showing three distinct layers of egg envelope and bifurcate microvilli. After Eckelbarger, K. 1984, In Fischer, A. and Pfannensteil, H.-D. (eds), Polychaete Reproduction: Progress in Comparative Reproductive Biology. Fortschritte der Zoologie: 29: 123–148, Fig. 43. C. Polydora cornuta. After Eckelbarger, K. 1984, In Fischer, A. and Pfannensteil, H.-D. (eds), Polychaete Reproduction: Progress in Comparative Reproductive Biology. Fortschritte der Zoologie: 29: 123–148, Fig. 42. D. Marenzelleria viridis, unfertilized egg showing germinal vescicle, cortical alveoli (A), and thick egg envelope (EE). After George, D. 1966. Biological Bulletin: 130: 76–93, Fig. 5. E. Ultrastructure of egg envelope of Marenzelleria viridis. Modified from Bochert, R. 1966a. Invertebrate Reproduction and Development 29: 57–69, Fig. 30.

#%& Reproductive Biology and Phylogeny of Annelida Bochert (1966a) described the ultrastructure of the thick honeycombed egg envelopes and large cortical alveoli of Marenzelleria viridis. Ten to 18 large, cortical alveoli or vesicles occur just below the surface and are connected cytoplasmically to pores in the envelope (Fig. 13.5D). The ultrastructure of the egg envelope of the mature oocyte of M. viridis suggests that the honeycomb appearance is due to furrowing of the surface (Fig. 13.5E). Bochert (1996a) illustrates the furrows as extending up to 4 µm below the surface of the egg envelope. Individual microvilli are single structures that become elongate and branch irregularly as development of the oocyte proceeds. The tips of the microvilli extend through the egg envelope where they terminate in spherical granules. According to Bochert (1996a), the high density of the tips of the microvillae (50–60 per µm2) greatly increases the available surface area of the oocytes. The total surface area produced by the honeycombed structural additions as well as the tips of the microvillae is many times the basic spherical area of the mature egg. Bochert suggested that the increased surface area might facilitate movement of molecules across the membrane during development. However, because the eggs of M. viridis are spawned into seawater where they are then fertilized and subsequently undergo embryonic and post-embryonic development, it is equally likely that the increased surface area plays a role in developmental processes as well (see below). To date, there have been no detailed studies concerning the fate of the egg envelope following fertilization and subsequent embryonic development. There is also no ultrastructural documentation concerning the nature and fate of the cytoplasmic connections between the cortical alveoli and the egg surface. With the light microscope, George (1966) clearly demonstrated that the alveoli play a role in post-fertilization events, including the shrinkage of the cytoplasm from the egg envelope toward the center. Thin strands of cytoplasm maintain a connection to the egg envelope and even pull portions of it downward forming depressions or craters on the surface. Allen (1904) described an identical process in Poecilochaetus serpens.

13.4 MORPHOLOGY OF THE MALE REPRODUCTIVE SYSTEM, SPERMATOGENESIS, AND MORPHOLOGY OF THE SPERM Retzius (1904) and Franzén (1956) defined two types of sperm: (1) “primitive,” referring to short-headed sperm that were spawned into seawater and (2) “aberrant,” referring to sperm that were modified and associated with copulation or a modified form of sperm transfer. Primitive sperm were subsequently referred to as “aquasperm” (Jamieson, 1986a–b) and “aquatic sperm” (Baccetti, 1979). Rouse and Jamieson (1987) and Jamieson and Rouse (1989) refined these definitions and introduced the terms “ect-aquasperm” and “ent-aquasperm” for the “primitive” types. Ectaquasperm are those that are freely spawned into seawater and that fertilize

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eggs in that medium. Ent-aquasperm are released and swim in seawater but do not fertilize in the external water being instead drawn into the tube or burrow of the female by inhalant feeding currents. To account for the aberrant sperm, Rouse and Jamieson (1987) introduced the term “introsperm.” Ect-aquasperm and introsperm are found among spioniforms. Among spioniforms, the ultrastructure of ect-aquasperm has been reported for Prionospio cf. queenslandica (Fig. 13.6A) by Rouse (1988) and Marenzelleria viridis (Fig. 13.6B) by Bochert (1996b). The reproductive biology and light microscopy investigations of the sperm of species of the genera Scolelepis, Aonides, Laonice, Malacoceros, Parascolelepis (Fig. 13.6C), and Spiophanes (Fig. 13.6B) suggest that they also have ect-aquasperm. Rouse (1999) reported that chaetopterids have ect-aquasperm. Introsperm are found in all genera of the Polydora-complex, as well as Microspio, Pygospio, Spio, and Streblospio where considerable data are available on spermiogenesis and a wide range of sperm morphology. Mature sperm of polydorids have elongate heads (Fig. 13.6E–F) and typically range from 59–74.5 µm long (Blake 1969). Sperm break away from aggregates of sperm plates when mature and lie free in the coelom. Ultrastructural details concerning spermatogenesis in Polydora may be found in Rice (1981) and Rouse (1988). The type of sperm among genera of the Spionidae sensu lato, with one exception, separates clearly between the subfamilies Nerininae (ectaquasperm) and Spioninae (introsperm). The only exception is Streblospio which has an unusual mode of larval brooding and produces introsperm. Streblospio is the only known genus of the newly named subfamily Nerininae to have introsperm. The morphology of spionid sperm has now been documented for more than 30 species (see Blake and Arnofsky 1999: Table 2). The structure of spionid ect-aquasperm includes a spherical or ovoid nucleus, a midpiece consisting of four large, rounded mitochondria that surround two centrioles, and a free flagellum or tail (Rouse 1988; Bochert 1996b). The acrosome is typically a small, cylindrical structure that rests in a depression on the anterior end of the nucleus (see Fig. 13.6A–D). In contrast, the morphology of introsperm includes various elaborations of the nucleus and midpiece. In polydorids, both the nucleus and midpiece are elongated; the acrosome is conical with a distinct substructure (see Fig. 13.6E–F). In Streblospio the nucleus is long and the midpiece is short; the acrosome is long and spiral (Rice 1981). Membranebound electron-dense bodies are present throughout the nucleus and midpiece of polydorids and the nucleus of Streblospio (Rice 1981; Rouse 1988). Other modifications include a spiral nucleus in Spio setosa (Simon 1967) and an unusually long nucleus and midpiece with an unusually short flagellum or tail in Boccardiella hamata (Blake 1965; Rice 1992). Rice (1981) postulated that inseminated females of polydorids and Streblospio benedicti should be able to store sperm for prolonged periods

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Fig. 13.6. Examples of spermatozoa in Spionidae. A–D. Ect-aquasperm. A. Prionospio cf. queenslandica. After Rouse, G. 1988. Acta Zoologica 69: 205–216, Fig. 1. B. Marenzelleria viridis. After Bochert, R. 1966. Acta Zoologica, 7: 191–199, Fig. 19. C. Spiophanes bombyx. After McEuen, F.S. 1979. M.S. Thesis, University of the Pacific, Stockton, California, Fig. 39. D. Parascolelepis cf. tridentata (After McEuen, F.S. 1979. M.S. Thesis, University of the Pacific, Stockton, California, Fig. 36.). E–F. Introsperm cross sections. E. Polydora ciliata. After Franzen, Å. 1974. Pp. 267–278. In Afzelius, B.A., The Functional Anatomy of the Spermatozoan. Pergamon Press, Oxford and New York, Fig. 13.5. F. Tripolydora sp. After Rouse, G. 1988. Acta Zoologica 69: 205–216, Fig. 28D. Abbreviations: Acr, acrosome; Nu, nucleus; mC, mitochondria; mP, middle piece.

without loss of viability. Such an adaptation would be ecologically important for species that produce multiple broods within a single season (= polytelic). The morphology of seminal receptacles has not been well documented but may be quite variable. McEuen (1979) described these structures in four spionids from northern California. He found that Streblospio benedicti had only three seminal receptacles per female, but these were large, occupying the entire length of a segment. Seminal receptacles for Pseudopolydora kempi

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and P. paucibranchiata were dorsal, present in all epitokal segments, and anterior to the upper nephridial canal. The seminal receptacle of P. paucibranchiata exits through a U-shaped duct that is lined with cuboidal cells (Fig. 13.7E). Seven seminal receptacles were encountered in Pygospio californica. The duct of the seminal receptacle for this species exits as a narrow curved channel lined by cuboidal cells (Fig. 13.7F).

Fig. 13.7. Spermatophore formation and morphology of seminal receptacles. A–D. Spermatophore formation in Polydora cornuta A. Diagrammatic cross section of male showing paired nephridia simultaneously discharging spermatophores, which fuse and form a paired structure. After Rice, S. 1978. Transactions of the American Microscopical Society 97: 160-170, Fig. 1. B. Paired spermatophores. After Rice, S. 1978. Transactions of the American Microscopical Society 97: 160-170, Fig. 3. C. Diagram of nephridium showing distinct regions as described in the text. After Rice, 1980. Zoomorphologie 95: 181-194, Fig. 1. D. Cell from region 5 where spermatophores are believed to form. After Rice, 1980. Zoomorphologie 95: 181-194, Fig. 2D. E. Seminal receptacle of Pseudopolydora paucibranchiata. After McEuen, F.S. 1979. M.S. Thesis, University of the Pacific, Stockton, California, Fig. 56. F. Seminal receptacle of Pygospio californica. After McEuen, F.S. 1979. M.S. Thesis, University of the Pacific, Stockton, California, Fig. 58.

13.5 MATING AND FERTILIZATION 13.5.1 Broadcast Spawners Chaetopterids spawn their eggs directly into seawater. Magelonids are believed to spawn directly into seawater (see below). Spionid polychaetes either spawn their gametes directly into seawater or males transfer sperm,

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usually in spermatophores, to females, where the eggs are fertilized as sperm are transferred to egg capsules. Broadcast spawning is believed to occur in most of the spionid genera referred to the subfamily Nerininae. These genera include: Paraprionospio, Prionospio, Dispio, Aonides, Aonidella, Lindaspio, Spiophanes, Rhynchospio, Scolecolepides, Malacoceros, Marenzelleria, Scolelepis, Parascolelepis, Laonice, Heterospio, Uncispio, Poecilochaetus, and Trochochaeta. Unfortunately, there is little more than scattered empirical and often anecdotal data to support this conclusion. Blake and Arnofsky (1999) noted that the absence of observations on spawning behavior in these genera is a major gap in an otherwise large and elaborate literature on reproduction and development. What few data are available was presented in Appendix 1 in Blake and Arnofsky (1999). It would appear that two patterns of broadcast spawning occur: (1) dissemination of eggs and sperm into the water column where fertilized eggs develop freely into larvae; (2) production of eggs and sperm by paired males and females resulting in an egg mass within which fertilized eggs develop to a stage where they enter the plankton as functional larvae. My own observations of eggs and larvae taken from the plankton in northern California strongly suggest that Spiophanes bombyx, S. duplex, and Dispio uncinata spawn their gametes directly into seawater where development proceeds in its entirety. This seems to be confirmed by the fact that the earliest planktic stages for these species ranged from fertilized eggs through pre-trochophores. George (1966), working in Nova Scotia, observed broadcast spawning in Marenzelleria viridis (as Scolecolepides). George suggested that spawning was stimulated by changes in salinity. Bochert and Bick (1995), however, concluded that spawning of M. viridis in the Baltic Sea was timed to decreasing water temperature because high densities of fertilized eggs were observed when the temperature dropped to 15°C. [N.B. In summer 1967, high densities of M. viridis larvae were observed in Penobscot Bay, Maine. These larvae were taken in plankton tows through lenses of low-salinity surface water. Larvae were rare or absent in deeper high-saline water (Blake, unpublished). These observations support the suggestion by George (1966)]. There is evidence that simple post-spawning cocoons or egg masses are formed by some species of Scolelepis and Parascolelepis. For example, Imajima (1959) observed simple external cocoons for P. yamaguchii (as Nerinides) anchored in the sediment. Blake (personal observation) observed the same structures for P. cf. tridentata (see below). It is also likely that mucous egg masses are formed after fertilization. Guérin and Kerambrun (1984) identified such an egg mass in a sibling species of Malacoceros fuliginosa. Formation of any post-spawning egg mass or cocoon would logically require pair formation among adults, but there are no observations to confirm this. Richards (1970), working in Barbados, found spermatophores of what she thought was Scolelepis squamata and concluded that fertilization was internal, but was not able to confirm this with actual observations.

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Very little is known about the reproduction of magelonids, although it is likely that they spawn their gametes freely into seawater. Wilson (1982), as part of a study of larval development of three species in British waters, provided some information on the timing of sexual maturity and on the size and form of eggs. For Magelona filiformis Wilson, mature gametes were collected from April–October, but the best fertilizations were made from gametes collected in August. Females are pink and males cream colored. Unfertilized eggs measure 100 µm in diameter. Fertilized eggs develop into swimming trochophores in 20 h. Mature adults of M. alleni Wilson were collected in August. Mature eggs are pink and measure 150 µm, with the resulting trochophores being approximately 1.5 times larger than those of M. filiformis at the same stage of development. The third species, studied by Wilson (1982), was M. mirabilis Johnston. Mature eggs are 100 µm in diameter and cream colored. Mature eggs that were used in successful fertilizations were collected between early May and the middle of August, with the best results obtained in August. These results suggest that all of the British species of Magelona exhibit seasonality in the maturation of gametes, with natural spawning likely in late summer.

13.5.2 Spermatophore Formation and Sperm Transfer Spermatophores have been described for Streblospio benedicti, Polydora cornuta, P. websteri, Tripolydora sp., Microspio mecznikowianus, Spio filicornis, and Pygospio elegans (Söderström 1920; Franzén 1956; Greve 1974; Rice 1978, 1980; Rouse 1988). The nephridia become highly modified in segments where gametes mature and eventually serve as gonoducts for passage of eggs and sperm out of the body (Fig. 13.7A). Depending upon the species, a pair of nephridia may join and have a common nephridiopore, or there may be two separate nephridiopores. In species where spermatophores are formed, sperm are concentrated and enclosed in discrete packets that are discharged from the male nephridia (Fig. 13.7A–B). Rice (1980) investigated the formation of spermatophores in the nephridia of mature male Polydora. The nephridia of Polydora are enlarged paired urogenital organs located in several segments. Rice divided the fully developed male nephridium into seven morphological regions (Fig. 13.7C): (1) nephrostome, (2) descending nephridial canal, (3) dorsal curvature, (4) U-shaped depressions, (5) large urn-shaped depressions with long, thin microvilli, (6) U-shaped depressions (Fig. 13.7D) as in region (4), and (7) ascending nephridial canal that terminates in the nephridiopore. Spermatophores are composed of a central sperm mass surrounded by tubules that form a capsule surrounding the sperm. The tubules are identical to microvilli found in areas 4, 5, and 6 of the nephridia, Rice postulated that the tubules were derived from the same microvilli and that spermatophores were actually produced in the nephridia. The shape and size of spermatophores vary among species.

#&" Reproductive Biology and Phylogeny of Annelida Rice (1978) demonstrated that spermatophores were transferred from males to females without pair formation or without the necessity of either the male or female leaving the safety of its tube. Spermatophores were released from the male and deposited outside the tube. These were then picked up by the ciliary currents generated by the palps of the female and carried into her tube. Sperm were then stored in seminal receptacles until egg spawning and capsule formation. The morphology of seminal receptacles has not been well documented. McEuen (1979) described seminal receptacle structure for several species including Pseudopolydora paucibranchiata (Fig. 13.7E) and Pygospio californica (Fig. 13.7F). These two species have dorsal seminal receptacles that are relatively small, but found in all epitokal segments, whereas, in Streblospio benedicti, these same structures extend completely across the dorsum of chaetigers 14–16 (McEuen, 1979).

13.5.3 Egg Capsule Formation The formation of egg capsules by spionids was initially described by Söderström (1920) and later confirmed by Rice and Reish (1976) for Polydora cornuta (as P. ligni). Mucus is extruded from each nephridiopore and contacts the wall of the tube. Eggs are then squeezed through the same nephridiopores. The two adjacent streams of mucous and their eggs coalesce into a single chamber which fills with additional eggs. Capsules produced on adjacent segments bond with one another, forming a beadlike string (Fig. 13.8A). Capsules of P. cornuta are attached to the tube by two thin extensions representing the paired nephridiopores. Species having only a single nephridiopore have a single attachment for the capsules. In P. cornuta, the individual capsules are loosely joined to one another; sometimes individual capsules are separate (Fig. 13.8A). In Dipolydora commensalis, the individual capsules are tightly joined to one another (Fig. 13.8B). In Boccardia proboscidea, the capsules formed on adjacent segments do not fuse with adjacent ones and remain separate. In Dipolydora quadrilobata, adjacent capsules fuse and form a single elongate cylinder (Fig. 13.8C). Gibson and Paterson (2003) reported that the egg capsules of Amphipolydora vestalis were smooth cylinders formed of seven fused capsules that lacked stalks and were attached to the tube wall by mucous.

13.6 DEVELOPMENT The spioniform polychaetes, and in particular the family Spionidae, are among the most extensively studied within the Polychaeta in terms of reproduction and larval development. To date, the larval development of more than 100 species, subspecies, and geographic variants have been partially or completely described (see Blake and Arnofsky 1999: Appendices 1 and 2). Comprehensive accounts of larval development that treat multiple species include those of Thorson (1946), Wilson (1928), Hannerz (1956), Blake (1969), and Blake and Woodwick (1975). Various

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Fig. 13.8. Egg capsules of Spioninae. After Blake, J.A. 1969. Ophelia 7: 1-63, Figs. 39C, 39B, and 28A, respectively. A. Polydora cornuta showing loosely joined individual capsules each attached to the tube with two thin extensions corresponding to two nephridiopores on the adult female. B. Dipolydora commensalis showing tightly joined capsules each attached to the tube with two thin extensions. C. Dipolydora quadrilobata Type I showing a long cylindrical capsule where capsular material from individual segments have merged. Each section of the elongate capsule is attached to the tube by a single thin extension that corresponds to a single nephridiopore on the adult female. Note developing larvae and unfertilized eggs.

morphological characteristics of larvae including body shape, pigment patterns, ciliary organization, and to some extent chaetae can be used to identify the planktic larvae of individual species. Larval morphology, when known, can serve to distinguish one species from another and provides an additional suite of characters to develop phylogenetic analyses.

#&$ Reproductive Biology and Phylogeny of Annelida A full range of developmental types is present among the spioniforms, including pure broadcast spawners having planktic or lecithotrophic larval development, brooding in capsules and cocoons, and viviparity. In brooders, development may be direct or continue in the plankton. The type of development is usually predictable within genera, having important implications for understanding spioniform phylogeny.

13.6.1 Seasonality of Reproduction and Development In general, most species of Spionidae that have been studied to date appear to reproduce during periods when water temperature is highest (Blake 1969; Levin 1984a; Levin and Creed 1986; Sato-Okoshi et al. 1990). Typically, such species are polytelic, i.e., reproducing more than once in a season. Many species are capable of establishing dense populations during the times they reproduce because a single female can produce sequential sets of gametes. Gudmundsson (1985), working in northeastern England, found that Polydora ciliata, Pygospio elegans, and Malacoceros fuliginosus were polytelic, whereas Spio martinensis was possibly monotelic, producing no more than one brood per year. Blake (1969) found that Dipolydora concharum and D. quadrilobata Type II (both as Polydora), reproduced during the winter months. Both species were found with egg capsules and larvae in the early spring months, suggesting that gametogenesis and spawning occurred when water temperature was lower. Blake and Arnofsky (1999) speculated that the Maine Type II populations of D. quadrilobata might actually be relics or isolates of a species adapted to a more northern, subarctic climate, where a spring/summer reproduction would occur at the same temperatures found along the coast of Maine in winter/spring. Data on the reproductive biology of a species throughout its latitudinal range is necessary before these kinds of questions can be addressed.

13.6.2 Larval Development of Broadcast Spawners According to George (1966), working in eastern Canada, the males and females of Marenzelleria viridis (as Scolecolepides) are readily distinguished from one another by the color of the gametes: sperm appear white and oocytes orange/brown. Specimens held in glass tubes were observed to discharge gametes through the nephridia and propel them away by ciliary currents. Summaries of developmental patterns for approximately 50 species distributed among 16 genera of broadcast spawners have been reported in the literature (Blake and Arnofsky 1999). A few of these species, such as Parascolelepis tridentata (Southern 1914) and P. yamaguchi (Imajima 1959) are known to produce external egg masses or jelly cocoons after spawning. These cocoons are anchored in the sediment by mucous extensions (see below, P. cf. tridentata). Because there are so few data on fertilization and spawning in this group, we cannot know if post-spawning egg mass

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formation might not be more common than simple dissemination of fertilized eggs into the water column. In order to illustrate the wide variety of larvae in this group, the development of several species that cover the range of larval morphology available in this very diverse group of spioniforms is presented below. Dispio uncinata Hartman, 1951. This species is a relatively uncommon but widely distributed species in American waters. During the early 1970s, larvae of this species were collected from plankton tows in Tomales Bay, California (Blake, unpublished). A brief account of the development of this species was presented in Blake and Arnofsky (1999); additional details are presented here. This study represents the only account of the development of a species of Dispio. Eggs and larvae of D. uncinata were encountered in Tomales Bay plankton from January–March 1972. Fertilized eggs or pretrochophores are large, 300 × 360 µm, and have an elegant honeycombed egg envelope (Fig. 13.9A–B). Two rows of depressions or craters on the surface of the eggs are connected to internal cortical alveoli by thin cytoplasmic strands. After 12 h at ambient temperatures, the pretrochophores develop into elongated ciliated trochophores that measure 300 × 400 µm (Fig. 13.9C). Two red eyes are apparent. The two rows of surface depressions are still apparent; the apical cilia, prototroch, and telotroch protrude through the egg envelope. Trochophores of this type have only limited swimming abilities. After 24 h, a 5-chaetiger lecithotrophic nectochaete with prominent prototroch and telotroch has developed (Fig. 13.9D). Two pairs of red eyes are present. At this stage of development, the egg envelope is still evident, but the honeycombed surface is obscured. The chaetae are all barbed capillaries, with those of chaetiger 1 long and extending posteriorly past the pygidial segment. After 48 h, the developing larvae have eight chaetigers and measure 280 × 570 µm. The body is uniformly tan in color caused by numerous oily globules on the surface of the cuticle. This oily appearance is diagnostic, permitting easy identification in the plankton. By this stage, the egg envelope is no longer apparent. The two pairs of red eyes are shifted, with the larger pair lateral and the smaller pair dorsal and medial to the larger pair. The prototroch and telotroch are still present, and gastrotrochs have developed on chaetigers 3–7. The provisional chaetae of chaetiger 1 have barbs and extend beyond the pygidium; chaetae of chaetigers 2–8 are short and smooth. The gastrotrochs provide these larvae with an improved swimming ability. Late-stage, 5-day-old, planktic nectochaetes have 11–12 chaetigers. They are slow swimmers and tend to settle to the bottom of the culture dishes. Provisional chaetae are lost, the prototroch and telotroch are present but reduced; gastrotrochs are lost (Fig. 13.9E). Nototrochs are present on chaetigers 2–5 and are anlage of dorsal cilia of adults. Short stubby palps are directed posteriorly; dorsal and ventral lobes are present on the pygidium.

#&& Reproductive Biology and Phylogeny of Annelida The latest stage that was obtained in laboratory culture was a 13chaetiger juvenile that measured 930 µm long (Fig. 13.9F). This specimen developed from a planktic larva that had settled and undergone metamorphosis. The body is light tan but otherwise unpigmented. The cuticle is smooth, in contrast to the oily appearance of the planktic

Fig. 13.9. Larval development of Dispio uncinata. A–B. Pretrochophores showing the thick egg envelope with honeycombed surface, paired rings of surface craters leading to internal alveoli, and embryo developing in the center. Originals. C. Fully developed trochophore with apical cilia, prototroch, and telotroch well developed and extending through the egg envelope; two red eye spots are apparent. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 7B. D. Lecithotrophic nectochaete with 5 chaetigers bearing provisional setae. The egg envelope is still apparent, four red eyes are present; prototroch and telotroch are well developed. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 7C. E. Eight-chaetiger larva ready to metamorphose. Short stubby palps are present, ciliary bands are reduced. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 7F. F. Thirteen-chaetiger juvenile showing forward-directed palps, broadly rounded prostomium with narrow caruncle, and newly developed nuchal cilia. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 7G. G. Bidentate hooded hook from juvenile figured in F. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 7G. Abbreviations: EE, egg envelope; pT, prototroch; tT, telotroch; nT, nototrochs.

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nectochaete from which it metamorphosed. The palps are still short, but directed anteriorly. The prostomium is broadly rounded on the anterior margin; a short caruncle with lateral nuchal organs is present. The notopodial postchaetal lamellae of chaetigers 2–4 are enlarged and bear serrated margins. Bidentate hooded hooks begin on chaetiger 9 (Fig. 13.9G). Blake and Arnofsky (1999) noted that whereas the adults of Dispio, Scolelepis, and Parascolelepis have pointed prostomia, this similarity is not apparent in the larval morphology of Dispio. The pointed prostomium of Scolelepis and Parascolelepis is established early in the development, whereas the prostomium of Dispio is entire in the larva and postlarva, becoming established much later in the adult. Further, the morphology of the egg membrane of Dispio bears rows of large craters or depressions, previously known only for Aonides. The eggs of Scolelepis and Parascolelepis may be either finely reticulated or smooth, but never have the heavy honeycombed appearance or the large surficial depressions. Therefore, the apparent close relationship between Dispio and the Scolelepis complex is superficial. Laonice sp. Larvae believed to belong to a species of Laonice were collected from a plankton sample near the entrance to Tomales Bay, California in May 1972 (Blake, unpublished). This larva is readily distinguished from other spionid larvae because of its overall dark internal coloration and slender form. The larva depicted in Figure 13.10A–B has seven chaetigerous segments and four achaetigers; it measures 700 µm long and 130 µm wide at chaetiger 3, the peristomial umbrella is slightly wider than chaetiger 3. The body of this larva has an overall brown color due to randomly dispersed cellular pigment. This pigment is also concentrated in the pharynx and intestine and on the pygidium; the internal organs appear nearly black. The prostomium is broadly rounded with a slight indentation along the anterior margin which bears two sensory cilia (Fig. 13.10 A–B). Two pairs of red eyes are present: a small circular dorsal pair and a larger, more oval ventrolateral pair. The prototroch is carried on an enlarged peristomial umbrella, similar to that found in Scolelepis and Trochochaeta. The prototroch consists of four patches of cilia on each side; these extend ventrally along the umbrella, and merge with the cilia of the vestibule (Fig. 13.10B). There is a dorsal cleft between the prostomium and umbrella, which bears nuchal cilia. Chaetiger 1 is large and contains a raised area from which emerge long provisional larval chaetae that extend posteriorly past the pygidium. Provisional chaetae on chaetigers 2–7 are much shorter (Fig. 13.10A); all provisional chaetae are barbed. The pygidium is darkly pigmented and bears protruding bacillary glands at the posterior margin. The telotroch consists of five patches of cilia with a dorsal gap. Some short cilia are located in the median dorsal region due to the dorsally located anal opening. The buccal morphology is distinctive. The lateral lips of the vestibule merge with the peristomial umbrella; at their anterior margin, the lips fuse,

#' Reproductive Biology and Phylogeny of Annelida forming a ridge that bears four sensory cilia (Fig. 13.10B). Two patches of longer cilia occur along the ventral margin near the lateral lips. A large neurotroch extends posteriorly in a depression to near the end of chaetiger 2; a small ciliated pit is in this depression at the posterior end of chaetiger 1. There are four patches of small accessory cilia on chaetiger 1 and two large gastrotrochs on chaetiger 2. From chaetiger 3, there are four gastrotrochs per segment to segment 10. Nototrochs are absent. The intestine has three distinct regions, all of which are darkly pigmented. The pharynx is lighter in color, well muscled, and partially eversible. The stomach-intestine and coiled proctodeum are darkly pigmented. The identity of this species is problematic and cannot be confirmed until later stages having adult characters are identified. However, these larvae are similar to those of Laonice cirrata (Sars, 1851) described by Hannerz (1956). The shape of the prostomium and relative size of the peristomial umbrella of Hannerz’s larvae are nearly identical to the complex ciliary patterns associated with the vestibule of the present larvae. The present larvae, however, are richly pigmented, whereas Hannerz (1956) reported his L. cirrata larvae as being unpigmented. Two species, L. cirrata, and L. nuchala Blake, 1996 have been reported from Central California (Blake, 1996). If L. cirrata is correctly identified from the eastern Pacific, then it is probable that these larvae belong to L. nuchala. Spiophanes cf. bombyx (Claparède, 1869). Larvae closely resembling those described for Spiophanes bombyx by Hannerz (1956) and observed in New England waters were found in Tomales Bay, California in November 1971 (Blake, unpublished). There are subtle differences in larval morphology that suggests several closely related species might be present among the widely distributed populations collectively referred to S. bombyx (Fig. 13.10 C–D). Large planktic larvae are thick and robust and have a characteristic swimming behavior where the larva holds itself in a more or less angular position. This behavior has been seen in larvae attributed to the same species in New England waters as well as in California (Blake, personal observation); Hannerz (1956) reported the same behavior in larvae from Swedish waters. A 13-chaetiger larva measuring 925 µm long and 240 µm wide is shown in dorsal view in Figure 13.10C and a 14-chaetiger larva measuring 1,025 µm long and 250 µm wide is shown in ventral view in Figure 13.10D. The prostomium is broadly rounded on the anterior margin and bears two short lateral horns. The palps are short, thick, and attached to lateral swellings, possibly corresponding to the umbrella of related genera. There are two pairs of red eyes, the lateral pair is larger and has a distinct lens. Body segments have well-developed noto- and neuropodial lamellae. The pygidium is large, bulbous, has several prominent bacillary glands, and bears two dorsal cirri. The prototroch and telotroch are prominent. Dorsal nuchal cilia are present as two large ciliated patches just anterior to the first

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Fig. 13.10. Diversity of larvae of Spionidae Nerininae from Tomales Bay, California plankton. A–B. Laonice sp. in dorsal (A) and ventral (B) views showing the slender shape, broad prostomium and moderately developed peristomial umbrella on which the prototroch develops. C–D. Spiophanes cf. bombyx in dorsal (C) and ventral (D) views showing thick, robust body form, weakly developed peristomial umbrella, and developing frontal horns on the prostomium. E–F. Scolelepis sp. in dorsal (E) and ventral (F) views showing thickened body with fusiform shape, well-developed peristomial umbrella, and pointed projection on anterior of prostomium. Original.

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chaetiger; a caruncle will develop between these cilia. Nototrochs are present from chaetiger 5; gastrotrochs from chaetiger 3. There is a long neurotroch that extends to the end of chaetiger 2 where a large ciliated pit is located. Pigmentation includes small black speckles of pigment on the dorsum, and ventrolateral reddish or russet spots located on the posterior margin of each segment; dark pigment lines the wall of the pharynx and proctodeum. Serrated provisional chaetae are present on all segments from chaetiger 1 and are seen in the 13-chaetiger larva. However, by the 14chaetiger stage adult chaetae are evident, including neuropodial hooded hooks from chaetiger 11. These larvae differ somewhat from those described by Hannerz (1956) in being thicker in shape. The nuchal organs are larger and rounded rather than narrow; the caruncle is wider. There are similarities in pigment. The small dorsal melanophores may be more regularly distributed in the California specimens. The brick-red lateral pigment reported by Hannerz may correspond to the reddish ventrolateral spots recorded here. Spiophanes duplex (Chamberlin, 1919). Larval stages of Spiophanes duplex (= S. missionensis) are common in Tomales Bay, California, plankton during spring and summer months. The species was encountered during weekly sampling from 1971–1975 (Blake, unpublished). Larvae were especially abundant in February 1971, April–June 1972, and August 1972. Identification of the earliest stages was possible only after they were cultured to a stage that overlapped morphology of later-stage planktic larvae. Larvae were cultured in the laboratory at a constant temperature of 15°C. The earliest stages found in the plankton were recently fertilized eggs and early to late embryos in various stages of cleavage (Fig. 13.11A). The fertilized eggs are oval in shape, approximately 150 × 260 µm, and have a finely reticulated egg envelope; the developing embryo itself is green in color and opaque. Early embryos and pretrochophores have an elongated shape inside the egg envelope (Fig. 13.11B–C) and measure approximately 150 × 300 µm; with differentiation, fine apical cilia develop and protrude through the egg envelope. The egg envelope persists until the early chaetigerous larvae, at which time cilia protrude through it, and the envelope is absorbed into the larval cuticle. Early embryos are relatively immobile until the prototroch and telotroch develop. The length and width of the developing larva remains similar to that of earlier embryos until the egg envelope is entirely incorporated into the developing larval cuticle. A 3-chaetiger stage measuring approximately 140 × 275 µm developed after three days (Fig. 13.11D–E). Remnants of the egg envelope are visible on the anterior end posterior ends. The body tissue has a translucent brown cast; the gut is a bright green color. The ciliation of the anterior end includes long tactile cilia, a well-developed prototroch, neurotroch, and prominent ciliated pit. The telotroch is also well developed and consists of nine patches of cilia with a dorsal gap. Two pairs of red eyes are present; the medial pair is

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smaller than the lateral pair. Provisional larval chaetae are lightly barbed; chaetae of chaetiger 1 are longest. These larvae are highly mobile. After the egg envelope is incorporated into the larval cuticle, size increases rapidly. A 10-chaetiger planktic larva is shown in Figure 13.11F–G. These larvae are 980 µm long and fully developed planktic nectochaetes. Newly collected larvae have characteristic bright green pigment in their guts and black pigment in the pharynx and proctodeum. Lipid droplets in the gut suggest they are storing reserves. The prostomium is rounded anteriorly; frontal horns are not yet developed; minute anlagen of the palps are visible. There are two pairs of dark red, granular eyes; the medial pair is oval and the lateral pair is larger and has a distinct lens. Ciliation includes a pair of oval patches on the dorsum of the head, just medial to the prototroch. Nototrochs are present on chaetigers 4–9; double rows of cilia begin on chaetiger 5, the anterior of which includes longer cilia than those of the posterior row. Ventrally, the prototroch merges with dense, fine oral ciliation. Posterior to the vestibule, a prominent ciliated pit is present. Gastrotrochs are present on chaetigers 2–4, with those of chaetiger 2 closely associated with the ciliated pit. Chaetae are all barbed provisional capillaries. A 22-chaetiger planktic larva is shown in Figure 13.11H–I. This larva is a pre-metamorphic stage that measures 2.3-mm long and 0.3-mm wide at chaetiger 3. These larvae have little dorsal pigment except for light greenyellow markings on the anterior margin of the prostomium and brownish pigment on the pygidial segment. The dorsal wall of the intestine bears prominent lipid droplets and has an elegant blue-green color. The pharyngeal wall is lined with a dark brown pigment. This color combination alone is sufficient to recognize the larvae of this species in plankton samples. Frontal horns have developed on the anterior margin of the prostomium and the palps are well developed. The four eyes and dorsomedial ciliary patches described previously are still present; the prototroch is reduced. Ventrally, the oral ciliation and ciliated pit are still prominent. Nototrochs occur from chaetiger 3 with a double row developing from chaetiger 5. These continue posteriorly along the body with the anterior row of cilia longer than the second. These cilia are more or less retained in adults as the dorsal ciliated organs. The pygidium is a large rounded lobe, lacking cirri; the telotroch is well developed with a wide dorsal gap. Gastrotrochs occur ventrally from chaetiger 2. All body segments have noto- and neuropodial lobes. Chaetae include fascicles of smooth capillary chaetae. The ventral fascicles of chaetiger 1 include large curved crookchaetae characteristic of Spiophanes species. Multidentate unhooded neuropodial hooks occur from chaetiger 14; thicker capillary chaetae in these fascicles may represent inferior sabre chaetae. Parascolelepis cf. tridentata (Southern, 1914). Cocoons containing eggs and early larval stages of a species closely resembling Parascolelepis tridentata were collected from sandy sediments of Tomales Bay and Bodega Harbor (California) in 1971 and 1972. Planktic larvae were collected from

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Fig. 13.11. Larval development of Spiophanes duplex. A. Recently fertilized egg showing thickened and honeycombed egg envelope. B–C. Pretrochophore showing elongation of embryo within the egg membrane. D–E. Three-chaetiger larvae in ventral (D) and dorsal (E) views showing remnants of egg envelope on anterior and posterior ends, well-developed prototroch and telotroch, oral ciliation and ciliated pit, and relative length of provisional larval chaetae. F–G. Ten-chaetiger larvae in dorsal view (F) and first four chaetigers in ventral view (G). Dark pigment in the pharynx and proctodeum together with lipid droplets in the intestine are apparent. All cilia typical of such larvae are developed including prototroch, telotroch, nototrochs, gastrotrochs, short neurotroch, and ciliated pit. H–I. A 22-chaetiger larva in dorsal (H) and first four chaetigers of ventral view (I). At this advanced stage of development, the prostomial horns, parapodial lamellae and adult chaetae are present. Original.

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Tomales Bay during the same period. Blake and Arnofsky (1999) presented a preliminary account of the development of this species; additional details are presented here. Females deposit the egg cocoons; there is no evidence of brooding. Pair formation between males and females was not observed, but it is likely that such behavior is necessary to ensure fertilized eggs. The egg cocoons are large, club-shaped, gelatinous structures that average 8.5 mm long and 3.5 mm wide (Fig. 13.12A). Cocoons are light green in color and anchored in the sediment by a thin ribbon of mucous. In the laboratory, fertilized eggs (Fig. 13.12B) were observed that within 1 d developed into early trochophores that were light green in color and moved slowly within the cocoons (Fig. 13.12C). These early stages measured 170 µm long and 140 µm wide. The egg envelope, while thick, is smooth, not reticulated or honeycombed. Initially, there is a gap between the fertilization membrane and the egg envelope (Fig. 13.12B), but by the time the early trochophore develops, this gap has been filled and the egg envelope and larval cuticle have merged (Fig. 13.12C). The early trochophore is spherical in shape with a large central yolk mass. Patches of cilia mark the location of prototroch and telotroch. Four eyes are present. The larvae develop within the cocoons until they have three chaetigers, at which time they hatch and enter the plankton as larvae. The 3-chaetiger larvae are shown in Figure 13.12D–E. These larvae measure 240 µm long and 175 µm wide. By this early stage, many of the characteristics of the larger planktic larvae are apparent, including initial development of the muscular tip on the prostomium, an expanded peristomium bearing the prototroch, an apical sensory area, an elaborate oral area or vestibule that bears long tactile cilia, rudiments of a ciliated pit, and barbed provisional chaetae. The intestine is usually dark green in color, probably imparted by a diet of phytoplankton. Five- and 14-chaetiger planktic larvae are shown in Figure 13.12F–I. The 5-chaetiger stages measure 370 µm long and the 14-chaetiger stages measure 850 µm long. The most conspicuous features of these larvae are the elaboration of the peristomium into an expanded umbrella-like structure and development of a retractile, muscular structure on the tip of the prostomium. The peristomial umbrella bears the prototroch and surrounds the ventrolateral part of the prostomium. The ridge containing the prototroch curves ventrally, forming two thickened lips that border the ciliated vestibule. The muscular tip of the prostomium is probably sensory in nature and may play a role in habitat selection at the time of metamorphosis. Similar prostomial structures have been observed in related species of Scolelepis (Hannerz 1956; Scheltema et al. 1997). The cilia of the prototroch and telotroch are large, powerful organs that serve to propel these robust larvae through the water. Gastrotrochs occur from chaetigers 3–9 (Fig. 13.12H); nototrochs are not apparent until late in development. A large ciliated pit occurs ventrally posterior to the mouth (Figure 13.12F and H). A ring of nine patches of cilia surrounds the anal

#'$ Reproductive Biology and Phylogeny of Annelida segment and comprises the telotroch. The barbed provisional chaetae are longest on chaetiger 1 in the early planktic larvae, but are not much longer than those on subsequent segments in later stages. The color of these larvae is distinctive: the pharynx has a band of orange in the 5-chaetiger stage that later expands into a larger orange-brown area, and most of the gut is darkly pigmented black to dark brown. The body surface has a light green cast with some irregularly spaced brown granular pigment. An 18-chaetiger specimen ready to undergo metamorphosis is transitional between the pelagic larva and postlarva and measures 1,330 µm long (Fig. 13.12J). The peristomial umbrella is reduced and largely replaced by thickened palps. Remnants of the prototroch are still visible, together with a few cilia of the telotroch on the posterior end. Nototrochs are visible from chaetiger 2, but are more likely the developing dorsal sensory organs of the adult. The tip of the anterior end is greatly elongated into a distinctly pointed and flexible prostomium. Multidentate hooded hooks begin on chaetiger 12; these have a main fang surmounted by two pair of apical teeth (Fig. 13.12K). Scolelepis sp. Larvae of a species of Scolelepis were in the plankton of Tomales Bay, California at various times during the spring and summer of 1975 (Blake, unpublished). Larvae observed ranged from small 3-chaetiger larvae to the larger 23-chaetiger stages depicted in Figure 13.10 E–F. The generic assignment of these larvae is confirmed by the neuropodial hooded hooks that develop in these large planktic stages. Hooks (not figured) have the main fang and shaft forming a wide angle; a pair of short apical teeth surmounts the fang. In contrast, the hooks of Parascolelepis have a large main fang that forms a right angle with the shaft; there may be several apical teeth (Maciolek 1987). The large planktic larvae are about 900 µm long and generally light brown in color with a gold-colored pygidial segment. The pharynx and intestine are black in color with diverticulae extending laterally into the middle and posterior segments. As is typical of Scolelepis larvae, there is a small flexible pointed projection on the anterior end of the prostomium. The four eyes are red and arranged in a transverse row across the prostomium (Fig. 13.10E). The peristomial umbrella is fleshy and bears the remnants of the prototroch; thick extensions are the anlagen of the palps. Ventrally, a short neurotroch extends posteriorly to chaetiger 2; a small ciliated pit is located in the middle of chaetiger 2 (Fig. 13.10F). A single gastrotroch is present on chaetiger 2; subsequent segments have upwards of six patches of cilia that comprise the gastrotrochs. Dorsally, nototrochs begin on chaetiger 2 and continue for at least 20 segments. The telotroch is well developed with a narrow dorsal gap. Provisional larvae chaetae are serrated and notopodial in position. Adult neurochaetae including hooded hooks are present. Adult specimens of Scolelepis referable to S. squamata (Müller) have been collected in nearby Bodega Bay. These larvae may actually be this local

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Fig. 13-12. Larval development of Parascolelepis cf. tridentata. A. Egg cocoon. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 8A. B. Early embryo (Original). C. Early trochophore showing presence of eyes, prototroch, and telotroch. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 8B. D–E. Three-chaetiger larvae in ventral (D) and dorsal (E) views showing beginning of an apical structure on the tip of the prostomium, development of the peristomial umbrella with the prototroch, prominent oral ciliation including a well-developed neurotroch leading to the ciliated pit, and telotroch. D, After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 8 C; E, original. F–G. Five-chaetiger larvae in ventral (F) and dorsal (G) views showing development of the apical prostomial musculature and further elaboration of the peristomial umbrella (Originals). H–I. Large 14-chaetiger fusiform-shaped larva in ventral (H) and dorsal (I) views showing full development of prostomium, peristomial umbrella, gastrotrochs, ciliated pit, and internal pigmentation H. Original. I. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 8D. J. An 18-chaetiger specimen in the process of metamorphosis showing a body that is becoming more slender. The peristomial umbrella is being converted to thickened palps, the prototroch and telotroch are reduced, and adult setae including neuropodial hooded hooks (K) are developed (Original). K. Multidentate hooded hooks typical for the genus Parascolelepis. Original.

#'& Reproductive Biology and Phylogeny of Annelida species, but the identification of S. squamata, a well-known European species may not be accurate. Trochochaeta franciscanum (Hartman, 1947). It is rare, but sometimes encountered in shallow muddy sediments of Tomales Bay, California. The original records of the species were from San Francisco Bay (Hartman, 1947). There are few published data on larvae of Trochochaeta. Hannerz (1956) described late-stage planktic larvae of T. multisetosum from the Gullmar Fjord, Sweden. Larvae of T. franciscanum were collected in Tomales Bay, California from January–May 1972. A preliminary account of the larvae was presented in Blake and Arnofsky (1999); additional details are presented here. These observations represent the first descriptions of development of early and late-stage planktic larvae and changes that take place at metamorphosis. The eggs of T. franciscanum have thick, egg envelopes with prominent cortical alveoli. Hartman (1947:168) carefully described the structure of what appear to be oocytes:

“They are flat, lens-shaped, circular or nearly so. The surface is smooth and covered with a thick membrane; beneath the surface there is a surface of flask-shaped vesicles [= cortical alveoli—JAB], numbering 21-29 in each ovum; these vesicles are clear and have their narrowed end directed toward the egg membrane. Near the center there is a large germinal vesicle or nucleus.” According to the magnification in Hartman’s Fig. 1e, these oocytes would be about 50 µm in diameter. In my own observations of preserved adult specimens, the coelomic oocytes are larger, 80–100 µm in diameter and appear to have a reticulated or honeycombed surface. Hannerz (1956) recorded mature eggs of T. multisetosum with a diameter of 200–225 µm. The planktic larvae are distinctive among spioniform larvae in having unusually long provisional chaetae on chaetiger 1. These chaetae extend up to 3–4 body lengths in the early 3–4 chaetiger larvae (Fig. 13.13A–B). Pure speculation as to the role of such chaetae is that they may serve in producing a larger rigid body form that provides protection and aids mobility. With continued growth, body length matches the long chaetae (Fig. 13.13C–E). Another unusual feature of Trochochaeta larvae is the very pronounced peristomial umbrella, similar to that of Parascolelepis and Scolelepis larvae (see above) and to a lesser extent, Laonice. The peristomial umbrella serves to support the prototroch and as such provides an enlarged ciliated ring around the anterior end of the larva. The prototroch is composed of powerful long cilia on the anterior side of the umbrella and shorter, posteriorly directed cilia on the posterior side. The shorter posterior cilia merge ventrally with the heavily ciliated vestibule that surrounds the mouth. Such a ciliated “wheel” presumably serves to capture and direct food particles to the vestibule and mouth.

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Other features of Trochochaeta larvae are more typical of spionid larvae, including a ventral ciliated pit posterior to the vestibule on chaetiger 2, followed by a short neurotroch. Both of these structures develop early and persist throughout development (Fig. 13.13B–E). Gastrotrochs begin to develop by the 3–4 chaetiger stage (Fig. 13.13C). Initially, there are two widely separated cells of cilia beginning on chaetiger 2 (Fig. 13.13C–D). The gastrotrochs of chaetiger 2 are located lateral to the ciliated pit and remain small and inconspicuous in late-stage larvae. From chaetiger 3, these cilia eventually include four patches or cells of cilia across the ventrum of each segment at least through chaetiger 11 in a 17-chaetiger larva (Fig. 13.13E); chaetigers 12–17 have only a single pair of ciliated cells. Nototrochs appear to be entirely absent. Four pairs of red eyes are present throughout development. Initially, the four red eyes are dorsal (Fig. 13.13A), but with development of the peristomial umbrella, the anterior pair of eyes is carried to a ventrolateral position and can be seen in ventral view (Fig. 13.13C–E). The palps first appear at the 10–12 chaetiger stage. The right palp develops ahead of the left palp. This situation persists through metamorphosis (Fig. 13.13F). The reason for such an unequal rate of palp development is unknown. The size of larvae depicted in Fig. 13.13 proceeds from the small 3chaetiger stage (Fig. 13.13A–G: 210 µm long), to the 8-segment or 4chaetiger stage (Fig. 13.13C: 440 µm long), the 11–12 chaetiger (Fig. 13.13D: 780 µm long), and the fully developed 17-chaetiger (Fig. 13.13E: 1.02 µm long). In the transition to the adult, modified notochaetae of chaetiger 3 and brush-tipped chaetae on chaetiger 4–7 are similar to those of the adults and are present in the largest planktic larvae (Fig. 13.13E). A newly metamorphosed juvenile is depicted in Figure 13.13F together with adult chaetae from the same specimen (Fig. 13.13G–J). Transition from larva to juvenile has not been previously reported for Trochochaeta. This juvenile metamorphosed in culture 7 d after the stage depicted in Fig. 13.13E. This specimen had 16 chaetigers, was contracted and measured 588 µm long and 300 µm wide. All specimens that underwent metamorphosis were similarly contracted. Changes at metamorphosis include reduction of the peristomial umbrella resulting in the ventrally located eyes shifting to a lateral position. Most of the prototroch was lost with only a few short cilia remaining. Palps have grown, but are still unequal in length. Ventrally, gastrotrochs have been lost. The mouth, with reduction of the peristomial umbrella appears as an opening between two lateral lips. The telotroch has been lost leaving the pygidium as a simple lobe. The first two chaetigers are obscured in the dorsal view illustrated. Careful inspection shows that chaetiger 2 bears modified spines (Fig. 13.13I). Chaetiger 3 is evident together with its large modified spines (Fig. 13.13J). Chaetigers 4-9 will become the adult thoracic segments but here have conspicuous notopodial post-chaetal lamellae and a lateral swelling that obscures the neuropodial lamellae. Notochaetae of the thoracic segments are simple capillaries at this stage; neurochaetae are the characteristic brush-tipped spines and fringed capillaries characteristic of the adult (Fig. 13.13G–H).

$ Reproductive Biology and Phylogeny of Annelida

Fig. 13.13 Larval development of Trochochaeta franciscanum. A–B. Three-chaetiger larvae in dorsal (A) and ventral (B) views showing long provisional setae and advanced development of the peristomial umbrella with prototroch. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Figs. 6A–B. C. Eight-segment larva in ventral view. Gastrotrochs and oral ciliation with neurotroch and ciliated pit fully developed. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 6C. D. Twelve-chaetiger larva (Original). E. Seventeen-chaetiger larva, ventral view, showing short left palp and with fully developed gastrotrochs. After Blake, J.A. and P.L. Arnofsky. 1999. Hydrobiologia 402: 57–106, Fig. 6D. F. Newly metamorphosed juvenile in dorsal view showing reduction of the peristomial umbrella, development of the left palp ahead of the right palp, and elaboration of adult parapodia and setae. Original. G–J. Adult chaetae present in juvenile. Original. Spines of chaetigers 2 and 3 represented by Figs I and J, respectively. Brush-tipped spines and fringed capillaries of neuropodia of thoracic chaetigers are shown in Figs. G–H.

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Development of Chaetopteridae. Gametes of chaetopterids are dispersed directly into seawater where fertilization takes place. Natural reproductive processes are poorly known, but laboratory procedures have been reported where gametes can be obtained and eggs fertilized (Costello et al. 1957; Eckberg and Hill 1996; Irvine and Martindale 1999). Enders (1909) provided a detailed account of larval development and metamorphosis in Chaetopterus variopedatus4 that was based both on larvae reared from fertilizations and later stages taken from the plankton. Werner (1953) and Cazaux (1965) also described larvae of C. variopedatus. A detailed account of the larval ontogeny of Chaetopterus was published by Irvine et al. (1999) who described the transition of larval segmentation to that of the adult. Bhaud and Cazaux (1987) describe development of pelagic larvae of other chaetopterids. Other papers depicting pelagic larvae of chaetopterids include Bhaud (1966), Mileikovsky (1967), and Scheltema (1974). In general, chaetopterid larvae are large, robust; sometimes more than 1-mm long and have a spherical body divided into three regions. When the nectochaetes are fully developed, they have a pair of palps, chaetae, and a powerful mesotroch that propels them through the water. Larvae with up to three mesotrochs have been seen (Blake, personal observations). The anterior end and oral opening are covered with fine cilia, and there may be additional sensory cilia in various regions on the body. The posterior end of the larvae becomes elongated late in development and this serves as an attachment organ during metamorphosis, although little is actually known of this process. According to Bhaud (1978), the modified spines of chaetiger 4 are similar in the adults and larvae and it should be possible to identify planktic larvae once the structure of the adult chaetae is known. This hypothesis was verified during a recent study of pelagic chaetopterid larvae collected from the waters of the Antarctic Peninsula. Mesotrochal larvae of chaetopterids were identified as Phyllochaetopterus monroi Hartman based upon the structure of the modified spines in chaetiger 4 (Scheltema et al., 1997; Blake, unpublished data). Scheltema (1974) demonstrated that larvae of at least two common species, Chaetopterus variopedatus and Spiochaetopterus costarum, were widely distributed in the plankton throughout the North and South Atlantic Ocean. Scheltema speculated that these dispersal patterns explained the wide distributions of these species and provided evidence for a regular pattern of gene flow between widely separated populations. Bhaud (1978) re-examined some of Scheltema’s specimens and photographs and determined that an additional species, Phyllochaetopterus socialis, another widely distributed species, was also present in the same plankton samples. 4

The taxonomy of Chaetopterus species requires revision. Numerous accounts of C. variopedatus in the embryological literature undoubtedly refer to other species. The embryological literature, therefore, is not accurate from a taxonomic point of view and numerous descriptions of larvae referred to C. variopedatus need to be reassessed. Comments relative to problems associated with taxonomy of Chaetopterus are mentioned in Petersen (1984a, b) and Irvine et al. (1999).

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Reproductive Biology and Phylogeny of Annelida

To illustrate the diversity of chaetopterid larvae, some examples have been taken from both published and unpublished sources (Fig. 13.14). Figure 13.14A–B represents an early planktic larva of Chaetopterus (Fig. 13.14A) and a juvenile after metamorphosis (Fig. 13.14B). These larvae have apical cilia, anterior and posterior mesotrochs, two palps, rudimentary segmentation, and a simple pygidium. The transition from planktic larvae to juveniles is complex in this genus. Figure 13.14 C–E represents two larval stages of Phyllochaetopterus solitarius. These larvae have no apical cilia, a single mesotroch, palps, well-developed segmentation with spines on chaetiger 4, and no evidence of a pygidium. Mesochaetopterus minutus has two mesotrochs (Fig. 13.14F–G); M. taylori has three mesotrochs (Fig. 13.14H), the first record of three such bands in chaetopterids. For these larvae, it is very evident that the mesotrochs are associated with anlage of the abdominal region. Both species exhibit rudiments of thoracic segmentation and a tapering posterior end. In M. taylori, there are what appear to be telotrochal cilia on the posterior end. Spiochaetopterus cf. costarum, which is relatively common in California waters, has a single mesotroch, well-developed thoracic segmentation with spines in chaetiger 4, palps, and rudiments of the abdominal segments (Fig. 13.14I). Notice that the bodies of both M. taylori and S. cf. costarum are depicted as covered with fine cilia. It is probable that all such planktic chaetopterid larvae are so ciliated. The oral morphologies of Mesochaetopterus and Spiochaetopterus are elaborated into broad ciliated vestibules, which is typical of all late-stage chaetopterid larvae. Irvine et al. (1999) confirmed that the two large ciliary bands of Chaetopterus are mesotrochs and not metatrochs because rather than being presegmental, these ciliary bands eventually come to lie within the segmented trunk region. In this regard, the mesotrochs of chaetopterids may be homologous to the nototrochs and gastrotrochs of other spioniform larvae. Irvine et al. (1999) determined that segmentation in Chaetopterus larvae is heterochronous, in that segments in each of the body regions develop at different times and rates rather than the more typical homochronous pattern found in other polychaetes where segmentation during larval development is sequential. Heterochrony has likely evolved together with the specialized body regions found in all chaetopterids. Development of Magelona. The most important work on development of Magelona is by Wilson (1982), who reviewed all previous accounts and provided detailed descriptions of both early and late larvae of three species from British waters. Pelagic larvae of M. filiformis, M. alleni, and M. mirabilis were collected from April–November, but most commonly occurred from July–November over a period of several years. Wilson (1982) was able to rear these larvae in laboratory culture and induce metamorphosis. Magelonid larvae are distinctive in having a broad prostomium with terminally attached larval tentacles, a long, narrow body, and stiff larval chaetae. Wilson found that the long larval tentacles are derived from the prototroch and associated tissues. The adult palps appear late in

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Fig. 13.14. Chaetopteridae Larvae. A–B. Chaetopterus larva (A) and benthic juvenile (B). After Bhaud, M. and Cazaux, C. 1987. Oceanis 13: 597–753, Figs. 57.1–2. C–D. Phyllochaetopterus socialis, planktic larva, ventral (C) and dorsal (D) views. After Bhaud M. and C. Cazaux. 1987. Oceanis 13: 597– 753, Fig. 57.6. E. Spine from chaetiger 4 of Phyllochaetopterus socialis (same, Fig. 57.7). F–G. Mesochaetopterus minutes. After Bhaud M. and C. Cazaux. 1987. Oceanis 13: 597–753, Figs. 58.2 and 58.4. Planktic larva (F) showing two mesotrochal bands. Newly metamorphosed juvenile (G). H. Mesochaetopterus taylori, planktic larva from California with three mesotrochal bands (Original). I. Spiochaetopterus cf. costarum, planktic larva from California. Original.

$" Reproductive Biology and Phylogeny of Annelida development and originate as thickenings at the bases of the larval tentacles. The larval tentacles and long larval chaetae are shed at metamorphosis. The first two species are relatively typical, in that both larval tentacles are fully developed and the larvae are symmetrical. For M. mirabilis, however, the left tentacle is a short stump, providing an asymmetrical appearance to the larva. According to Wilson (1982), it should be possible to recognize the larvae of different species of magelonids in the plankton based upon both larval and developing adult characters. During the 1970s, larvae of at least two species of magelonids were collected from plankton of Tomales Bay, California (Blake, unpublished). The first of these is believed to be Magelona pitelkai Hartman (Fig. 13.15A– C), the most common intertidal species in central and northern California. Figure 13.15A–B represents early larvae with terminally attached larval tentacles and mouth. Distinctive dark pigment in the posterior segments of these larvae may be observed in later stages. A late-stage larva, in which the larval tentacles have been shed and replaced by the developing adult tentacles, is shown in Figure 13.15C. The prostomium is entire and somewhat blunted on the anterior margin, a feature that is characteristic for adult M. pitelkai. Figure 13.15D is that of a late-stage larva of another species having a strongly rounded anterior margin on the prostomium. This second species is probably M. sacculata Hartman, a subtidal species known from central and southern California.

13.6.3 Larval Development of Viviparous Species Among spionids, the best known example of viviparity is Streblospio benedicti which is widely distributed on all three coasts of North America, where its mode of larval brooding and development have been extensively studied (Dean 1965; Levin 1984b; Eckelbarger 1986). The females of this species brood their young in dorsal pouches (Fig. 13.16A), with larvae eventually released into the plankton. Collier and Jones (1967) determined that the dorsal brood pouches were thin-walled, dorsolateral extensions of the coelom, thus explaining how eggs can be transported from the coelomic ovaries to the dorsal brood pouches. Blake and Arnofsky (1999) provided a summary of S. benedicti literature. Two types of development have been described for S. benedicti: lecithotrophic and planktotrophic. Two forms of larval development in a single species is an example of poecilogony. Other examples are treated in Section 13.6 (see below). Lecithotrophic larvae are released from the brood pouches having 9–12 chaetigers (550–650 µm), and settle within hours or at most a few days. These larvae lack provisional chaetae, have poorly developed ciliary bands, and are weak swimmers (Fig. 13.16B). Despite the extensive literature on the larvae of S. benedicti, morphological details of the oral structures and cilia have not been described for either larval type. Metamorphosis is relatively rapid, with competent larvae developing thickened palps and branchiae (Fig. 13.16C), but still retaining cilia until the first mucous tube is

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Fig. 13.15. Magelona larvae from Tomales Bay, California. A–C. Magelona pitelkai Early larvae showing terminally attached larval tentacles (A–B). Late-stage larva showing adult tentacles (C). D. Late stage larva of Magelona sacculata. Original.

constructed. Lecithotrophic populations occur in the SE United States, Gulf of Mexico, and California (Blake and Arnofsky 1999). Planktotrophic larvae are released from the brood pouches after they have attained 4–9 chaetigers (200–300 µm long). These larvae have welldeveloped serrated provisional chaetae, and highly developed ciliary tracts (Fig. 13.16D–E) and remain in the plankton for up to 45 d, growing to 450– 550 µm in length before settlement and metamorphosis. Provisional chaetae are lost just prior to metamorphosis (Fig. 13.16F). Planktotrophic development occurs in populations along the eastern United States and in the Gulf of Mexico (Blake and Arnofsky 1999). Larvae of the European species, S. shrubsoli, develop directly from eggs carried in dorsal grooves on the body of the female (Fonseca-Genevois and Cazaux, 1985). Development continues until the 14-chaetiger stage at which

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Fig. 13.16. Larvae of Streblospio benedicti. A. Section of body of female from central California showing lecithotrophic larvae in brood pouches. B. Lecithotrophic larva from central California recently released from brood pouch. Prototroch and telotroch evident, provisional larval chaetae entirely absent. C. Transitional stage between lecithotrophic larva and juvenile undergoing metamorphosis. D–E. Planktic larvae from New England recently released from brood pouch. F. Late-stage larva from New England just before metamorphosis. Originals.

time the juveniles leave the female to crawl away and burrow into the substratum.

13.6.4 Larval Development of Brooded Species Having Egg Capsules Pygospio elegans Claparède, 1863. This species is a widely distributed spionid occurring in intertidal sediments of bays and estuaries. The species is typically found in middle to upper intertidal zones and is tolerant of low salinities. The reproduction and development of this species are well

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known and have been reported by several authors (see Blake and Arnofsky, 1999). The species typically alternates asexual and sexual phases with the largest increase in populations probably due to asexual reproduction (Armitage, 1979). However, Bolam (2004) only found sexual reproduction in a population in Scotland. Pygospio elegans exhibits poecilogony with two types of reproduction and larval development reported in the literature—with and without nurse eggs. In the first type, developing larvae feed on the extrinsic yolk available from the nurse eggs. Such larvae are retained for extended periods in the capsules until they escape into the plankton late in their larval life (Söderström, 1920; Hannerz, 1956; Rasmussen, 1973; Gudmundsson, 1985; Morgan et al., 1997). In the second type, without nurse eggs, larvae are released into the plankton with about 3–4 chaetigers and continue development as planktotrophic larvae (Gudmundsson, 1985; Morgan et al., 1997; Bolam, 2004). Morgan et al. (1997) validated poecilogony for P. elegans by comparing genetic divergence in 14 enzymatic loci. Results clearly showed that a single species was involved. Different stages in the development of larvae that depend upon nurse egg feeding or adelphophagia for nutrition (Type I) are shown in Fig. 13.17. All figures are taken from Rasmussen (1973) from Danish waters. The nurse eggs of Pygospio elegans are fragile and break up into yolk granules as the larvae derived from the fertilized embryos begin to move around in the capsules (Fig. 13.17A–B). Figure 13.17C shows an embryo engulfing nurse egg fragments. The extrinsic yolk that is engulfed is stored in the embryos as if it were derived from intrinsic sources. In effect, these embryos become encapsulated lecithotrophic larvae until they escape into the plankton (Fig. 13.17C–D). In some cases, developing larvae do not feed on the available nurse egg yolk supply. Such larvae develop in capsules to the 3-chaetiger stage and have the appearance of Type II larvae (Fig. 13.17F). The fate of these larvae is not clear; it is likely the larger lecithotrophic larvae developing in the same capsules (Fig. 13.17C) cannibalize them. A late planktic stage is shown in Fig. 13.17G; a settling stage is shown in Fig. 13.17H. The two latter stages are identical whether or not they were derived from Type I or II early larvae. Pygospio californica Hartman, 1936. Larvae of P. californica were collected from the plankton at Bodega Bay, California near the Coast Guard Station in early January 1972 and at Lawson’s Landing in Tomales Bay, California in January and April 1972 (Blake, unpublished). In separate collections, egg capsules similar to those of P. elegans were collected in high intertidal sediments in Bodega Harbor. None of these was successfully maintained in the laboratory. There was also evidence that P. californica reproduced asexually in the same manner as P. elegans. A 3-chaetiger larva measures 354 µm long and 120 µm wide. In dorsal view, the prostomium is broadly rounded along the anterior margin (Fig. 13.18A). There are two pairs of black eyes: a median circular pair and a lateral crescent-shaped pair. Dorsally, the prototroch has two patches of cilia

$& Reproductive Biology and Phylogeny of Annelida

Fig. 13.17. Larval development of Pygospio elegans (After Rasmussen, E. 1973. Ophelia 11: 1–495, Figs. 30A, 29B, 29C, 30B, 29D, 31A, 31B, and 32A, respectively). A. Egg capsules showing embryos feeding on nurse egg fragments. B. Diagram showing nurse egg breaking into yolk fragments. C. Individual embryo engulfing extrinsic yolk. D. Egg capsule showing three large, lecithotrophic 3chaetiger embryos and one smaller larva that has apparently has not been feeding on the extrinsic yolk. E. Late pre-chaetiger embryo showing cilia around vestibule used to engulf yolk particles. F. Threechaetiger planktic larva. G. Late-stage planktic larva recently released from capsule. H. Early benthic stage, just after metamorphosis.

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on each side of a developing nuchal ridge that extends to the posterior margin of chaetiger 1. The prototroch continues ventrally as several ciliated patches and merges with the oral cilia of the vestibule (Fig. 13.18B). Two patches of fine cilia arise close to the dorsal ridge and represent the developing nuchal organs. Ventrally, the buccal region is simple, with ciliation limited to the lateral lips and the area anterior to the vestibule. A short neurotroch is on an elevated ridge and extends posteriorly to a large ciliated pit on the anterior edge of chaetiger 2; where the ridge merges with chaetiger 1 there are tufts of cilia on either side (Fig. 13.18B). Three chaetigerous segments bear long serrated larval chaetae. A fourth segment is developing. Nototrochs are absent at this stage; a single gastrotroch consisting of four patches of cilia is present on chaetiger 3. The telotroch consists of seven patches of cilia with a wide dorsal gap. The anal segment is very glandular and has a slight russet color. The pharynx is transparent and colorless. The stomach-intestine is pigmented green internally (from food) but has granular black pigment in the epithelial lining. The proctodeum is black in color, short and straight. A 7-chaetiger larva develops from the previous 3-chaetiger stage in 5– 7 d at 12°C. This larva measures 510 µm long and 164 µm wide. In dorsal view, the prostomium has elongated, but otherwise the head retains the same general shape although with a slight depression on the anterior margin (Fig. 13.18C). The position of the eyes and ciliation remains the same except for the nuchal area, where the cilia are finer (Fig. 13.18D). The only other changes on the prostomium are the appearance of two long sensory cilia that arise anteriorly and point backwards along the lateral ridges and two groups of fine cilia that have developed above the median eyes (Fig. 13.18D). Provisional chaetae are dorsal and present on all seven segments; short neurochaetae have appeared (Fig. 13.18 C–D). Nototrochs begin on chaetiger 3 and continue on subsequent segments including the developing eighth segment. A dorsal pigment pattern of granular black pigment spots is present on the anterior and posterior margins of chaetigers 4–8. This pigment intensifies posteriorly and is always more prominent on the posterior margin of the segments. Black pigment also occurs on the dorsal side of the pygidium. Russet color has intensified on the anterior margin of the prostomium and pygidium. Dorsally, the pygidium has a median cleft and the telotroch has a dorsal gap in the cilia (Fig. 13.18D). Ventrally, the raised area of chaetiger 1 has fused with the lateral lips and is completely covered with cilia of an expanded neurotroch (Fig. 13.18C). The prototroch and the remaining buccal ciliation are unchanged. Short anlagen of the palps are present on the posterior lateral sides of the peristomium. Gastrotrochs are present on chaetigers 3, 5, and 7. The pygidium is glandular. Internally, the vestibule-pharynx area is lined with a distinct black pigment. The stomach-intestine and proctodeum are darkly pigmented as previously described. A 13-chaetiger larva measures 850 µm long and 210 µm wide (Fig. 13.18 E–F); this larva is approximately seven weeks older than the previously

$ Reproductive Biology and Phylogeny of Annelida

Fig. 13.18. Larval development of Pygospio californica from northern California. A–B. Three-chaetiger planktic larvae in ventral (A) and dorsal (B) views. C–D. Seven-chaetiger planktic larvae in ventral (C) and dorsal (D) views. E–F. Thirteen-chaetiger planktic larvae in ventral (E) and dorsal (F) views. G. Newly settled benthic juvenile. Originals.

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described 7-chaetiger stage. Dorsally, the prostomium has retained the same approximate shape and ciliary patterns with the following exceptions: (1) deeper russet pigmentation occurs along the anterior margin, (2) nuchal organs are larger and more prominent, and (3) the lateral eyes and their adjacent sensory cilia have shifted in their relative positions (Fig. 13.18F). The palps have elongated and there is a small ciliated patch at their point of insertion. Ventrally, the buccal region is very similar to the previous stage. The ciliated pit and neurotroch are still present. There is an additional patch of cilia along the lateral margin of the neurotroch. Provisional chaetae are in the process of being replaced by adult chaetae. Adult chaetae include only capillaries; neuropodial hooded hooks are not apparent. Noto- and neuropodial lobes are apparent on all segments. Notopodial lobes each bear a single internal bacillary gland (Fig. 13.18F). Nototrochs are present from chaetiger 3, and consist of up to seven closely spaced cells per segment. The pygidium and telotroch have retained their earlier configuration. Dorsal pigment is more intense and consists of scattered black granular from chaetiger 6 (Fig. 13.18F). This pigment is concentrated on the anterior and posterior margins of each segment, and intensifies posteriorly; similar pigment is present ventrally on the surface of chaetigers 3–6. The pygidium has a definite russet cast, and has black pigment on either side of the telotroch. Gastrotrochs occur on chaetigers 2–5, 7, 8, 11, and 12 in the patterns shown in Figure 13.18E. In younger specimens, there is also ciliation on segment 6, so it appears that this stage is the beginning of ciliary regression. A newly settled benthic juvenile is slender in appearance (Fig. 13.18G). One specimen that metamorphosed in the laboratory measured 2.06 mm long and 0.24 mm wide for 23 chaetigers. The prostomium is conical with diffuse russet pigment on the anterior margin. The four eyes are arranged in a rhomboid configuration and situated on the nuchal crest. A caruncle extends posteriorly to chaetiger 2 and bears two ovoid nuchal organs on either side. Palps are fully developed. The segments are all well developed and have adult parapodial and chaetal characteristics. The first chaetiger is reduced and retains only rudiments of parapodia and chaetae. A remnant of the larval nototroch occurs on chaetiger 2, but otherwise, nototrochs are present only on chaetigers 14–20 (Fig. 13.18G). This may be an indication of later branchial development but, at this stage, branchiae are absent. Notochaetae are simple capillaries. The neurochaetae are also capillaries until chaetiger 13, where hooded hooks begin. The dorsal pigmentation found in the planktic larval stages has disappeared. The anal segment is glandular and still bears both the russet and black pigment. The pygidium is composed of two large glandular lobes and two small lobes, each of which contains a single bacillary gland. Sensory cilia project from the pygidial lobes. The gut has developed into a long pharnyx-esophagus, which has lost much of its black pigment. The stomach is still dark and has well-developed diverticulae. The intestine has clear walls and is somewhat convoluted as is the proctodeum, which still has black pigment in its walls.

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Reproductive Biology and Phylogeny of Annelida

Boccardia columbiana Berkeley, 1927. This species is widespread in the eastern Pacific and occurs in numerous intertidal habitats where it bores into hermit crab shells, coralline algae, and tests of Balanus (Woodwick, 1963). Adult specimens with egg capsules in the burrows were collected from several intertidal habitats in northern California, including the ocean side of Tomales Point, and from burrows in shells of Tegula funebralis from tests of Balanus from rocky areas north of Dillon Beach, California. The developmental biology and larval morphology of B. columbiana are very similar to the closely related sibling species B. proboscidea (Woodwick 1977; Blake and Kudenov 1981; Gibson 1997). However, all eggs of B. columbiana are fertilized and eventually develop into planktic larvae, whereas most populations of B. proboscidea studied to date exhibit nurse egg feeding of the larvae, with release of the larvae delayed until extrinsic yolk reserves are exhausted. Egg capsules were collected at several widely spaced times of the year suggesting a continuous reproduction. The specimens on which the major efforts were concentrated were collected on 2 July 1969 from Tomales Point and on 24 February 1970, from Dillon Beach. Egg capsules are single and attached to the wall of the tube by two thin extensions (Fig. 13.19L). Several capsules usually occur together along the wall of the adult tube but are not joined to one another as in related species of Polydora (Blake, 1969). Eggs are pink, spherical in shape, and measure approximately 110–115 µm in diameter. There are approximately 50–60 eggs per capsule. All of the eggs are fertilized; there are no nurse eggs. Development within the capsules. The earliest embryo with recognizable morphology removed from an egg capsule was a pretrochophore that measured 120 µm long and about 100 µm wide (Fig. 13.19A–B). These early embryos have large ventral ciliary patches, a ciliated mouth or vestibule, and a dark yolk mass. After 24 h, an embryo having the characteristics of a trochophore is 130 µm long with some suggestion of segmentation developing at the level of the ventral ciliary patches (Fig. 13.19C–D). The vestibule is deep and heavily ciliated. Two areas of fine cilia lateral to the vestibule are the prototroch. Several isolated patches of cilia on the posterior end represent the developing telotroch. After 48 h, the larvae have three developing chaetigerous segments and are 160 µm long (Fig. 13.19E–F). These larvae are capable of slow gliding movement along the bottom of a culture dish if released from the capsule. They have two black eyes, each cup-shaped. The prototroch has developed as a prominent band of large cilia organized into distinct patches and a few isolated smaller patches that are below the main band; these smaller cilia are either lost or absorbed into the main telotroch with development. Ventrally, the lateral lips of the vestibule have developed with the prototroch extending ventrally across these lips. The telotroch has further developed and has seven individual patches of cilia with a wide dorsal gap. Provisional serrated chaetae are well developed in each of the three segments and protrude through the cuticle on chaetiger 1.

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Fig. 13.19. Larval development of Boccardia columbiana from Dillon Beach, California. A–B. Early pretrochophore embryos removed from egg capsules, ventral ciliary patches are evident (Originals). C– D. Encapsulated trochophores showing development of prototroch and telotroch C, original; D, After Blake, J.A. and Arnofsky, P.L. 1999. Hydrobiologia 402: 57–106, Fig. 11B. E–F. Early 3-chaetiger larvae removed from egg capsules Original. G–H. Four-chaetiger removed larvae from egg capsules G, original; H, After Blake, J.A. and Arnofsky, P.L. 1999. Hydrobiologia 402: 57–106, Fig. 11C. I–J. Fourchaetiger planktic larvae at a stage approximating release from egg capsules I, original; J, After Blake, J.A. and Arnofsky, P.L. 1999. Hydrobiologia 402: 57–106, Fig. 11D. The first of the large dorsal chromatophores is present. K. Large pelagic planktic larva showing dorsal chromatophores, nototrochs from chaetiger 3, and gastrotrochs from chaetigers 5 and 7. After Blake, J.A. and Arnofsky, P.L. 1999. Hydrobiologia 402: 57–106, Fig. 11E. Chaetiger 5 has modified spines at this stage. L. Egg capsule After Blake, J.A. and Arnofsky, P.L. 1999. Hydrobiologia 402: 57–106, Fig. 11A.

$" Reproductive Biology and Phylogeny of Annelida After 72 h, the larvae have four chaetigers and are 170 µm long (Fig. 13.17G–H). Yolk reserves are still apparent, but these larvae are capable of strong swimming if released from the capsules. Six eye spots are apparent, including an inner circular-shaped granular pair and a small middle pair located beside a larger cup-shaped lateral pair. The two lateral ciliated patches have shifted to a more dorsal location commensurate with further development and elaboration of the prototroch. Black pigment has developed on the dorsum of the pygidium. Development in the plankton. Four-chaetiger larvae continue to develop in the capsules until their yolk reserves are exhausted after which they are released into the plankton. The 4-chaetiger larvae (Fig. 13.19 I–J) are 280 µm long, slender in shape and very strong swimmers. They are strongly photopositive. The serrated provisional chaetae are long and serve to provide the larvae with a streamlined and rigid shape. The pigment pattern of B. columbiana larvae includes a row of large branching chromatophores along the dorsal midline. The first of these chromatophores is present over chaetiger 2. Black pigment is present on the dorsal side of the pygidial segment. Two narrow ciliated areas posterior to the medial eyes are precursors of the nuchal cilia. There are no gastrotrochs or nototrochs at this stage. The prototroch and telotroch are well developed and are used to move the larvae through the water. The ventral ciliary patches are reduced to two slender areas posterior to the mouth. A podial lobe is present on chaetiger 1. A pair of small tubercles was observed on chaetigers 3 and 4, but these were not observed in later stages and their function is unknown. An 11-chaetiger larva is 560 µm long and thick, and fusiform in shape (Fig. 13.19K). Dorsal provisional notochaetae characteristic of planktic spionid larvae are prominent on all segments except chaetiger 5; ventral provisional chaetae are present from chaetiger 6. Adult chaetae are developing and include hooded hooks from chaetiger 7 and modified spines of chaetiger 5. Modified spines of chaetiger 5 include a bristle-topped spine and a falcate spine; both are characteristic for this species. Gastrotrochs occur on chaetigers 3, 5, and 7 in this larva, but will also develop on chaetigers 10 and 13 in later planktic stages; nototrochs begin on chaetiger 3 and continue on subsequent segments. The telotroch is formed of approximately seven patches of cilia leaving a wide dorsal gap. The prototroch extends dorsally to the level of the lateral eyes; and ventrally to the oral opening. Nuchal cilia are well developed. Pigmentation is well developed. The entire body has a greenish cast; the dorsal pigmentation consists of a single row of branching chromatophores from chaetiger 2. Laterally, small black pigment spots are located between the parapodia of chaetigers 7 and 8. Some dark granular pigment occurs dorsally on the pygidium. Three pairs of black eyes are present, similar to those described for the 4-chaetiger stage. Reticulate black pigment is present anterior to the medial eyes. Dipolydora brachycephala (Hartman, 1936). Larvae identified as Dipolydora brachycephala were taken from plankton at Lawson’s Landing,

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Tomales Bay, California during December 1971 and again from January– April 1972. They were most abundant from December–February of 1971 and 1972. These larvae resemble those of P. caulleryi described by Blake (1969) from New England but differ in several aspects including the pigmentation. Adults of the two species are nearly indistinguishable and the difference in larval morphology represents the only firm evidence at present that two sibling species are represented in North America. Because of these differences, D. brachycephala is here re-established as a distinct species. A planktic larva with 4-chaetigerous segments measures 320 µm long and 120 µm wide (Fig. 13.20 A–B). Provisional chaetae are serrated and exceptionally long, often extending well past the posterior end of the body. The prototroch extends ventrally from the dorsum to near the lateral lips of the vestibule. Tactile cilia are located in various areas of the head. Six black eye spots are disposed as in Figure 13.20A; in later stages the two lateral eye spots will merge. The lips of the vestibule are heavily ciliated. A broad, triangular-shaped neurotroch extends posteriorly, terminating in a ciliated pit in the middle of chaetiger 2 (Fig. 13.20A). A single patch of cilia lies posterior of the ciliated pit. A gastrotroch on chaetiger 3 is composed of five separate patches of cilia. The telotroch has a wide dorsal gap. The anal segment contains distinctive black pigment, most prominent dorsally (Fig. 13.20B), but continuing slightly around to the ventral position. Parapodial lobes are developing on chaetigers 3–4. Nuchal ciliation is present as two circular areas on either side of the head, just posterior to the lateral eyes. A 10-chaetiger larva measures 740 µm long and 210 µm wide (Fig. 13.20C–D). In dorsal view, the prostomium is bluntly shaped and small enough so that the lateral lips of the vestibule can be seen beneath it. It bears three pairs of eyes, disposed as in Figure 13.20D. The prototroch is well developed and present as two distinct ciliary cells on each side of the developing nuchal ridge. Finely ciliated, oval nuchal organs and ciliated ridges whose function is unknown are present in that same area (Fig. 13.20D). Palps buds have developed. Provisional larval chaetae are present on all 10 chaetigers with the longest being notopodial; adult chaetae are not present. A lateral notopodial lobe is present on all chaetigers. Nototrochs begin on chaetiger 3 and continue posteriorly with each band consisting of 3–4 ciliated cells (Fig. 13.20D). Gastrotrochs occur on chaetigers 3, 5, 7 and 10 (Fig. 13.20C). The telotroch consists of five ciliated cells with a wide dorsal gap. The dorsal pigment pattern consists of granular black pigment distributed laterally between chaetigers 1 and 2. From chaetiger 4 posteriorly, dorsolateral “spots” of pigment occur in a triangular configuration at the anterior half of each segment. There is also russet pigment on the prostomium and pygidium, with some diffuse dark pigment also on the pygidium. In ventral view, the larva has a distinctive buccal region where the flexible lateral lips of the vestibule exhibit a deep median fold. There are long sensory cilia as well as nuchal cilia, and the prototroch continues ventrally as three cells of cilia on each side of the

$$ Reproductive Biology and Phylogeny of Annelida mouth. A small neurotroch extends ventrally to the ciliated pit in the middle of chaetiger 2 (Fig. 13.20C). Internally, there is a pharynx (which is partially eversible and has black pigment in its wall), a stomach intestine, and a proctodeum. As this larva develops in the plankton, the pigment intensifies with the smaller spots on the lateral borders forming lateral lines (both russet); black pigment of the pygidium intensifies until the telotroch cells stand out as clear areas. A 15-chaetiger larva is 1.1 mm long and 0.40 mm wide (Fig. 13.20E–F). Dorsally, the prostomium retains a blunt shape with the lateral lips of the vestibule still protruding beyond the lateral margins. Of the three pairs of eyes, the two lateral pairs are closely associated and appear to be merged (Fig. 13.20F). A single long sensory cilia is present lateral to each of the lateral-most eyes. The prototroch is well developed. The nuchal organs are prominent along the narrow caruncle that has developed between the prostomium and chaetiger 1. The shape of the nuchal ridge indicates that the earlier ciliated ridge is now part of the nuchal organs. The palps are longer, but are relatively short for this stage of development. Each palp is glandular in appearance. All chaetigerous segments still bear the larval chaetae, but in addition, chaetiger 5 bears internal modified chaetae that are of the characteristic form for this species. Bidentate neuropodial hooded hooks are present from chaetiger 7 (Fig. 13.20E). Nototrochs begin on chaetiger 3, continuing posteriorly and consisting of seven patches of cilia per band. Gastrotrochs occur on chaetigers 3, 5, 7, 10, and 15. The telotroch is unchanged, but the pygidium has developed a deep cleft. The dorsal pigment pattern has intensified so that granular black pigment now occurs lateral and intersegmental on chaetigers 1–7. Beginning with chaetiger 7, the posterior half of each segment has diffuse pigment that becomes intense at about chaetiger 11 (Fig. 13.20F). In addition, a double row of black lines along the anterior margin of each segment extends from chaetiger 5 posteriorly, fading into other pigmentation at chaetiger 10. An 18-chaetiger juvenile represents the stage just following settlement and metamorphosis (Fig. 13.20G). This specimen is 1.6 mm long and 0.4 mm wide. In dorsal view, the prostomium has elongated, causing the positions of the eyes and other features of the earlier stages to shift. There is only two pairs of eyes, with the small lateral pair of earlier stages now incorporated into the larger lateral pair. A well-developed caruncle has developed over chaetiger 1 and is bordered by prominent nuchal cilia (Fig. 13.20G). A small remnant of the prototroch is obscured by the elongate and forwardly directed palps. Parapodia have developed further, and all adult chaetae are now present, with the provisional chaetae having been shed. The modified chaetae of segment 5 have broken through the cuticle and are seen to have an elongate, bristle-topped beak (Fig. 13.20H). The nototrochs are present as seven distinct cells per segment, beginning on chaetiger 3, and represent the dorsal organs of the adult. A continuation of the nototrochs has become the branchial ciliation on chaetiger 7–10. The pygidium is 4-lobed as in the adult and, because of very fine pigmentation,

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Fig. 13.20. Larval development of Dipolydora brachycephala from Tomales Bay, California. A–B. Planktic 4-chaetiger larvae in ventral (A) and dorsal (B) views. C–D. Ten-chaetiger larvae in ventral (C) and dorsal (D) views. E–F. Late-stage 15-chaetiger planktic larvae in ventral (E) and dorsal (F) views. G. Benthic juvenile just following metamorphosis, showing anteriorly directed palps, caruncle, and branchiae from chaetiger 7. Remnants of the telotroch are present. H. Modified spines from chaetiger 5 from same juvenile (G). Original.

$& Reproductive Biology and Phylogeny of Annelida the lobes appear grey. The telotroch is also reduced, with only a few cilia remaining. The dorsal pigment has faded appreciably, leaving only the black intersegmental pigmentation and the diffuse granular pigment on the posterior half of posterior segments (Fig. 13.20G). The entire body has taken on a light brown cast, and russet pigment is still present on the margins of the prostomium and caruncle. In ventral view, the buccal region has progressed towards the adult morphology. With the exception of the prototroch, all of the ventral ciliation is still present, though somewhat reduced in the case of the neurotroch and ciliated pit. The internal structure is fully developed and pigmented: the vestibule is black and granular, the esophagus is unpigmented, the stomach is brown and fades into the clear intestine; and the proctodeum is dark russet. 13.6.4.1 Diversity of planktic larvae of the polydora complex A collage of polydorid larvae (Fig. 13.21) illustrates the wide variety of body shapes and pigment patterns that have been observed in planktic larvae of the Polydora complex (Blake 1966; unpublished; Blake and Woodwick 1975). Boccardia tricuspa (Hartman, 1939). This species is a relatively common shell and coralline borer in the eastern Pacific. Egg capsules were collected from coralline algae in July 1964 at Cayucos, California (Blake, unpublished). Eggs are pink and contained in small capsules that are joined beadlike in a string and attached to the inner lining of the burrow. All eggs are fertilized with larvae being released into the plankton at the 3-chaetiger stage. Larvae of B. tricuspa were collected from the plankton in Tomales Bay, California, in September–October 1971. A late-stage 18-chaetiger planktotrophic larva is shown in Figure 13.21A. This larva is thick and fusiform in shape, measuring 900 µm long and 300 µm wide. The shape of these larvae, together with the single dorsal row of chromatophores, is similar to that of larvae of other Boccardia species (example: B. columbiana, see above; B. proboscidea by Woodwick 1977; Blake and Kudenov 1981, and Gibson 1997). The prostomium is rounded on the anterior margin with lateral raised areas containing the prototroch; the anterior margin is pigmented a golden brown color. Palps arise from the posterior lateral margin of the peristomium; these are relatively long and extend posteriorly for about six segments. Each palp is densely covered with sensory cilia; the ventral groove is also heavily ciliated. There are three pairs of black eyes. The innermost pair is round in shape; the two outer pairs are often difficult to separate from one another, but one pair is cup-shaped and appears to overlie the third irregularly-shaped pair. The nuchal ridge or developing caruncle is thick and terminates bluntly on the anterior margin of chaetiger 2. Ciliated nuchal grooves are present on either side of the caruncle. Nototrochs begin on chaetiger 2 and continue on succeeding chaetigers to the end of the body. There are usually six patches of cilia per nototrochs; however, on chaetigers 7–11 that bear developing branchiae, there are eight patches of cilia. Dorsally the prototroch has three distinct ciliary patches,

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continuing to a ventrolateral location. The telotroch has prominent patches of cilia separated by a dorsal gap. Provisional chaetae are neither long nor serrated at this stage of development. Characteristic modified chaetae were observed in chaetiger 5, with the major spines consisting of a falcate spine and 1–2 spines with the tricuspid form that is typical for this species. The dorsal pigment pattern is dominated by a single row of large, branching black chromatophores from chaetiger 2. The pygidial segment contains golden brown pigment and isolated black pigmented patches. The entire dorsal surface of the larva is covered with numerous green-pigmented cells. The mouth region is complex. The maturation of the animal has caused migration of the ventral and lateral buccal lips. The lateral lips have shifted medially, and in an older specimen have fused to form the anterior boundary of the vestibule. The ventral lip has shifted anteriorly and the articulations that are characteristic of the adult are beginning to develop. There is no neurotroch, although the buccal area is covered with fine cilia that direct water currents and food into the vestibule. Gastrotrochs consist of eight patches of cilia and occur on chaetigers 7, 10, and 13. The ventral pigment of this species is distinctive; it forms a pattern of nine large cells on chaetigers 7–12 which looks like a random pattern, but which is actually exact in orientation for each specimen examined. It appears as dark red to black in transmitted light, but pale yellow to light green in reflected light. The only other pigmentation in this larva is a russet tinge in the prostomium, vestibule, and pygidium, and the pale red color of the esophagus. These large larvae appear to be ready for metamorphosis, but apparently delay this event due to lack of a suitable substrate. Carazziella califia Blake, 1979. Larvae of a species of Carazziella were encountered in plankton tows at Lawson’s Landing in Tomales Bay, California, on 1 November 1971 and again in July and August 1972. These larvae closely resemble several species of Boccardia, but the nature of the specialized chaetae of chaetiger 5 and the first appearance of hooded hooks on chaetiger 8 identify this as a species of Carazziella, most certainly C. califia described by Blake (1979) from Monterey Bay. Adults presumably also occur in the vicinity of Tomales and Bodega Bays. A 14-chaetiger planktotrophic larva is 950 µm long (Fig. 13.21B). The body of these larvae is thick and weakly fusiform. The prostomium is bluntly conical. There are four black eyes in a trapezoidal arrangement, with the central pair being located on the nuchal ridge or developing caruncle. Ciliated nuchal organs appear as flattened oval areas between the palp bases and the caruncle. Dorsally, the prototroch is located anterior to the palp insertion points, extending ventrally to merge with the cilia of vestibule. A short neurotroch extends from the mouth to anterior of a ciliated pit on chaetiger 2. Parapodia are well developed with the notopodia being the larger. Notochaetae are long serrated provisional chaetae while the neurochaetae are adult winged capillaries. Bidentate hooded hooks begin on chaetiger 8. Modified chaetae are present in chaetiger 5, including one spine with an expanded bristle-topped tip and another falcate spine also with bristles that

$  Reproductive Biology and Phylogeny of Annelida

Fig. 13.21. Larvae of six species of the Polydora-complex, all in dorsal view. A. Boccardia tricuspa, late-stage planktic larva from Tomales Bay, California (Original). Large fusiform shape and dorsal row of chromatophores together with tricuspid spines from chaetiger 5 (not shown) are diagnostic. B. Carazziella califia, late-stage planktic larva from Tomales Bay, California. After Blake, J.A. and Arnofsky, P.L. 1999. Hydrobiologia 402: 57–106, Fig. 12A. Similar to Boccardia species, but shape is more slender and modified chaetae of chaetiger 5 include two types of bristle-topped spines. C. Late-stage larvae of Dipolydora concharum from New England. After Blake, J.A. 1969. Ophelia 7: 1-63, Fig. 25A. Larvae are slender with dorsal pigment including a central group of pigmented cells and transverse bands on the anterior margin of most chaetigers. D. Dipolydora quadrilobata Type II 9-chaetiger planktic larvae from New England. After Blake, J.A. 1969. Ophelia 7: 1-63, Fig. 35B. Dorsal pigment consists of a pair of curved melanophores on each segment from chaetiger 4 and a wedge of black pigment on the pygidium. E. Polydora websteri, 12-chaetiger larva from New England. After Blake, J.A. 1969. Ophelia 7: 1-63, Fig. 8. Classic pigment pattern for this and related species includes transverse bands of pigment across anterior chaetigers and paired chromatophores on middle and posterior chaetigers. F. Pseudopolydora paucibranchiata, 13-chaetiger planktic larva from California. After Blake, J.A. and Woodwick, K.H. 1975. Biological Bulletin 149: 109–127, Fig. 18. The shape is distinctly fusiform and the dorsal pigment pattern consists of a medial chromatophore on chaetiger 1, followed on subsequent chaetigers by lateral chromatophores and medial reticulated green pigment.

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cloak the tip. Developing branchiae are present on chaetigers 7–9. Nototrochs are present from chaetiger 3, continuing posteriorly as seven ciliary patches per segment, although those of chaetiger 3 appear to be regressing. Gastrotrochs consist of five ciliated patches on chaetigers 3, 5, 7, 10, and 13. The dorsal pigment pattern consists of a single row of chromatophores from chaetiger 4, continuing posteriorly. Additional pigment is granular russet (reflective yellow) on the posterior borders of the pygidium. There is no ventral pigment. The pygidium has a median cleft; the telotroch is prominent. The gut consists of a black pharynx, followed by a clear esophagus, stomach-intestine with oil globules, and noticeable segmental diverticulae, and a clear proctodeum. This larva was capable of extruding part of its pharyngeal region, and increased its use of this “proboscis” when placed on a glass slide. Dipolydora concharum (Verrill, 1880). This species is a common shellborer in New England waters. In studies by Blake (1969, 1971), the species (as Polydora concharum) was collected from shells of the sea scallop Placopecten magellanicus. Blake (1969) described the larval development from the Damariscotta River Estuary, Maine. The species produces egg capsules in January and February; planktic larvae occurred from February– April. The 15-chaetiger larva is 900 µm long and has an elongated, slender shape (Fig. 13.21C). The dorsal pigment pattern is diagnostic for this species. Dorsal bands begin on chaetiger 2 and continue to the end of the body; these are separated into two medial bands and two lateral spots. A medial group of pigmented cells is present dorsally and these occur on all segments from chaetiger 3 (Fig. 13.21C). Dorsal pygidial pigment consists of two large and one small patch that form a dark triangle. Additional black pigment occurs dorsolaterally on the head and margins of the first chaetigers. Nototrochs occur on chaetiger 3 and subsequent segments. Each row has four patches of cilia. Gastrotrochs consisting of four patches each occurs on chaetigers 3, 5, and 7. Palps are short and inconspicuous. Two patches of nuchal cilia are present on the dorsal side of the head. Adult chaetae are developing. The modified chaetae of segment 5 consist of a heavy curved spine and a limbate chaeta. Hooded hooks are present from chaetiger 7. Dipolydora quadrilobata (Jacobi, 1883)—Type II. Blake (1969) collected two distinct types of egg capsules and early larvae from different locations on the Maine coast and described the larval development of Dipolydora quadrilobata (as Polydora). Developmental Type I collected from Cobscook Bay in northern Maine consisted of fully brooded larvae that carried out their entire development in egg capsules sustained by unfertilized nurse eggs. The developing larval stages did not develop provisional chaetae typical for larvae that become planktic. Developmental Type II collected from Lamoine Beach in central Maine; included egg capsules contained developing larvae without nurse eggs. Type II larvae developed to a 4–5 chaetiger stage and then were released into the water where they developed as typical planktotrophic spionid larvae. The

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Reproductive Biology and Phylogeny of Annelida

presence of two distinct types of larval development is another example of poecilogony. A 9-chaetiger Type II larva measuring 900 µm long is shown in Figure 13.21D. These larvae are thick and slightly fusiform in shape. They have a distinctive dorsal and pygidial pigment pattern, and a dark green intestine. Provisional larval chaetae have a single row of serrations. Adult chaetae are developing but are obscured by the larval chaetae. Ciliation consists of a prototroch that is divided ventrally into two portions and dorsally extends to about the level of the lateral eye spots. A sensory cilium arises ventrally from the prototroch at about the level of the palps. Sensory cilia occur anterior to the inner pair of circular eyes. Long tactile cilia arise from the region of the lateral eyes and extend well beyond the head in some individuals. The neurotroch extends into chaetiger 2 and terminates in a small ciliated pit. Nototrochs occur from chaetiger 3. Gastrotrochs occur ventrally on chaetigers 3, 5, 7, 10, 13–15, consisting of four patches of cilia. The telotroch consists of five patches of cilia with a dorsal gap. Polydora websteri Hartman, 1943. This species is a widespread shell borer that has often been recorded as a pest of commercial shellfish. Blake (1969) described larvae of P. websteri from the Damariscotta Estuary in Maine. Egg capsules were obtained from March–July, with planktic larvae present from April–September over a 3-yr period from 1966–1968. A 12chaetiger larva measures 740 µm and a 17-chaetiger larva 1,300 µm (Fig. 13.21E). The overall shape of these larvae is slender, but thicker than Dipolydora concharum. The prostomium is rounded anteriorly and has yellow-brown granular pigment. The lateral eye spots are so branched on some specimens that they cover large areas of the prostomium (Fig. 13.21E). Four large tactile cilia originate from the eyes. Nuchal ciliation occurs alongside the caruncle, which extends to the anterior margins of chaetiger 1. Provisional chaetae are present on all segments except chaetiger 5 where modified chaetae are present. Gastrotrochs are reduced on chaetigers 3 and 5, but are present on segments 7, 10, and 13. Nototrochs are present from chaetiger 3. The telotroch exhibits a wide dorsal gap. The dorsal pigment pattern consists of two rows of black chromatophores from chaetiger 3 with those of 3–6 being band-shaped and the remaining ones branched. Pseudopolydora paucibranchiata (Okuda, 1937). This species is widely distributed in the north Pacific and is common in estuaries of central California. Blake and Woodwick (1975) described the reproduction and larval development of P. paucibranchiata in detail. The species has a typical early development where embryos are retained in egg capsules in the tube of females until they are released into the plankton with three fully developed chaetigers. There are no nurse eggs in the capsules (Blake and Woodwick, 1975). A planktic 13-chaetiger larva in dorsal view is depicted in Fig. 13.21F. This larva is 640 µm long and distinctly fusiform in shape. These larvae have a characteristic swimming behavior, with the posterior end held lower than the prostomium and the elongate palps held posteroventrally but may be recurved with tips directed anteriorly. The

Spionida

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head region is enlarged due to the broad prostomium and expanded lateral lips of the vestibule. Palps arise from the dorsolateral region of the peristomium; each has a ventral ciliated groove. Three pairs of black eyes are present in more or less a straight line; the two inner pairs are circular and the larger, lateral pair is cup-shaped. A nuchal ridge or developing caruncle is present, with patches of rapidly beating nuchal cilia on either side. The prototroch is well developed with several closely associated long tactile cilia. Nototrochs are present from chaetiger 3, each with 4–6 patches of cilia. Gastrotrochs occur on chaetigers 3, 5, 7, and 11 with four patches of cilia on chaetiger 3 and six on chaetigers 5, 7, and 11. The telotroch encircles the anal segment, but there is a dorsal gap. A neurotroch extends halfway into chaetiger 1; several small patches of cilia are present lateral to the neurotroch on the ventral side of chaetiger 1. The pigment pattern is highly characteristic. The dorsal surface of chaetigers 2–11 is covered with a reticulate green pigment. Black pigment is heavy on the ventral side of the peristomium. Light brown, non-reflective pigment occurs on the anal segment and on the margins of the peristomium and lips of the vestibule. The cilia of the telotroch occur in unpigmented areas. Two medial chromatophores occur ventrally; one is on or about chaetiger 6 and the other is on the anal segment; the former is black while the latter is iridescent yellow by reflected light. Two prominent areas of reflective yellow pigment occur dorsolaterally on the peristomium. Similar reflective pigment occurs dorsally with the lateral chromatophores from chaetigers 3–10.

13.6.5 Direct Development Pygospiopsis dubia (Monro, 1930). There have been few studies where direct development has actually been reported for spioniform polychaetes. Studies of development refer to either (1) demersal spawning with ultimate development of a planktic larva, (2) brooding in egg capsules or cocoons where larvae are released to the plankton at various stages of development, or (3) viviparity where embryos brooded on special structures on the body are eventually released as planktic larvae or postlarval stages. The latter situation occurs in some populations of Streblospio benedicti and S. shrubsoli and approximates direct development (Blake and Arnofsky 1999; this paper). Söderström (1920) reported that larvae of Boccardia natrix from Patagonia and sub-Antarctic localities were brooded in epitokous segments of females and suggested that the development was direct (see Blake 1983). Buzhinskaja and Jorgensen (1997) reported that larvae of Trochochaeta carica from the Kara Sea were entirely brooded within the tube. During a cruise to the eastern side of the Antarctic Peninsula in May 2000 to study marine sediments formerly covered by the Larsen Ice Shelf, larval, postlarval, and adult specimens of Pygospiopsis dubia were recovered from a single benthic sample from a water depth of ca. 500 m just south of the Prince Gustav Channel. The species appears to exhibit direct development. The morphology of P. dubia is unusual among spioniforms. As noted in the earlier phylogenetic review, the genus appears to be isolated within the

$ " Reproductive Biology and Phylogeny of Annelida Spionidae and is perhaps a sister group to the two larger clades, Spioninae and Nerininae, that have been identified. I postulate that this species is brooded in the tubes of females. Whether or not egg capsules are present is unknown. The earliest stage of development recovered from the sample was a 14-chaetiger non-planktic larva (Fig. 13.22A). This specimen measures 780 µm long and 270 µm wide. It is short, thick, and does not appear capable of swimming in the plankton. A pair of very short palps is present. The prostomium is short and divided into two lobes that extend over the developing mouth; ventrally there are two lips. Both noto- and neurochaetae are present; they are not serrated as in provisional chaetae of planktic spionid larvae. Nototrochs are absent. Gastrotrochs are present from about chaetiger 3, but the distribution of these is not clear; a telotroch is present. Nototrochs are absent. These larvae should be capable of slow crawling or swimming movements within the tube. A 16-chaetiger post larva is shown in Figure 13.22B. This specimen is 1,165 µm long and 390 µm wide. The prostomium is very blunt, and similar to that of the adult; a single intact palp is considerably longer than in the previous stage, but still shorter than in other spionids at this stage of development. Two pairs of short and blunt branchiae are present on chaetigers 7–8. There is no evidence of dorsal ciliary organs. Evidently, the broad lamellate adult branchiae and the dorsal cilia develop later. Figure 13.22C is of the anterior end of one of the adult females recovered from the sample. These specimens exhibit a short, blunt prostomium with an occipital tentacle and short caruncle, elongate dorsal cirri on chaetigers 2–3, and elaborate branchiae from chaetiger 7 that are fused with dorsal lamellae. There have been very few observations of early developmental stages of spioniforms from polar and deep-water habitats where we would expect to find evidence of direct development. In addition to the present report on Pygospiopsis dubia from Antarctica and that of Buzhinskaja and Jorgensen (1997) on Trochochaeta carica from the Kara Sea, a few specimens of a related species of Pygospiopsis (P. occipitalis Blake) from off California in 1200 m suggest a similar mode of direct development (Blake, unpublished). Detection of such larval forms requires the use of gentle elutriation methods and very fine sieves to extract these delicate life stages from the sediments. Care must also be taken to avoid extreme temperature changes and the samples must be placed on ice or in a refrigerator as soon as possible. These methods were used in the collection of the P. dubia described here.

13.6.6 Poecilogony and Adelphophagia Some spionid polychaetes are able to vary their pattern of larval development by having (1) populations that have short periods of brooding followed by long periods of planktotrophicc development, or (2) populations that have long periods of brooding where developing larvae

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Fig. 13.22. Development of Pygospiopsis dubia from the Weddell Sea sector of the Antarctic Peninsula. A. Scanning electron micrograph (SEM) of 14-chaetiger benthic larva in lateral view. Chaetae present on this specimen are non-serrated, suggesting direct development to adult. B. SEM of 16-setiger postlarva showing development of palps and of branchiae on chaetigers 7–8. C. SEM of anterior end of adult female in dorsal view taken from the same samples as the larvae and post-larvae. Original.

$ $ Reproductive Biology and Phylogeny of Annelida feed on unfertilized eggs (adelphophagia) followed by abbreviated periods in the plankton or none at all. In a few cases, asexual reproduction occurs as an additional form of reproduction. Species exhibiting such variability in their mode of larval development throughout their geographic range are said to exhibit poecilogony. Blake and Arnofsky (1999), who reported on eight spionid species having poecilogony, reviewed this subject: Boccardia proboscidea, Dipolydora quadrilobata, Pygospio elegans, Pseudopolydora kempi, Spio decoratus, S. martinensis, S. setosa, and Streblospio benedicti. Except for S. benedicti, all of these species have populations that have been reported with both the long planktotrophic larval development and adelphophagic development. Not all species exhibiting adelphophagia have been reported to also have non-adelphophagic populations. The different forms of development of S. benedicti and P. elegans were discussed in earlier sections (see above). The non-fertilized eggs of adelphophagic populations are called nurse eggs and two distinct types have been reported. In Dipolydora quadrilobata, Polydora hoplura, P. nuchalis, and Boccardia proboscidea, the form of the unfertilized eggs appears to be identical to that of fertilized eggs suggesting that either a shortage of sperm or gamete incompatibility led to nonfertilization; they are engulfed whole by developing larvae (Wilson 1928; Woodwick 1960, 1977; Blake 1969), In contrast, unfertilized eggs of Pygospio elegans and Pseudopolydora kempi are fragile, readily breaking up into small yolk granules that are devoured by developing larvae (Rasmussen 1973; Blake and Woodwick 1975). Nurse egg formation for Amphipolydora vestalis reported by Gibson and Paterson (2003) appears to be similar to that of P. kempi and Pygospio elegans. These authors reported that eggs destined to be nurse eggs cleaved unequally forming loose balls of blastomeres that subsequently broke apart into individual blastomeres that were then consumed by developing embryos. Adelphophagia probably occurs in at least half of the species of the Polydora- complex and related genera studied to date (Blake and Arnofsky, 1999). Adelphophagia has been considered a form of lecithotrophic development because the adult simply deposits yolk into nurse eggs instead of into the cytoplasm of normally developing oocytes (Woodwick 1977; Blake and Kudenov 1981). Radashevsky (1994) distinguished between different forms of lecithotrophy and introduced the terms exolecithotrophic for species with nurse eggs and adelphophagia, and endolecithotrophic for species that store yolk in normally developing eggs but do not have planktotrophic larvae. There is evidence that some species of Polydora may be in the incipient stages of establishing poecilogony in their populations. For example, Blake (1969) reported that P. cornuta (as P. ligni), which was normally observed to have all eggs in its capsules developing into larvae, sometimes had individual capsules with unfertilized eggs. In this example which was near the end of the breeding season, two of 11 capsules in a tube had unfertilized eggs and the developing larvae fed on them. Additional examples for P. cornuta and similar examples in P. websteri have been observed

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subsequently (Blake unpublished). Radashevsky and Cárdenas (2004) reported that 10% of the capsules of P. rickettsi contained unfertilized eggs that were consumed by developing larvae. These observations suggest a mechanism whereby adelphophagia could have evolved. Initially, a failure of fertilization or depletion of sperm in the seminal receptacles would result in some eggs being non-fertilized and available as an extrinsic yolk source. This seems very reasonable in a polytelic species such as P. cornuta where a single female is capable of rapidly producing multiple broods in a single summer season. Near the end of a breeding season, females might be depleted of sperm and produce more capsules with fewer fertilized eggs. Such a pattern could be selected if the larvae being brooded for longer periods required less time in the plankton when required phytoplankton species were less abundant. The shift to a permanent adelphophagic mode of development and the eventual evolution of a completely different type of nurse egg such as found in Pygospio elegans and Pseudopolydora kempi would require a modification of oogenesis to produce eggs that would be completely non-viable. An intermediate stage in this process appears to be found in Boccardia proboscidea. Blake and Kudenov (1981) and Gibson (1997) observed that Boccardia proboscidea within the same population was capable of producing planktotrophic larvae only after variable periods of being lecithotrophic. At the same time, other larvae are produced that develop entirely lecithotrophic in egg capsules, surviving on nurse eggs. Gibson and Gibson (2004) performed a series of experiments with Boccardia proboscidea to determine if morphogenesis differed among the different modes of reproduction. These authors determined that when embryos fed on nurse eggs, offspring with an accelerated onset of juvenile traits were produced relative to offspring that were planktotrophic. Some offspring that ate nurse eggs had accelerated development, hatched as benthic juveniles, and thus appeared to be morphologically preadapted for a benthic lifestyle. In contrast, larvae lacking nurse eggs hatched as planktotrophic larvae, spending 15 d or more in the plankton before developing juvenile characteristics and settling into the sediment/benthos. Based on their results, Gibson and Gibson (2004) proposed that poecilogony in B. proboscidea evolved through sequential heterochrony with accelerated morphogenesis of juvenile traits. B. proboscidea, therefore, maintains local populations by early onset of juvenile morphology in adelphophagic larvae. At the same time planktotrophic larvae in the same population or others, disperse the species over a greater geographic range. Many species probably shift seasonally between intrinsic and extrinsic yolk production and this shift might be related to seasonal flux of organic matter available as phytoplankton. The relationship of spionid reproduction to phytoplankton cycles and organic flux has not been investigated, but is probably required before the evolution and ecology of the reproductive plasticity exhibited by spionids will be fully understood. It is possible that some reported examples of poecilogony in spionids may actually be examples of sibling species. Morgan et al. (1997) validated

$ & Reproductive Biology and Phylogeny of Annelida poecilogony in Pygospio elegans from populations in England and France by comparing genetic divergence in 14 enzymatic loci. No fixed genetic differences were detected. Other examples of apparent poecilogony will need to be verified by similar techniques.

13.7 ASEXUAL REPRODUCTION Two types of asexual reproduction occur in spioniforms: architomy and paratomy (Table 13.1). Architomy is the simplest form of asexual reproduction and includes fragmentation of the body into individual segments or groups of segments, which then regenerate into new individuals (Fig. 13.23A). Paratomy involves the division of the body into two distinct halves, with the reconstitution of missing parts by regeneration. Sometimes the second half (stolon) remains attached to the first half (stock) while regenerating. Additional divisions may also occur, resulting in chains of stolons being proliferated from the original stock parent. Spioniforms exhibiting paratomy tend to be very small, usually with a reduced and defined number of segments, whereas species having architomy are larger and have numerous segments. To date, architomic asexual reproduction in spioniforms appears to be restricted to one chaetopterid species and several species of the Spioninae. Architomy has been described for Phyllochaetopterus prolifica (Potts, 1914). The worms autotomize into anterior and posterior parts that in turn regenerate the missing parts. Potts found up to six individuals in a single tube. It is likely that new individuals resulting from fragmentation cut open the tube and produce side branches. Architomy has been reported in the laboratory for Dipolydora caulleryi and D. socialis by Stock (1964), but has not been observed in the field. Pygospio elegans has been widely reported as having architomy (Rasmussen 1953, 1973; Bregenballe 1961; Muus 1967; Hobson and Green 1968; Armitage 1979; Wilson 1983; Anger 1984; Gibson and Harvey 2000; Golam 2004). Architomy also occurs in the closely related species P. californica (Blake, unpublished). Blake (1983) reported architomy for Amphipolydora abranchiata, from off Argentina in 100 m. Gibson and Paterson (2003) report asexual fragmentation in A. vestalis from New Zealand. Radashevsky and Nogueira (2003) described architomic fragmentation in Dipolydora armata. In all of these accounts, architomic fragmentation results in a worm breaking up into four or more separate parts, each of which regenerates missing anterior and posterior ends. In Pygospio elegans the parent body is split into fragments through transverse fission. Each fragment regenerates into a separate individual. Gibson and Harvey (2000) described details of morphogenesis during postfission regeneration. These authors found that each fragment retained the original anterior-posterior polarity and that regeneration followed a defined sequence of events over an 8 d period. These included wound healing (1 d), development of a blastema to regenerate lost tissues and body

Variable Variable

Variable

Architomy Architomy

Architomy

Architomy

Paratomy

Paratomy

Paratomy

Paratomy

Paratomy

Paratomy

Pygospio californica Amphipolydora abranchiata Amphipolydora vestalis

Dipolydora armata

Polydora tetrabranchia

Polydorella prolifera

Polydorella stolonifera

Polydorella kamakamai

Polydorella smurovi

Polydorella dawydoffi

Between segments 15–16, 16-17, or 17–18 Between segments 10–11 Between segments 10–11 Between segments 10-11 Between segments 10–11 Between segments 11–12

Variable

N/A N/A

Variable Variable

Phyllochaetopterus prolifica Architomy Pygospio elegans Architomy

>5

1

3

1

1

1-2

N/A

N/A

N/A N/A

Number of stolons

Table 13.1. Asexual reproduction in spioniform polychaetes Species Type Location of fission zone

?

Yes

Yes

?

?

Yes

Yes

Yes

Yes ?

? Yes

Sexual reproduction

South China Sea, Vietnam

Red Sea

Southeastern Australia Philippines

Western Australia

North Island, New Zealand Widespread in tropical and subtropical seas North Carolina

California Argentina

Eastern Pacific Northern Europe; Massachusetts; California; Washington

Locality

Radashevsky, 1996

Tzetlin et al. 1985

Williams, 2004

Blake and Kudenov, 1978

Augener, 1914; Blake and Kudenov, 1978

Campbell, 1955

Radeshevsky and Nogueira, 2003

Gibson and Paterson, 2003

Potts, 1914 Rasmussen, 1953; 1973; Hobson and Green, 1968; Armitage, 1979; Wilson, 1983; Anger, 1984; Gibson and Harvey, 2000; Golam, 2004 Blake, unpublished Blake, 1983

References

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$! Reproductive Biology and Phylogeny of Annelida

Fig. 13.23. Asexual reproduction. A. Architomic regeneration of Pygospio elegans. Rasmussen, E. 1953. Nature 171: 1161, unnumbered figure. B–C. Paratomic asexual reproduction of Polydorella kamakamai. After Williams, J.D. 2004. Journal of Natural History 38: 1339–1358, Fig. 5A–E, showing pattern of stolonization and regeneration.

regions (2–3 d), segmentation (3–6 d), and differentiation of regenerated segments into specific structures such as palps and pygidial cirri (4–8 d). Gibson and Harvey (2000) found this sequence of regeneration and differentiation to be the same regardless of where the original fragmentation took place. Fragments having the original head had a higher survivorship than fragments containing the original posterior end. For Amphipolydora vestalis, Gibson and Paterson (2003) reported that 4–6 fragments from a single parent regenerate their bodies within eight days. In most populations of Pygospio elegans that have been studied, both sexual and asexual reproduction occurs. This strategy ensures that once colonized by settling larvae, populations could be expanded and maintained asexually. Armitage (1979), working with populations from two different localities in Tomales Bay, California, found that both sexual and

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asexual phases of P. elegans were controlled primarily by temperature, in that both forms of reproduction were accelerated following seasonal lows. These results support earlier observations by Rasmussen (1953) from Denmark that asexual reproduction in P. elegans increased with increasing spring temperatures. Armitage (1979) also observed that sexual reproduction was more prevalent in an intertidal sand flat with a gradual slope and homogenous sediments rather than at a second site that was more heterogeneous. In contrast, Anger (1984) working with North Sea and Baltic Sea populations was unable to correlate either temperature or salinity with reproduction. She also found that a western Baltic population favored asexual reproduction exclusively, whereas eastern Baltic and North Sea populations exhibited both sexual and asexual reproduction. The differences between the results of Armitage (1979) and Anger (1984) are probably an artifact of differences in approach to data collection. Armitage conducted a long-term field investigation, whereas Anger maintained specimens in static laboratory conditions. Paratomy has been reported for Polydora tetrabranchia and five closely related species of Polydorella: P. prolifera, P. stolonifera, P. smurovi, P. dawydoffi, and P. kamakamai (Campbell 1955; Blake and Kudenov 1978; Tzetlin et al. 1985; Radashevsky 1996; Williams 2004). Polydora tetrabranchia is a shell borer, whereas the five Polydorella species construct tubes on the surface of sponges. According to Campbell (1955), asexual reproduction in Polydora tetrabranchia occurs by transverse fission of the stock animal. Regeneration of new posterior and anterior ends proceeds while the separated sections (stolons) are still connected, providing an appearance of two joined individuals. A chain of three individuals was found in a laboratory experiment, but no more than two joined individuals were ever observed in the field. Asexual reproduction proceeded year round and approximately one-third of all specimens collected were regenerating anterior or posterior ends. Radashevsky (1996) and Williams (2004) reviewed paratomy in Polydorella species. In P. prolifera, P. stolonifera, P. kamakamai, and P. smurovi, the fission and growth zone occurs between segments 10 and 11 (Fig. 13.23B–F); whereas, the growth zone appears between segments 11 and 12 in P. dawydoffi. The first three species have 15 segments; the latter two species have 16. Radashevsky (1996) has reported chains of 5–6 stolons for P. dawydoffi. In P. stolonifera, regeneration of a stolon begins with the development of a new anterior end with small palp buds that appear in a growth zone between chaetigers 10 and 11. Eventually the section of the worm anterior to the growth zone breaks away and regenerates a new posterior end, while the stolon differentiates into a fully functional and normal appearing individual (Blake and Kudenov 1978). A similar pattern occurs in the other species (see Williams 2004). In P. kamakamai a chain of three stolons develops from the stock animal. The third stolon subsequently develops a secondary stolon with continued regeneration (Fig. 13.23B–C).

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Reproductive Biology and Phylogeny of Annelida

Sexual reproduction has been reported for P. smurovi and P. kamakamai but is likely to occur in all four species because a dispersive larval stage would be needed to colonize new sponges. Williams (2004), however, speculated that in lieu of sexually produced larval stages, adults of Polydorella species might leave their burrows and move to adjacent sponges, presumably by swimming or drifting. Such waterborne movement of adults has been observed for benthic infaunal polychaetes (Dauer et al. 1982), but has not been documented for epifaunal species.

13.8 ACKNOWLEDGMENTS Research on California polychaete larvae cited in this chapter was supported by the National Science Foundation under Grant OCE-71-00497A02 to James A. Blake while he was on the faculty of the Pacific Marine Station, University of the Pacific at Dillon Beach, California. Completion of this review was supported by NSF Grant No. DEB-0118693 (PEET) to James A. Blake, University of Massachusetts, Boston. Jason Williams prepared the SEMs of Pygospiopsis and provided some of his images of Polydorella for one of the figures. I am grateful to Greg Rouse for inviting me to contribute this chapter and to Nancy Maciolek for her careful review of the manuscript and helping with the graphics.

13.9 LITERATURE CITED Allen, E. J. 1904. The anatomy of Poecilochaetus Claparède. Quarterly Journal of Microscopical Science, new series 48: 79–151. Anger, V. 1984. Reproduction in Pygospio elegans (Spionidae) in relation to its geographical origin and to environmental conditions: a preliminary report. In Fischer, A. and Pfannenstiel, H. -D. (eds), Polychaete Reproduction: Progress in Comparative Reproductive Biology. Fortschritte der Zoologie 29: 45–51. Armitage, D. L. 1979. The Ecology and Reproductive Cycle of Pygospio elegans Claparède (Polychaeta: Spionidae) from Tomales Bay, California. M.S. Thesis, University of the Pacific, Stockton, California. 81 pp. Baccetti, B. 1979. The evolution of the acrosomal complex. Pp. 305–329. In Fawcett, D. W. and Bedford, J. M. (eds), The Spermatozoon: Maturation, Motility, Surface Properties, and Comparative Aspects. Urban and Schwarzenberg, Baltimore. Bhaud, M. 1966. Étude du développement et de l’écologie de quelques larves de Chaetopteridae. Vie et Milieu 17: 1087–1120. Bhaud, M. 1978. Morphological variations of the modified chaetae of chaetopterids during ontogenesis. Ophelia 17: 199–206. Bhaud, M. and C. Cazaux. 1987. Description and identification of polychaete larvae: their implications in current biological problems. Oceanis 13: 597–753. Blake, J. A. 1965. Spionid polychaetes from Morro Bay, California: A Taxonomic and Biological Study. M. A. Thesis, California State University, Fresno, Fresno, California. 55 pp., 10 pls. Blake, J. A. 1969. Reproduction and larval development of Polydora from northern New England (Polychaeta: Spionidae). Ophelia 7: 1–63. Blake, J. A. 1971. Revision of the genus Polydora from the east coast of North America (Polychaeta, Spionidae). Smithsonian Contributions to Zoology 75:1–32.

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Blake, J. A. 1979. Four new species of Carazziella (Polychaeta: Spionidae) from North and South America, with a redescription of two previously described forms. Proceedings of the Biological Society of Washington 92: 466–481. Blake, J. A. 1983. Polychaetes of the family Spionidae from South America, Antarctica and adjacent seas and islands. Biology of Antarctic Seas XIV. Antarctic Research Series 39: 205–288. Blake, J. A. 1996. Chapter 4. Family Spionidae. Pp. 81–223. In Blake, J.A., Hilbig, B., and Scott, P.H. (eds). Taxonomic Atlas of the Santa Maria Basin and Western Santa Barbara Channel. Vol. 6. Annelida Part 3. Polychaeta: Orbiniidae to Cossuridae. Santa Barbara Museum of Natural History. Blake, J. A. and Arnofsky, P. L. 1999. Reproduction and larval development of the spioniform Polychaeta with application to systematics and phylogeny. Hydrobiologia 402: 57–106. Blake, J. A. and Arnofsky, P. L. 2000. Systematics and phylogeny of the spioniform Polychaeta. Bulletin of Marine Science 67: 657 (abstract). Blake, J. A. and Kudenov, J. D. 1978. The Spionidae (Polychaeta) from southeastern Australia and adjacent areas, with a revision of the genera. Memoirs of the National Museum of Victoria 39: 171–280. Blake, J. A. and Kudenov, J. D. 1981. Larval development, larval nutrition, and growth for two Boccardia species (Polychaeta: Spionidae) from Victoria, Australia. Marine Ecology Progress Series 6: 175–182. Blake, J. A. and Woodwick, K. H. 1975. Reproduction and larval development of Pseudopolydora paucibranchiata (Okuda) and Pseudopolydora kempi (Southern) (Polychaeta: Spionidae). Biological Bulletin 149: 109–127. Bochert, R. 1996a. An electron microscopic study of oogenesis in Marenzelleria viridis (Verrill, 1873) (Polychaeta: Spionidae) with special reference of large cortical alveoli. Invertebrate Reproduction and Development 29: 57–69. Bochert, R. 1996b. An electron microscopic study of spermatogenesis in Marenzelleria viridis (Verrill, 1873) (Polychaeta: Spionidae). Acta Zoologica 77: 191–199. Bochert, R and Bick, A. 1995. Reproduction and development of Marenzelleria viridis (Polychaeta: Spionidae). Marine Biology 123: 763–773. Bolam, S. F. 2004. Population structure and reproductive biology of Pygospio elegans (Polychaeta: Spionidae) on an intertidal sandflat, Firth of Forth, Scotland. Invertebrate Biology 123: 260–268. Borowski, C. 1995. New records of Longosomatidae (Heterospionidae) (Annelida, Polychaeta) from the abyssal southeast Pacific, with description of Heterospio peruana sp. n. and general remarks on the family. Mitteilungen aus dem Hamburgischen Zoologischen Museum und Institut 92, Ergänzungsband 1: 129–144. Bregenballe, F. 1961. Plaice and flounder as consumers of the microscopic bottom fauna. Meddelelser fra Kommissionen for Danmarks Fiskeri- og Havunderersøgelser, new series 3(6): 133–182. Buzhinskaja, G. N. and Jørgensen, L. L. 1997. Redescription of Trochochaeta carica (Birula, 1897) (Polychaeta, Trochochaetidae) with notes on its reproductive biology and larvae. Sarsia 82: 69–75. Campbell, M. A. 1955. Asexual Reproduction and Larval Development in Polydora tetrabranchia Hartman. Ph.D. Dissertation, Duke University. 67 pp. + 35 pls. Cazaux, C. 1965. Développment larvaire de Chaetopterus variopedatus (Renier). Travaux de l’Institut de Biologie Marine de l’Université de Bordeaux 102 (1A): 1–31. Cazaux, C. 1985. Reproduction et développement larvaire de l’annélide polychète saumâtre Streblospio shrubsolii (Buchanan, 1890). Cahiers de Biologie Marine 26: 207–221.

$!" Reproductive Biology and Phylogeny of Annelida Collier, M. and M. L. Jones. 1967. Observations on the reproductive and general morphology of Streblospio benedicti Webster. Biological Bulletin 133: 462. Costello, D. P., Davidson, M. E., Eggers, A., Fox, M. H., and Henley, C. 1957. Methods for Obtaining and Handling Marine Eggs and Embryos. Marine Biological Laboratory, Woods Hole, Massachusetts. 247 pp. Dauer, D. M., Ewing, R. M. Sourbeer, J. W., Harlan, W. T., and Stokes, Jr. T. L. 1982. Nocturnal movements of the macrobenthos of the Lafayette River, Virginia. International Revue Der Gesamten Hydrobiologie 67: 761–775. Dean, D. 1965. On the reproduction and larval development of Streblospio benedicti. Biological Bulletin 128:67–76. Dorsett, D. A. 1961. The reproduction and maintenance of Polydora ciliata (Johnst.) at Whitstable. Journal of the Marine Biological Association of the United Kingdom 41: 383–396. Eckberg, W. R. and S. D. Hill. Aug. 9, 1996. Chaetopterus—oocyte maturation, early development, and regeneration. Marine Models Electronic Record. [serial online]. Available from http://www.mbl.edu/BiologicalBulletin/MMER/ECK/ EckTit.html Eckelbarger, K. J. 1980. An ultrastructural study of oogenesis in Streblospio benedicti (Spionidae), with remarks on diversity of vitellogenic mechanisms in Polychaeta. Zoomorphologie 94: 241–263. Eckelbarger, K. J. 1983. Evolutionary radiation in polychaete ovaries and vitellogenic mechanisms: their possible role in life history patterns. Canadian Journal of Zoology 61(3): 487–504. Eckelbarger, K. J. 1984. Comparative aspects of oogenesis in polychaetes. In Fischer, A. and Pfannensteil, H. -D. (eds), Polychaete Reproduction: Progress in Comparative Reproductive Biology. Fortschritte der Zoologie: 29: 123–148. Eckelbarger, K. J. 1986. Vitellogenic mechanisms and the allocation of energy to offspring in polychaetes. Bulletin of Marine Science 39 (2): 426–443. Eckelbarger, K. J. 1988. Oogenesis and the Female Gametes. In Westheide, W. and Hermans, C. O. (eds), The Ultrastructure of Polychaeta. Microfauna Marina 4: 281– 307. Eckelbarger, K. J. 1992. Polychaeta: Oogenesis. Pp.109–127. In Harrison, F.W. and Gardiner, S. L. (eds), Microscopic Anatomy of Invertebrates, Vol. 7: Annelida. WileyLiss, Inc. Eckelbarger, K. J. 1994. Diversity of metazoan ovaries and vitellogenic mechanisms: Implications for life history theory. Proceedings of the Biological Society of Washington. 107: 193–218. Eckelbarger, K. J. and J. P. Grassle. 1983. Ultrastructural differences in the eggs and ovarian follicle cells of Capitella (Polychaeta) sibling species. Biological Bulletin 165: 379–393. Enders, H. E. 1909. A study of the life history and habits of Chaetopterus variopedatus. Journal of Morphology 20: 479–532. Fonseca-Genevois, V. da and C. Cazaux. 1987. Streblospio benedicti Webster, 1879 (Annèlide Polychéte) dans l’estuaire de la Loire: biologie et écologie. Cahiers de Biologie Marine 28: 231–261. Franzén, Å. 1956. On spermiogenesis, morphology of the spermatozoon, and biology of fertilization among invertebrates. Zoologiska Bidrag från Uppsala 31: 355– 480, pls. 1–6. Franzen, Å. 1974. Sperm ultrastructure in some Polychaeta. Pp. 267–278. In Afzelius, B. A. (ed.), The Functional Anatomy of the Spermatozoan. Pergamon Press, Oxford and New York.

Spionida

$!#

George, J. D. 1966. Reproduction and early development of the spionid polychaete Scolecolepides viridis (Verrill). Biological Bulletin 130: 76–93. Gibson, G. 1997. Variable development in the spionid Boccardia proboscidea (Polychaeta) is linked to nurse egg production and larval trophic mode. Invertebrate Biology: 116: 213–226. Gibson, G. D. and A. J. F. Gibson. 2004. Heterochrony and the evolution of poecilogony: generating larval diversity. Evolution 58: 2704–2717. Gibson, G. D. and Harvey, J. M. L. 2000. Morphogenesis during asexual reproduction in Pygospio elegans Claparède (Annelida, Polychaeta). Biological Bulletin 199: 41–49. Gibson, G. D. and I. G. Paterson. 2003. Morphogenesis during sexual and asexual reproduction in Amphipolydora vestalis (Polychaeta: Spionidae). New Zealand Journal of Marine and Freshwater Research 37: 741-752. Gibson, G. D. and Smith, H. L. 2004. From embryos to juveniles: morphogenesis in the spionid Boccardia proboscidea (Polychaeta). Invertebrate Biology 123: 136–145. Green, K. D. 1982. Uncispionidae, a new polychaete family (Annelida). Proceedings of the Biological Society of Washington 95: 530–536. Greve, W. 1974. Planktonic spermatophores found in a culture device with spionid polychaetes. Helgoländer Meeresuntersuchungen 26: 370–374. Gudmundsson, H. 1985. Life history patterns of polychaete species of the family Spionidae. Journal of the Marine Biological Association of the United Kingdom 65: 93–111. Guérin, J. -P. and P. Kerambrun. 1984. Role of reproductive characters in the taxonomy of spionids and elements of speciation in the ‘Malacoceros fuliginosus complex.’ In Fischer, A. and Pfannenstiel, H. -D. (eds), Polychaete Reproduction: Progress in Comparative Reproductive Biology. Fortschritte der Zoologie 29: 317–333. Hannerz, L. 1956. Larval development of the polychaete families Spionidae Sars, Disomidae Mesnil and Poecilochaetidae n. fam. in the Gullmar Fjord (Sweden). Zoologiska Bidrag från Uppsala 31: 1–204. Hartman, O. 1944. Polychaetous annelids Part VI. Paraonidae, Magelonidae, Longosomidae, Ctenodrilidae, and Sabellariidae. Allan Hancock Pacific Expeditions 10(3): 311–389, pls. 27–42. Hartman, O. 1947. Disoma franciscanum, a new marine annelid from California. Journal of the Washington Academy of Science 37: 160–169. Hartman, O. 1965. Deep-water benthic Polychaetous annelids off New England to Bermuda and other North Atlantic areas. Allan Hancock Foundation, Occasional Papers No. 38: 1–378. Hobson, K. D. and R. H. Green. 1968. Asexual and sexual reproduction of Pygospio elegans (Polychaeta) in Barnstable Harbor, Massachusetts. Biological Bulletin 135: 410. Imajima, M. 1959. A description of a new species of the Spionidae (Polychaeta), Nerinides yamaguchii n. sp., with notes on its development. Journal Hokkaido Gakugei University 10: 155–159. Irvine, S. Q., Chaga, O. and Martindale, M. 1999. Larval ontogenetic stages of Chaetopterus: Developmental heterochrony in the evolution of chaetopterid polychaetes. Biological Bulletin 197: 319–331. Irvine, S. Q. and Martindale, M. Q. Nov. 1, 1999. Laboratory culture of the larvae of spionidan polychaetes Marine Models Electronic Record. [serial online]. Available from http://www.mbl.edu/BiologicalBulletin/MMER/ECK/IrvTit.html. Jamieson, B. G. M. 1986a. Onychophoran–euclitellate relationships: evidence from spermatozoal ultrastructure. Zoologica Scripta 15: 141–155.

$!$ Reproductive Biology and Phylogeny of Annelida Jamieson, B. G. M. 1986b. Yasuzumi Memorial Lecture. Some recent studies on the ultrastructure and phylogeny of annelid and uniramian spermatozoa. Development, Growth and Differentiation 28: 25–26. Jamieson, B. G. M and Rouse, G. W. 1989. The spermatozoa of the Polychaeta (Annelida): an ultrastructural review. Biological Reviews 64: 93–157. Levin, L. A. 1984a. Life history and dispersal patterns in a dense infaunal polychaete assemblage: community structure and response to disturbance. Ecology 65: 1185– 1200. Levin, L. A. 1984b. Multiple patterns of development in Streblospio benedicti (Webster) from three coasts of North America. Biological Bulletin 166: 494–508. Levin. L. A. and E. L. Creed. 1986. Effect of temperature and food availability on reproductive responses of Streblospio benedicti (Polychaeta: Spionidae) with planktotrophic or lecithotrophic larvae. Marine Biology 92: 103–113. Maciolek, N. J. 1987. New species and records of Scolelepis (Polychaeta: Spionidae) from the east coast of North America, with a review of the subgenera. Bulletin of the Biological Society of Washington No. 7: 16–40. McEuen, F. S. 1979. Observations on the Reproductive Morphology of some California Spionid Polychaetes. M. S. Thesis, University of the Pacific, Stockton, California. 42 pp. Mesnil, F. 1896. Études de morphologie externe chez les annélides. I. Les spionidens des côtes de la marche. Bulletin Scientifique de la France et de la Belique 29: 110– 287, pls. 7–15. Mesnil, F. 1897. Études de morphologie externe chez les annélides. II. Remarques complémentaires sur les Spionidiens. La familie nouvelle des Disomidiens. La place des Aonides (senus Tauber, Levinsen). Bulletin Scientifique de la France et de la Belique 30: 83–100. Mesnil, F. 1925. Classification, affinities et systématique des Spionidiens. Bulletin de la Société Zoologique de France 49: 672–680. Mileikovsky, S. A. 1967. Larval development of Spiochaetopterus typicus M. Sars (Polychaeta, Chaetopteridae) from the Barents Sea and taxonomy of the Family Chaetopteridae and the Order Spiomorpha. Doklady Akademiya Nauk SSSR 174:733–736. [English translation of Vol. 174, with text published on pp. 403–405] Morgan, T. S., Wilson, A. J. Rogers, A. and Paterson, G. J. L. 1997. Variable life history in the polychaete worm, Pygospio elegans. Marine Biological Association of the United Kingdom, Annual Report for 1997, pp. 37–38. Muus, B. J. 1967. The fauna of Danish estuaries and lagoons: distribution and ecology of domination species in the shallow reaches of the mesohaline zone. Meddelelser fra Kommissionen for Danmarks Fiskeri- og Havunderersøgelser, new series 5(1): 1–316. Orrhage, L. 1964. Anatomische und morphologische Studien über die Polychätenfamilien Spionidae, Disomidae, und Poecilochaetidae. Zoologiska Bidrag från Uppsala 36: 335–405. Petersen, M. E. 1984a. Chaetopterus variopedatus (Annelida: Polychaeta: Chaetopteridae): another victim of the “characteristic species” disease. American Zoologist 24: 64A. Petersen, M. E. 1984b. Chaetopterus variopedatus (Renier) (Annelida: Polychaeta: Chaetopteridae): a species complex. What species are being used at MBL? Biological Bulletin 167: 513. Pettibone, M. H. 1963a. Revision of some genera of polychaete worms of the family Spionidae, including the description of a new species of Scolelepis. Proceedings of the Biological Society of Washington 76: 89–104.

Spionida

$!%

Pettibone, M. H. 1963b. Marine polychaete worms of the New England Region. I. Families Aphroditidae through Trochochaetidae. Bulletin of the United States National Museum 227, Part 1: 1-356. Potts, F. A. 1914. Polychaeta from the N. E. Pacific: The Chaetopteridae. With an account of the phenomenon of asexual reproduction in Phyllochaetopterus and the description of two new species of Chaetopteridae from the Atlantic. Proceedings of the Zoological Society of London 1914: 955–914, pls. 1–6. Radashevsky, V. I. 1994. Life history of a new Polydora species from the Kurile Islands and evolution of lecithotrophy in polydorid genera (Polychaeta: Spionidae). Ophelia 39: 121–136. Radashevsky,V. I. 1996. Morphology, ecology and asexual reproduction of a new Polydorella species (Polychaeta: Spionidae) from the South China Sea. Bulletin of Marine Science 58: 684–693. Radashevsky, V. I. and Cárdenas, C. A . 2004. Morphology and biology of Polydora rickettsi (Polychaeta: Spionidae) from Chile. New Zealand Journal of Marine and Freshwater Research 38: 243–254. Radashevsky, V. I. and Nogueira, J. M. de M. 2003. Life history, morphology and distribution of Dipolydora armata (Polychaeta: Spionidae). Journal of the Marine Biological Association of the United Kingdom 83: 375–384. Rasmussen, E. 1953: Asexual reproduction in Pygospio elegans Claparède (Polychaeta Sedentaria). Nature 171: 1161. Rasmussen, E. 1973. Systematics and ecology of the Isefjord marine fauna (Denmark). Ophelia 11: 1–495. Retzius, G. 1904. Zur Kenntnis der Spermien der Evertebraten. I. Biologische Untersuchungen von Gustaf Retzius. Neue Folge 11: 79–102. Rice, S. A. 1978. Spermatophores and sperm transfer in spionid polychaetes. Transactions of the American Microscopical Society 97: 160–170. Rice, S. A. 1980. Ultrastructure of the male nephridium and its role in spermatophore formation in spionid polychaetes. Zoomorphology 95: 181–194. Rice, S. A. 1981. Spermatogenesis and sperm ultrastructure in three species of Polydora and in Streblospio benedicti (Polychaeta: Spionidae). Zoomorphology 97:1– 16. Rice, S. A. 1992. Polychaeta: Spermatogenesis and spermiogenesis. Pp. 129–151. In Harrison, F. W. and Gardiner, S. L. (eds), Microscopic Anatomy of Invertebrates, Vol. 7: Annelida. Wiley-Liss, Inc. Rice, S. A. and D. J. Reish. 1976. Egg capsule formation in the polychaete Polydora ligni: confirmation of a hypothesis. Bulletin of the Southern California Academy of Sciences 75: 285–286. Richards, S. L. 1970. Spawning and reproductive morphology of Scolelepis squamata (Spionidae: Polychaeta). Canadian Journal of Zoology 48: 1369–1379. Rouse, G. W. 1988. An ultrastructural study of the spermatozoa from Prionospio cf. queenslandica and Tripolydora sp.: two spionid polychaetes with different reproductive methods. Acta Zoologica 69: 205–216. Rouse, G. W. 1999. Polychaeta, Including Pogonophora and Myzostomida. Pp. 81– 124. In Jamieson, B.G.M. (ed.), Reproductive Biology of the Invertebrates. Volume 9, Part B, Progress in Male Gamete Ultrastructure and Phylogeny. Wiley-Liss, Inc. New York. Rouse, G. W. and K. Fauchald. 1997. Cladistics and polychaetes. Zoologica Scripta 26: 139–204. Rouse, G. W. and B. G. M. Jamieson. 1987. An ultrastructural study of the spermatozoa of the polychaetes Eurythoe complanata (Amphinomidae), Clymenella sp. and

$!& Reproductive Biology and Phylogeny of Annelida Micromaldane sp. (Maldanidae), with definition of sperm types in relation to reproductive biology. Journal of Submicroscopical Cytology 19: 573–584. Sato-Okoshi, W., Sugawara, Y. and T. Nomura. 1990. Reproduction of the boring polychaete Polydora variegata inhabiting scallops in Abashiri Bay, North Japan. Marine Biology 104:61–66. Scheltema, R. S. 1974. Relationship of dispersal to geographical distribution and morphological variation in the polychaete family Chaetopteridae. Thalassia Jugoslavica 10: 297–312. Scheltema, R. S., Blake, J. A., and Williams, I. P. 1997. Planktonic larvae of spionid and chaetopterid polychaetes from off the west coast of the Antarctic Peninsula. Bulletin of Marine Science 60: 396–404. Sigvaldadóttir, E., Mackie, A. S. Y., and Pleijel, F. 1997. Generic interrelationships within the Spionidae (Annelida: Polychaeta). Zoological Journal of the Linnaean Society 119: 473–500. Simon, J. L. 1967. Reproduction and larval development of Spio setosa (Spionidae: Polychaeta). Bulletin of Marine Science 17: 398–431. Söderström, A. 1920. Studien über die Polychätenfamilie Spionidae. Inaugural Dissertation, Uppsala, Almquist and Wicksells. 288 pp. Stock, M. W. 1964. Anterior Regeneration in Spionidae. M. S. Thesis, University of Connecticut, Storrs, Conneticut. 91 pp. Thorson, G. 1946. Reproduction and larval development of Danish marine bottom invertebrates, with special reference to the planktonic larvae in the sound (Øresund). Meddelelser fra Kommissionen for Danmarks Fiskeri- og Havunderersøgelser, Series Plankton 4: 1–523. Tzetlin, A., Britayev, B. and Temir, A. 1985. A new species of the Spionidae (Polychaeta) with asexual reproduction associated with sponges. Zoologica Scripta 14: 177–181. Werner, B. 1953. Beobachtungen über den Nahrungserwerb und die Metamorphose der Metatrochophora von Chaetopterus variopedatus Renier u. Claparède (Polychaeta sedentaria). Helgoländer Wissenschaftliche Meeresuntersuchungen 4: 225–238. Williams, J. D. 2004. Reproduction and morphology of Polydorella (Polychaeta: Spionidae), including the description of a new species from the Philippines. Journal of Natural History 38: 1339–1358. Wilson, D. P. 1928. The larvae of Polydora ciliata Johnston and Polydora hoplura Claparède. Journal of the Marine Biological Association of the United Kingdom 15: 567–589. Wilson, D. P. 1982. The larval development of three species of Magelona (Polychaeta) from localities near Plymouth. Journal of the Marine Biological Association of the United Kingdom 62: 385–401. Woodwick, K. H. 1960. Early larval development of Polydora nuchalis Woodwick, a spionid polychaete. Pacific Science 14: 122–128. Woodwick, K. H. 1963. Comparison of Boccardia columbiana Berkeley and Boccardia proboscidea Hartman (Annelida, Polychaeta). Bulletin of the Southern California Academy of Sciences 62: 132-139. Woodwick, K. H. 1977. Lecithotrophic larval development in Boccardia proboscidea Hartman. Pp. 347–371. In Reish, D. J. and Fauchald, K., (eds) Essays on Polychaetous Annelids in Memory of Dr. Olga Hartman. Allan Hancock Foundation, University of Southern California, Los Angeles.

14

CHAPTER

Problematic Annelid Groups Günter Purschke

14.1 PHYLOGENY AND SYSTEMATICS Introduction. The title of this chapter refers to the fact that the groups considered below are of still unknown or uncertain phylogenetic position, and most of them have been placed incertae sedis at various positions in recent phylogenetic analyses (Rouse and Fauchald 1997; Glasby et al. 2000; Rouse and Pleijel 2001). These groups, namely Polygordiidae, Protodrilida, Nerillidae, Dinophilidae, Diurodrilidae, Aeolosomatidae, Potamodrilidae, Parergodrilidae and Hrabeiella periglandulata, mainly comprise meiofaunal or interstitial species. They are characterized by small body dimensions and a seemingly simple organization. Most of the species inhabit marine sediments in intertidal and subtidal regions, but limnic, terrestrial, as well as continental ground water species are also included in this assemblage of polychaetes. Formerly some of these taxa were thought to belong to a single systematic group of more or less primitive annelids, Archiannelida (e.g., Westheide 1990). Because of their simple organization they were considered to represent a primitive state in Annelida, close to the annelid stem species. However, modern investigations have revealed that they neither form a monophyletic group nor retain many plesiomorphies (Hermans 1969; Jouin 1971; Westheide 1985, 1990). Instead these groups have proved to be highly derived taxa, most likely secondarily simplified in certain characters, but highly specialized in others and miniaturized in the course of invading meiofaunal habitats (Jouin 1967, 1968, 1978-79; Westheide and Riser 1983; Westheide 1985; Bunke 1986; Purschke and Jouin 1988; Nordheim 1989b, 1991a, 1991b; Purschke and Jouin-Toulmond 1993; Kristensen and EibyeJacobsen 1995; Purschke and Müller 1996; Rota 1998; Hessling and Purschke 2000; Müller and Westheide 2002; Purschke 1985a, b, 1990a, b, 1992, 1993, 1999, 2002, 2003). Moreover, most of them are obviously not closely related to one another. As a consequence, the concept Archiannelida has been eliminated from modern zoological systems. However, the phylogenetic relationships of most groups remained obscure and unresolved (see e.g., Zoologie, Fachbereich Biologie/Chemie, University of Osnabrueck, D-49069 Osnabrueck, Germany

$" Reproductive Biology and Phylogeny of Annelida Glasby et al. 2001; Rouse and Fauchald 1997), and it is conceivable that some groups may fall into well-known taxa comprising “typical” polychaetes. Although for some of these families no autapomorphies have yet been found (see Fauchald and Rouse 1997), they are usually regarded as being monophyletic. Dinophilidae. Probably the most famous example is Dinophilidae, which today usually are regarded as representative of progenetically evolved Dorvilleidae (Eunicida) (Åkesson 1977; Eibye-Jacobsen and Kristensen 1994; Westheide 1982, 1985, 1990; Westheide and Riser 1983). This proposed relationship is based on their general resemblance to juvenile stages of larger species of Dorvilleidae and Eunicida. Since such larval characters mostly consist of absences or reductions two explanations are conceivable: (1) The group formed by uniting such dorvilleid taxa exhibit an increasing progenetic organization, with Dinophilidae as the most derived clade, is in fact monophyletic (see Eibye-Jacobsen and Kristensen 1994) or (2) the group simply represents a morphological grade that is not monophyletic (Struck et al. 2002a, 2005). It cannot be excluded that there are more than one evolutionary pathway leading to progenetic taxa within Dorvilleidae and Eunicida. Moreover, since there are no specific larval synapomorphies for Dinophilidae and Dorvilleidae, a completely different origin of Dinophilidae is also possible. Analyses using molecular data produced no evidence of this proposed relationship (Struck et al. 2002a). Unfortunately, this investigation does not allow us to give an alternative classification of Dinophilidae. After critical evaluation of all studies, given the obvious lack of clear morphological synapomorphies, the phylogenetic relationship of Dinophilidae should still be regarded as an open question (Struck et al. 2002a), hence their inclusion in this chapter. Diurodrilus. Formerly Diurodrilus was also placed within Dinophilidae, but affinities remain uncertain (Kristensen and Niilonen 1982; Westheide 1990). The placement of Diurodrilus as another progenetic taxon of Dorvilleidae (Rouse and Pleijel 2001), however, needs to be confirmed by clear apomorphic characters which have still not been found (Kristensen and Eibye-Jacobsen 1995; Kristensen and Niilonen 1982; Westheide 1990). Molecular data are lacking. Nerillidae. It has been suggested that Nerillidae is part of Aciculata (Rouse and Fauchald 1997; Westheide 1990; Worsaae and Kristensen 2003). As is the case for Dinophilidae, a progenetic origin can also be inferred for Nerillidae. However, they do not resemble any juvenile stage of extant polychaetes in any obvious respect (Westheide 1990; Westheide and Purschke 1996). Repeatedly, a resemblance to juvenile Onuphidae has been brought into discussion (Westheide 1990) and this warrants further investigation, as does some resemblance with Aberranta (Rouse and Pleijel 2001; Worsaae et al. 2005). The phylogeny within the taxon, consisting of approximately 50 species assigned to 18 genera, is still unresolved. A phylogenetic analysis based on morphological and molecular data has been published recently (Worsaae 2005).

Problematic Annelid Groups

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Protodrilida. Another group that has been considered well established is Protodrilida, comprising Protodriloides, Protodrilus, Parenterodrilus and Saccocirrus (Purschke and Jouin 1988). A close relationship between Saccocirrus and Protodrilus has been accepted since Goodrich (1901).The suggested close relationship of Saccocirrus to Spionida (Hatschek 1893) received support from anatomical and ultrastructural investigations on the nervous system, sense organs and prostomial appendages in the two groups (Orrhage 1974; Purschke 1990a, 1993; Purschke and Jouin-Toulmond 1993, 1994). A close relationship between Saccocirrus and Polygordius, considered to be valid since Marion and Bobretzky (1875), was not supported. Currently Polygordius is regarded as not allied with Protodrilida but instead is of uncertain position (Westheide 1990). A close relationship of Polygordius to Opheliidae has been discussed repeatedly (see Hermans 1969; Rouse and Pleijel 2001; Westheide 1990). In recent analyses using 18S rDNA sequences the only taxa falling into one cluster are Saccocirrus and Polygordius, whereas neither a close relationship of any Protodrilida, either to each other, nor to any of these to Spionida, was found (Bleidorn et al. 2003; Rota et al. 2001; Struck et al. 2002a, b; Struck 2003). In an extended analysis, using sequences of more than 150 polychaetes, these taxa cluster far apart from each other in various clades but without significant support (Struck 2003, unpublished). Thus, their relationships need to be clarified by further molecular and morphological studies. Aelosomatidae, Potamodrilidae, Parergodrilidae. The remaining taxa, Aelosomatidae and Potamodrilidae (commonly united as Aphanoneura), Parergodrilidae and Hrabeiella periglandulata, have been discussed as either belonging to Clitellata, or at least being related to them (see e.g., Purschke et al. 2000). These hypotheses were based on presence or absence of different clitellate characters (Struck et al. 2002b). However, absence of a true clitellum, different structure of the spermatozoa and genital organs, as well as occurrence of modified and more or less internal nuchal organs precluded inclusion within Clitellata (Hessling and Purschke 2000; Purschke 1986, 1999, 2000; Purschke et al. 2000; Rota and Lupetti 1997; Rota 1998). Therefore, similarities between these taxa and Clitellata were regarded as convergently evolved, most likely as structural adaptations to similar environments. It should be noted that structural resemblance is greatest with Hrabeiella. Since these correspondences do not only represent absences or possible losses but rather specific structures as well, the possibility of a sister-group relationship of Hrabeiella to Clitellata has been brought into discussion again (Purschke 2003). Such a relationship is neither rejected nor significantly supported in phylogenetic analyses using various molecular markers (Jördens et al. 2004; Struck 2003). On the other hand, Parergodrilidae obviously is not to be related to any of the taxa mentioned above. Its monophyly is supported by molecular and morphological data (Purschke 1999; Struck et al. 2002b). According to recent molecular studies they always fall into a clade comprising Orbiniidae and Questidae (Bleidorn et al. 2003; Purschke et al. 2004; Struck 2003; Struck et

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Reproductive Biology and Phylogeny of Annelida

al. 2002b). This position appears to be stable, it is supported by high bootstrap values, and cannot be rejected on the basis of morphological evidence. The validity of Aphanoneura, consisting of Aeolosoma, Rheomorpha, Nectohelmis, Hystricosoma and Potamodrilus, was questioned by Bunke (1967, 1985, 1986), who excluded Potamodrilus from this group without indicating a probable sister group. Following an investigation of the central nervous system and sense organs this view was not supported (Hessling and Purschke 2000; Purschke and Hessling 2002). The same conclusion was reached by Marotta et al. (2003) following an investigation of the spermatozoa in Aeolosoma singulare. Aphanoneura is also validated by molecular studies using 18S rDNA and cytochrome Oxidase I in which Aeolosoma, Rheomorpha and Potamodrilus were always found to be a monophyletic group (Struck 2003; Struck and Purschke 2005). A relationship to Clitellata is not supported in any recent molecular analysis (Rota et al. 2001; Struck et al. 2002b; Struck 2003). Moreover, such a clitellate relationship as suggested by Timm (1981) and Brinkhurst and Nemec (1987) is also not supported by the structure of the nervous system, which exhibits several specific (apomorphic) clitellate features (Hessling and Purschke 2000; Purschke and Hessling 2002). These findings clearly support the rejection of a close relationship of Aphanoneura and Clitellata, and similarities have to be regarded as convergences.

14.2 ANATOMY WITH REFERENCE TO THE REPRODUCTIVE SYSTEM Dinophilidae. Dinophilidae comprising Dinophilus, Trilobodrilus and, most likely, Apharyngtus, are small animals with a trunk consisting of only a small and constant number of achaetigerous segments. Various numbers of segments have been attributed to dinophilid species because of their indistinct outer metameric annulation and the lack of chaetae. However, recent investigations on the central nervous system showed that the number of segments has been overestimated; only six pairs of ganglionic concentrations were found in Trilobodrilus and Dinophilus (Müller and Westheide 2002). Dinophilids are gonochoristic, with the exception of Trilobodrilus hermaphroditus, which is the only hermaphroditic species known in the taxon (Riser 1999; Westheide 1990). Dinophilus gyrociliatus is a highly dimorphic species with dwarf males lacking a gut system and measuring only 50 mm in length. The reproductive system appears to be rather similar in the group (Jägersten 1943; Westheide 1971, 1990; Westheide and Schmidt 1974). The male organs consist of an unpaired testis, paired seminal vesicles with paired spermioducts, and an unpaired copulatory organ (Jägersten 1943, 1944; Westheide 1971, 1990; Westheide and Schmidt 1974). The testis extends anteriorly as paired lobes from an unpaired ventral part situated below the gut; likely it represents a coelomic cavity in which spermiogenesis takes place. The vesicles, one pair in Dinophilus and two

Problematic Annelid Groups

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pairs in Trilobodrilus, are anterior dilatations of the ciliated gonoducts. In the vesicles mature spermatozoa accumulate. The male ducts merge in the unpaired copulatory organ, which mainly is a multicellular gland made up of two to four different types of gland cells (Westheide 1988). Central projections of epithelial and gland cells form the penis proper, which bears an unpaired opening. According to Westheide (1990) the copulatory organ develops out of an invagination of the epidermis and underlying body musculature. The copulatory organ is innervated from two penis ganglia which form a fiber mass around the organ in D. gyrociliatus and T. hermaphroditus (see Müller 1999; Müller and Westheide 2002; Windoffer and Westheide 1988). The females possess one, two or four ovaries that are located in a similar position in the body. In T. hermaphroditus the female organs are situated in front of the male organs (Riser 1999). The ovaries are made up of coelomic cavities (Westheide 1990). There are only a few developing oocytes visible at the same time; there may be about 10 mature oocytes in T. axi, but only two in A. punicus and one in T. hermaphroditus (Riser 1999; Westheide 1971; Westheide and Schmidt 1974). The existence of female ducts has been reported only for some species (Westheide 1990); confirmations by ultrastructural investigations are desirable. Diurodrilidae. Knowledge about the reproductive organs of Diurodrilus is scanty. Sexes are separate. Females carry one or two mature oocytes, and paired rows of vitellogenic oocytes have been observed in D. subterraneus and D. westheidei (Kristensen and Niilonen 1982; Mock 1981). Female gonoducts were stated to be absent by Mock (1981). In the males spermatozoa develop in the coelom and are discharged via two seminal vesicles followed by short ciliated funnels opening into a cloaca (Kristensen and Eibye-Jacobsen 1995). Nerillidae. In Nerillidae individuals are minute; the group includes the smallest metazoans with a complete set of internal organs. Many taxa do not exceed 0.5 mm and adult Nerillidium gracile may be only 0.3 mm long. Species have 7–9 chaetigerous segments, which may or may not have cirri (Westheide 1990; Westheide and Purschke 1996; Müller 2002). Most species are gonochoristic, but there are a number of hermaphroditic species as well, sometimes within the same genus. Gonoducts are simple and consist of paired ciliated oviducts and spermioducts (Fig. 14.1A–D; Jouin 1968; Westheide 1990). Oocytes and spermatocytes develop freely in the body cavity. In hermaphroditic species they occur together and usually the female ducts are situated posteriorly, opening in chaetiger 8 in most cases. There are two to three pairs of spermioducts, which may open separately or in a common pore; their openings are found between segments 5 and 7 and may be surrounded by additional gland cells (Fig. 14.1A). The arrangement of ducts is highly diverse and taxon specific (e.g. Jouin 1967, 1968). In Meganerilla clavata a pair of pits has been described close to the male openings, presumably for the storage of sperm (Jouin 1968). Generally the ducts possess a more or less distinct funnel (Fig. 14.1C) and open after a short distance in the following segment. As a rule the ducts are formed

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Fig. 14.1. Genital organs. A, B. Nerilla antennata. Anti-acetylated-α-tubulin staining; confocal laser scanning microscope micrographs. A. Male with three pairs of spermioducts (sd) opening in common genital pore (arrow). B. Female with one pair of separate oviducts; ne nephridium, sn segmental nerve, vcb ventral ciliary band, vnc ventral nerve cord. C–D. Troglochaetus beranecki. C. Densely ciliated funnel of oviduct, arrows point to junctions between funnel cells; n nucleus, r ciliary rootlet. D. Oviduct formed by single cell; note thin epidermal cell. E. Saccocirrus sp. Cross section of penis in penial sheath. Abbreviations: ci, cilia; coe, coelomic cavity; cu, cuticle; ep, epidermal cell; gl, gland cell; iep, inner epithelium; ne, nephridium; oep, outer epithelium; ov, oviducts; ps, penial sheath; sf, seminal funnel; sn, segmental nerve; sp, spermatozoa; vcb, ventral ciliary band; vnc, ventral nerve cord. Figs A, B, Courtesy of M.C.M. Müller and K. Worsaae, Figs C–E, original.

Problematic Annelid Groups

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by one cell, as seen in cross section (Fig. 14.1D; see Westheide 1988 for diversity of gonoducts). Nephridia are lacking in segments with gonoducts, indicating that these organs are homologous (Worsaae and Müller 2004). Polygordiidae. Polygordiids are fairly large compared with the other taxa discussed in this chapter and may be up to 10 cm long, with more than 200 segments (Rouse and Pleijel 2001; Westheide 1990). They are threadlike, roundish, with almost no external signs of segmentation and thus resemble large nematodes. This resemblance also applies to their locomotion, as a result of the absence of circular muscle fibers. Usually species are gonochoristic; reports of a hermaphroditic species have been called into question (Westheide 1990). Gonads develop in a large number of segments and may start between segment 20 and 70 and extend to the end of the body. Male gonads usually begin in different segments than the female organs. Since gonoducts have not been observed, it is suggested that gametes are released by rupture of the body wall. Life cycle data indicate that at least some species (e.g. Polygordius lacteus) reproduce more than once in their lifetime (Nordheim 1984). Protodrilida. Species of this taxon are thread-like and possess multisegmented trunks from about 20 segments and 2 mm in length (Protodrilus minutus) to more than 200 segments and up to 8 cm in length in Saccocirrus major (see Jouin and Rao 1987; Nordheim 1989a; Pierantoni 1908; Westheide 1990). A pair of palps is characteristic for all members of the group (Fig. 14.2A). The palps are highly mobile in Saccocirrus, Protodrilus and Parenterodrilus, supplied with internal coelomic cavities and originate ventrolaterally, whereas they are less mobile in Protodriloides, without coelomic cavities and arise anteriorly from the prostomium (Jouin 1966; Purschke and Jouin 1988; Purschke 1993). Two pygidial adhesive lobes are characteristic and additional segmentally arranged adhesive glands are present in some species such as Protodrilus adhaerens and Protodriloides symbioticus. Sexes are separate and usually the number of fertile segments is large. The simplest genital organs are found in the males of Protodriloides, where each fertile segment is supplied with a pair of spermioducts (Jouin 1966). In a large female of Protodriloides chaetifer consisting of a total of 46 segments fertile segments extend from chaetiger 20 to 44 with a total number of 17 large oocytes. In a female of Protodriloides symbioticus comprising 15 segments fertile segments were found from segment 7 backwards, with a total number of 4–10 mature oocytes. Epidermal glands are present in the fertile regions of both species; those of the females produce a cocoon surrounding the eggs at spawning (Jouin 1966; Swedmark 1954). In Protodrilus, structure and arrangement of genital organs are speciesspecific in both sexes, but is not sufficiently known for every species (Jägersten 1952; Jouin 1970; Nordheim 1989a; Pierantoni 1908). In females the fertile region may extend from segment 10 and backwards. The number of oocytes per segment usually varies from 2 to 40–60, depending on the species (Nordheim 1989a). In P. haurakiensis up to 160 eggs per segment

$"$ Reproductive Biology and Phylogeny of Annelida

Fig. 14.2. Genital organs in Protodrilidae. A. Protodrilus oculifer. Mature male; arrow points to beginning of fertile region, slightly squeezed. B–C. Schematic representation of lateral organs, sperm funnels, genital openings and fertile segments. B. Protodrilus hatscheki. C. Parenterodrilus taenioides. D. Protodrilus helgolandicus Spermatophore attached to female, oc oocyte. E–G. Lateral organs. E. Protodrilus ciliatus. Interference contrast micograph showing one lateral organ with numerous gland cell necks (arrowheads). F. Protodrilus purpureus. First (arrowhead) and second (arrow) lateral organ, SEM micrograph. Inset: Enlargement of densely ciliated organ in Protodrilus ciliatus. G. Protodrilus helgolandicus. Cross section through lateral organ. Abbreviations: cc, ciliated cell; cu, cuticle; ecm, extracellular matrix; ep, epidermis; gl, gland cell; n, nerve; p, palp; sp. spermatozoa; spd, sperm duct. A original, B courtesy of C. Jouin-Toulmond, modified from Jouin, C. 1970. Cahiers de Biologie Marine 11: 367-434, Fig. 6, C modified from Jouin-Toulmond, C. and Purschke, G. 2004. Zoomorphology 123: 139-146, Fig. 3A. D–G courtesy of H. v. Nordheim.

Problematic Annelid Groups

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were reported (Nordheim 1989a). Some species possess a number of short ciliated oviducts. Eggs are deposited either via these oviducts, via the nephridia, by shedding of posterior segments, or by rupture of the body wall (Nordheim 1983, 1991a). The male organs consist of testes, short ciliated spermioducts and so-called lateral organs (Jägersten 1952; Jouin 1970; Nordheim 1989a, 1991b). The latter, present in some anterior segments, are elongated grooves into which a large number of gland cells open (Fig. 14.2B). Usually the first lateral organs occur in front of the fertile segments that normally are found from segments 9–13 and backwards (Fig. 14.2A, B). These lateral organs may be discontinuous or continuous and three to five pairs of male ducts open in the posterior lateral organs (Fig. 14.2B). These organs serve in spermatophore formation. The structure of the male organs is basically similar in Parenterodrilus taenioides but the females are still unknown in this species (Fig. 14.2C; Jouin-Toulmond and Purschke 2004). Species of Saccocirrus have either bilateral or paired unilateral unpaired genital organs (Brown 1981). In the latter group gonads either are on the left side or male and female gonads may be on different sides of the body (female organs left, male organs right). The fertile region is restricted to the middle of the trunk and there may be more than 100 fertile segments (Brown 1981; Jouin and Rao 1987; Westheide 1990). Each fertile segment is supplied with a complete set of genital organs. In females they consist of an oviduct which joins a seminal receptacle, in males there is a spermioduct, seminal vesicle and a protrusible penis, which lies in an epidermal pouch just behind the parapodia (e.g., Brown 1981). In the penis the duct is ciliated and some mature spermatozoa are found in this duct and the seminal vesicle. The genital opening is situated subdistally (Purschke unpublished). The organ consists of epithelial, sensory and glandular cells (Fig. 14.1E). Stylet-like structures of probable cuticular origin have been described for some species at the light microscopic level (e.g. S. minor, S. heterochaetus, S. krusadensis, see Aiyar and Alikunhi 1944; Brown 1981; Jouin 1975), while in others such structures are absent (e.g. S. tridentiger, see Brown 1981). The cuticular nature of these rods has not been confirmed by ultrastructural investigations; preliminary observations in a Saccocirrus sp. with unilateral gonads indicate the presence of small intracellular rootlet-like structures, somewhat resembling skeletal elements found in the copulatory organ of Microphthalmus cf. similis (see Westheide 1979). In Saccocirrus heterochaetus genital chaetae are present that differ between males and females (Jouin 1975). Aeolosomatidae and Potamodrilidae. Species of Aphanoneura are small worms which are 0.3–10 mm long and only 60–110 µm across. All species are hermaphroditic and possess only a few fertile segments. Most species of Aeolosomatidae likely reproduce exclusively by paratomy (Bunke 1967). If genital organs are developed, there is one pair of ovaries in a midbody segment, but only one of them produces mature oocytes. Only one mature oocyte has been found at a time. There are no special

$"& Reproductive Biology and Phylogeny of Annelida female ducts, but the ventral epidermis below the mature oocytes becomes glandular and forms a genital pore. Mostly there are three pairs of small seminal receptacles ventrally in the anteriormost chaetigers. Pairs of testes may be found in anterior and posterior segments. Male gonoducts are absent and spermatozoa are released through the nephridia (Bunke 1967). In Potamodrilus only sexual reproduction occurs. The female organs consist of a pair of ovaries situated in the fifth segment, an unpaired ventral pore surrounded by prominent glands, and an unpaired seminal receptacle just in front of the female atrium. Male gametes develop in segments 4 and 5 and are discharged through a pair of highly convoluted spermioducts, which unite and form a single ventral male pore in front of the female opening. Parergodrilidae. Both species are comparatively stout and possess up to ten chaetigers. Sexes are separate. Female organs are only known from histological investigations (Karling 1958; Reisinger 1925; Rota 1998). In Stygocapitella subterranea they consist of an unpaired ovisac of coelomic origin containing a pair of ovaries. A pair of female ducts leads to the female opening in the furrow between chaetigers 9 and 10. Part of the latter is differentiated into a seminal receptacle (Karling 1958). In the following segments ventral cocoon-forming glands are found. In Parergodrilus heideri the ovisacs are paired as well, but the female openings are situated at the posterior end and are directly associated with prominent cocoon-forming glands. Each oviduct forms a seminal receptacle close to the ovisac, containing only a limited number of sperm in each individual (Reisinger 1925, 1960; Rota 1997, 1998; Purschke 1999, 2002). Usually there is only one, comparatively large, mature oocyte at a time, but vitellogenic oocytes are always present in each ovary. The male organs of S. subterranea are made up of a huge seminal vesicle containing the paired testes and numerous developing stages of male gametes. A pair of long convoluted spermioducts supplied with prominent funnels extend from the vesicle and open ventrally, close to the chaetae of chaetiger 9. The sperm ducts are associated with a pair of prominent multicellular prostate glands. The different gland cells open either close to the male pore or into the posterior portion of the male duct (Karling 1958). True copulatory organs are absent. The main differences between the two species are: a paired seminal vesicle, two pairs of spermioducts which open into a common genital atrium together with prostate gland cells, and a pair of large copulatory chaetae in P. heideri, and an unpaired vesicle with only one pair of sperm ducts and absence of copulatory chaetae in S. subterranea (Reisinger 1960; Purschke 2002). Males of P. heideri have two segments more than females but the same number of chaetigers as in S. subterranea. Hrabeiella. Individuals of Hrabeiella periglandulata are about 2 mm long and possess 15 chaetigers. The species is a simultaneous hermaphrodite. The male organs are found in chaetiger 5. The paired organs consist of testes, unequally developed sperm sacs (seminal vesicles), and ciliated spermioducts. The ducts extend ventrally and merge into a midventral

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penial bulb associated with a large prostate gland (Fig. 14.3; Purschke unpublished; Rota and Lupetti 1997; Rota 1998). The testes are situated laterally and stages of spermatogenesis are found in the vesicle. The larger vesicle extends dorsally and may lie above the gut (Fig. 14.3). The unpaired male copulatory organ, about 100 µm in length and 30 µm in diameter, is shifted to one side of the body. The female organs are unpaired and comprise an ovary in chaetigers 12 and 13 as well as an ovisac which may extend into chaetiger 15 (Rota and Lupetti 1997). Oviducts and seminal receptacles have not been observed.

14.3

OOGENESIS

Information about the oogenesis of these polychaetes is scarce and usually limited to light microscopic observations. So generally it cannot be ascertained for every taxon whether oogenesis is extraovarian or intraovarian and which pattern of vitellogenesis occurs (see Eckelbarger

Fig. 14.3. Genital organs. Hrabeiella periglandulata. Cross section through chaetiger 5 showing left seminal vesicle and unpaired copulatory organ, TEM micrograph. Abbreviations: co, copulatory organ; cy, cytophore with spermatids; ep, epidermis; g, gut; te, testis; vnc, ventral nerve cord; vs, seminal vesicle. Micrograph courtesy of K. Rainer.

$# Reproductive Biology and Phylogeny of Annelida 1988). In dinophilids nurse cells have been observed that fuse with developing oocytes (Traut 1969; Riser 1999). In D. gyrociliatus eggs of two sizes are produced: the larger, accounting for about 60–70% of the total number, develop into females, the smaller into dwarf males. Whereas formerly sex determination was considered to be progamous, sex determination is now thought to reside in the male gametes since a univalent sex chromosome was detected. Large eggs are selectively fertilized by female-determining spermatozoa and vice versa (Martin and Traut 1987; Westheide 1990). In Nerillidae oogenesis most likely is extraovarian (Fig. 14.4A, B). Absorption of smaller oocytes by large oocytes has been observed in Mesonerilla biantennata (Jouin 1968). In Polygordius oogenesis very likely is intraovarian and nurse cells have been described by Hempelmann (1906). Mature oocytes accumulate in the coelom after they

Fig. 14.4. Oogenesis. Troglochaetus beranecki. A. Oocyte at prophase of first meiotic division showing synaptonemal complexes (arrows). B. Vitellogenic oocyte with large nucleolus. Abbreviations: n, nucleus; nu, nucleolus; y, yolk granule. A, B original.

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have been released through the peritoneum enclosing the ovary. In Parergodrilidae developing oogonia and vitellogenic oocytes are in close contact with blood spaces. Usually a thin peritoneal lining separates oocytes from the blood. Follicle cells are absent, and oogenesis is clearly intraovarian (Purschke unpublished). In Hrabeiella oocytes are associated with follicle cells inside the ovary (Rainer, unpublished).

14.4 SPERMATOGENESIS AND SPERMATOZOA Introduction. Observations on spermatogenesis are comparatively rare, but the ultrastructure of the spermatozoa is known for a comparatively large number of species (Figs. 14.5, 14.6). Most species have so-called modified spermatozoa (Franzén 1977a), which have also been termed “introsperm” (Jamieson and Rouse 1989). Fine-structural analyses revealed that not only filiform spermatozoa are of this type; some of those resembling ectaquasperm are actually introsperm-like as well, hence the term entaquasperm (Rouse and Jamieson 1987). In general the diversity between the taxa considered here is high. Spermatogenesis generally occurs in the seminal vesicles. In several species large cytophores such as are typical of many annelids are formed (Fig. 14.6L), whereas in others spermatids develop in tetrads. Dinophilidae. Ultrastructure of spermatozoa in Dinophilidae is basically the same in all species investigated: Dinophilus gyrociliatus, Trilobodrilus axi and T. heideri (see Franzén 1977b; Scharnofske 1986). Spermatozoa are filiform and are made up of an elongated head piece, midpiece and tail, which are not sequentially arranged but show a high degree of overlap. The axoneme begins beside the acrosomal vesicle in Trilobodrilus or just below it in Dinophilus, the nucleus forms a long rod along the axoneme, as do the four long mitochondria. A pair of supporting structures below the nucleus and an annulus at the transition between midpiece and tail proper is only described for Trilobodrilus (Scharnofske 1986). Spermatids develop in typical morulae. Diurodrilus. Spermiogenesis in Diurodrilus subterraneus produces an extremely strange spermatozoon, although seemingly of the ect-aqusperm type according to light microscopic observations (Mock 1981; Kristensen and Eibye-Jacobsen 1995). Spermatids develop in tetrads and no cytophores are formed. The mature spermatozoon is characterized by a very large acrosome composed of several compartments that partly envelop the nucleus and the mitochondria. In the nuclear region the plasma membrane carries peculiar mushroom-like bodies. This region is followed by the flagellum, which may be divided into three distinct regions. A typical midpiece is absent or incorporated in the nuclear region. Nerillidae. The highest diversity of sperm structure is found in Nerillidae (Franzén and Sensenbaugh 1984; Purschke and Tzetlin, unpublished). It ranges from typical ect-aquasperm in Trochonerilla mobilis to highly particular introsperm in Nerilla antennata. In other species, such as

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Reproductive Biology and Phylogeny of Annelida

Fig. 14.5. Spermatozoa. Introsperm; schematic drawings after electron microscopic observations, not to same scale. A. Saccocirrus sp. Original. B–C. Protodrilidae, Parenterodrilus taenioides. B. Euspermatozoon; C. Paraspermatozoon. Modified from Jouin-Toulmond, C. and Purschke, G. 2004. Zoomorphology 123: 139-146, Fig. 3B, D. D. Hrabeiella periglandulata. Combined and redrawn from Rota, E. and Lupetti, P. 1997. Tissue and Cell 29: 603-609.Figs. 26, 29, 32, 40, and Nienhüser and Rainer, unpublished E. Parergodrilus heideri. Modified from Purschke, G. 2002. Zoomorphology 121: 125-138, Fig. 6. F. Stygocapitella subterranea. Modified from Purschke, G. and Fursman, M. 2005. Zoomorphology (124: 137-148), Fig. 6. Abbreviations: av, acrosomal vesicle; ax, axoneme; m, mitochondrion; n, nucleus; se, supporting element.

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Nerillidium troglochaetoides and Troglochaetus beranecki, spermatozoa appear to be ect-aquasperm but are highly specific as well (Fig. 14.6A, B; Purschke unpublished). In N. troglochaetoides this mainly refers to the complex acrosome (Fig. 14.6B), whereas in T. beranecki the entire cell is of untypical structure (Fig. 14.6C). In the latter species the spherical anterior part comprises a series of globular vesicles occupying most of the space, an electron-dense nucleus of similar size, and a single minute mitochondrion. Below the nucleus the two centrioles are found, the distal one of which gives rise to an axoneme made up of single microtubules only. This indicates that spermatozoa are most likely immobile. In these two species spermatids develop in tetrads; cytophores are not formed. Spermatids are found all over the body cavity. Spermiogenesis appears to be similar in N. antennata (Franzén and Sensenbaugh 1984). However, the spermatozoa are elongated and thread-like. Head and flagellum are parallel and connected to one another by specific structures, so that the flagellum has the typical form only in its distal region. Polygordiidae. The only taxon with typical ect-aquasperm is Polygordius (see Franzén 1977b): it comprises a small cone-shaped acrosome situated above the spherical nucleus, five spherical mitochondria, and a typical flagellum. Protodrilida. Protodrilida possess spermatozoa which exhibit taxonand species-specific features. In Protodriloides they are aflagellate roundish cells (Jouin 1978–79; Fig. 14.6K). Specific features are an irregularly shaped nucleus, several small mitochondria, a number of electron-dense vesicles originating from the Golgi complex, and several smaller electron-lucent vacuoles. The most characteristic feature of Protodrilus and Parenterodrilus is the co-occurrence of two sperm types, called euspermatozoa and paraspermatozoa (Fig. 14.5B, C; Franzén 1977b; Nordheim 1989b; JouinToulmond and Purschke 2004). Both types are filiform and are between 100 and 250 µm long. Paraspermatozoa are regarded as infertile and may comprise up to 20% of the mature gametes (Nordheim 1989b). The most characteristic feature of the euspermatozoa is a complex midpiece made up of nine supporting elements arranged in a specific pattern and two inconspicuous mitochondrial derivatives (Figs. 14.5B, 14.6J). The former are absent in paraspermatozoa which possess two long mitochondria with cristae in the midpiece. In both sperm types the acrosome is simple and consists of an acrosomal vesicle and a small basal acrosomal rod. Between midpiece and tail an annulus region with specific substructures is present (Nordheim 1989b). Co-occurrence of two sperm types, euspermatozoa and paraspermatozoa, is rare in Annelida; another example is Tubificinae, a taxon of oligochaetous Clitellata (e.g., Ferraguti et al. 2002). In Saccocirrus there is only one type of spermatozoa of different structure, although it is thread-like as well (Fig. 14.5A). Preliminary electron microscopic observations indicate that they are made up of a short simple acrosome, followed by a comparatively short nucleus and a long midpiece. In the midpiece there are three mitochondria and two supporting rods arranged

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Fig. 14.6 contd

Problematic Annelid Groups

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around the central axoneme (Fig. 14.6I). The tail region is most likely comparatively short. Aphanoneura. The ultrastructure of spermatozoa of Aphanoneura was studied in Aelosoma litorale, A. marcusi, A. singulare and Potamodrilus fluviatilis by Bunke (1985, 1986), Gluzman (1994, 1997) and Marotta et al. (2003). Spermatogenesis follows the general pattern and cytophores with 128 spermatids [termed “spermatozeugmata” by Gluzman (1994)] are formed in A. litorale. Since the coelom is not divided by septa, they are found throughout the animal. Mature spermatozoa in all species are filiform and the acrosome consists of a simple elongated vesicle followed by a cylindrical nucleus. Several differences between the two species have been observed, including the absence of a midpiece in P. fluviatilis where a few rod-shaped mitochondria are situated beneath the nucleus (Fig. 14.6H) and, with respect to the longitudinal axis of the cell, an obliquely positioned distal centriole giving rise to a curved flagellum. In Aeolosoma spp. a short midpiece is present, characterized by a cylindrical mitochondrial derivative originating from two mitochondria during the spermatogenesis (Fig. 14.6 D–F). In P. fluviatilis the posterior part of the nucleus is surrounded by a layer of small vesicles; somewhat larger vesicles are present in the midpiece and the basal nuclear region in Aeolosoma spp. (Bunke 1985, 1986; Fig. 14.6D, E, G). In contrast to Bunke (1985, 1986) Marotta et al. (2003) identified some characters as possible synapomorphies for Aeolosoma and Potamodrilus: the mature spermatozoon of the latter resembles a late spermatid of Aeolosoma. Presence of an acrosomal tube as well as an axial rod as reported by Gluzman (1997), both indicative for clitellate sperm, could not be verified by Bunke (1986) and Marotta et al. (2003). Parergodrilidae. In Parergodrilidae mitotic divisions in the testes produce spermatogonia which are released into the seminal vesicle where they undergo further divisions. In Parergodrilus heideri spermatids develop on small cytophores, each carrying only four developing spermatids (Purschke 2002). In contrast to previous light microscopical observations, Fig. 14.6 contd

Fig. 14.6. Spermatozoa. A–B. Nerillidium troglochaetoides. A. Entire spermatozoon; arrows point to flagellum. B. Enlargement of head showing complex acrosome partly surrounding (arrowheads) nucleus; arrow points to empty apical vesicle. C. Troglochaetus beranecki. Head with small nucleus, several acrosomal vesicles and minute mitochondrion. D–F. Aeolosoma litorale. D. Longitudinal section with nucleus and mitochondrion. Note vesicles surrounding nucleus and mitochondrion. E– F. Cross sections. G–H. Potamodrilus fluviatilis; cross sections. G. Nucleus surronded by single layer of vesicles. H. Mitochondria situated arround basal part of nucleus. I. Saccocirrus sp., Midpiece and tail region; arrowhead points to supporting element. J. Protodrilus purpureus. Midpiece of euspermatozoon with supporting elements (arrowheads). K. Protodriloides symbioticus. Aflagellate spermatozoon with irregularly shaped nucleus, presumed acrosomal vesicles and vacuoles. L–M. Hrabeiella periglanduata. L. Cytophore with late spermatids. M. Midpieces of late spermatids, arrowheads point to accessory tubules, arrows to electron-dense rod. Abbreviations: a, acrosome; av, acrosomal vesicles; c, centriole; cy, cytophore; m, mitochondria; n, nucleus; sp, spermatids; v, vesicles. A–C, G-I, L,M original, D–F courtesy of D. Bunke, J courtesy of H. v. Nordheim, K courtesy of C. Jouin-Toulmond.

$#$ Reproductive Biology and Phylogeny of Annelida the spermatozoa are not of the ect-aquasperm type (Fig. 14.5E): the head comprises an elongated slightly curved acrosome and nucleus, and in the midpiece eight mitochondria surround the distal centriole, which gives rise to the axoneme. A small proximal centriole is oriented parallel to the distal one. Acrosome and tail are set off by two prominent infoldings of the cell membrane. In Stygocapitella subterranea spermatozoa are filiform (Fig. 14.5F) and with a length of about 300 µm they are among the longest spermatozoa observed in annelids. They develop on huge cytophores bearing at least 128 spermatids. The long acrosome only comprises an acrosomal vesicle which is basally folded inwards. In mature spermatozoa the chromatin of the nucleus is not completely condensed. In the long midpiece there is a single circular mitochondrion, which originates by fusion of a multiple number of mitochondria as are present in P. heideri (Purschke 1999; Purschke and Fursman 2005). Hrabeiella. In Hrabeiella periglandulata spermatozoa are filiform as well (Fig. 14.5D; Rota and Lupetti 1997). The elongated conical acrosome consists of an acrosomal vesicle and a central rod-shaped perforatorium. The nucleus is about 25 µm long and has an asymmetric tip extending into the acrosome. The midpiece is made up of a single filiform mitochondrion, the axoneme and seven electron-dense rods surrounding the axoneme. The circle includes the mitochondrion. In addition there are 27 accessory tubules arranged in two groups in the midpiece. The latter are shorter than the electron-dense rods and in late spermatids these accessory tubules branch off from the midpiece (Fig. 14.6M), so that in the seminal vesicles numerous tubules are to be seen among the late spermatids. The tail is comparatively short. The spermatozoa develop on typical cytophores in high numbers (Fig. 14.6L).

14.5 MATING AND FERTILIZATION Dinophilidae. In Dinophilidae transfer of sperm is by direct hypodermic injection through any point of the epidermis and fertilization is internal (Westheide 1990). Each female is inseminated by one or a few males. Opening of the epidermis most likely occurs histolytically by the secretion of the penis glands; in Dinophilus gyrociliatus certain gland cells produce bundles of needle-like structures, which have been suggested as possibly serving to open the epidermis of the female mechanically in the initial phase of copulation (Scharnofske 1984; Westheide 1988). In the dimorphic D. gyrociliatus first copulations take place inside the cocoon with the still juvenile sisters. The life span of the dwarf males is about one to two weeks and most likely they also copulate with non-sister individuals after having left the cocoons (Schmidt and Westheide 1972). Diurodrilidae. In Diurodrilidae no information is available on the reproductive biology, but the structure of sperm indicates some kind of sperm transfer and internal fertilization (Kristensen and Niilonen 1982; Kristensen and Eibye-Jacobsen 1995).

Problematic Annelid Groups

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Nerillidae. The different types of genital structures found in Nerillidae suggest different modes of sperm transfer, as does the occurrence of gonochoristic and hermaphroditic species. However, observations on this are rare. In contrast to many other species of small body size, fertilization mostly appears to be external, since no spermatozoa have been found in females. In Paranerilla limicola bundles of spermatozoa (spermatozeugmata) are released in the mud by the males and then females release their eggs as well (Jouin and Swedmark 1965). It is assumed that many species show various kinds of pseudocopulation characterized by discharge of sperm, either close to the females during the release of eggs, or directly onto the eggs (Jouin and Swedmark 1965; Westheide 1990). This is also supported by the occurrence of immobile sperm (e.g., Troglochaetus beranecki; see above). In Nerilla antennata tiny spermatophores are formed, which are attached to the eggs or deposited in their vicinity (Jouin 1968, 1971). According to Maganini (1982) only the presence of spermatophores on sand grains induces egg deposition in their vicinity. Larger spermatophores are described in Trochonerilla mobilis, but nothing is known about their deposition and fertilization in this species (Tzetlin and Saphonov 1992). Polygordiidae. For Polygordiidae external fertilization and release of gametes into the open water by rupture of the body wall have been suggested (Westheide 1990). Protodrilida. Internal fertilization is characteristic of Saccocirrus and Protodrilus and can be assumed for Parenterodrilus as well. However, sperm transfer in the former is by true copulation and sperm is stored in the seminal receptacules. In S. uchidai synchronous mass spawning has been observed during high tide in the surf zone and eggs are distributed into the water. Although it is suggested that all species have direct transfer of spermatophores in Protodrilus, only in one species, Protodrilus rubropharyngeus, have dorsal organs for spermatophore reception been found. In other species spermatophores are randomly positioned on the epidermis of the females (Nordheim 1983, 1989a, 1991a). In the spermatophores both types of spermatozoa are present (see above). The thin wall of the spermatophores, formed by the secretion of the glands bordering the lateral organs, attaches to the female epidermis, and the sperm content is released through the body wall. The paraspermatozoa are probably involved in histolysis of the tissues during penetration of the body wall, while the euspermatozoa most likely represent the fertilizing gametes (Nordheim 1989b). This is in contrast to the function of the two types of spermatozoa in Tubificinae, in which parasperm form the external sheath of spermatozeugmata (Ferraguti et al. 2002). In both cases eusperm are regarded to be the fertilizing spermatozoa while parasperm have different functions. Whereas most species obviously deposit clusters of adhesive eggs freely on the sediment grains, others produce cocoons secreted by specific glands. In contrast, fertilization is external in Protodriloides (Jouin 1978–79). Large yolky oocytes are laid within a cocoon that is attached to sand grains. In P. symbioticus the cocoon contains four to ten eggs; in

$#& Reproductive Biology and Phylogeny of Annelida P. chaetifer, up to 17. Males discharge aflagellate spermatozoa onto the cocoon that must penetrate this cover in order to fertilize the eggs (Fig. 14.7C–E). Aphanoneura. Sperm transfer or copulation has not been observed in Aphanoneura, but direct transfer of sperm into the seminal receptacles of the partner has been inferred (Bunke 1967, 1986). The reproductive biology of Potamodrilus is virtually unknown (Bunke 1985). Parergodrilidae. In Parergodrilidae the occurrence of seminal receptacles, which always contain spermatozoa in mature females, and the structure of the male genital system, indicate a high probability of direct transfer of sperm and internal fertilization (Rota 1998; Purschke 1999). However, this has never been observed and nothing is known about the function of the genital chaetae in Parergodrilus heideri or of the huge prostate glands in Stygocapitella subterranea. The mode of sperm transfer most likely is different between the two species as seen from their different genital organs. A puzzling feature of P. heideri is the rarity of male individuals: usually females are collected (Rota 1997, 1998; Purschke 1999, 2002). This has led to the assumption that besides gonochoristic individuals there may exist hermaphroditic individuals as well. This was rejected by Reisinger (1960) but the reasons for this phenomenon still remain speculative. Hrabeiella. In Hrabeiella periglandulata reproduction is easily achieved in the laboratory but mating and egg deposition have not been observed (Rota 1998; Rota and Lupetti 1997). Occasionally couples of individuals are seen lying close together, suggesting mating (Rota and Lupetti 1997; Rainer, unpublished). Since neither oviducts nor seminal receptacles have been observed, several possibilities of sperm transfer and mode of fertilization remain conceivable.

14.6 DEVELOPMENT Dinophilidae. Following spawning in Dinophilidae eggs release mucopolysaccharides that expand in seawater; eggs laid at the same time thus stick together and form a common envelope, usually called a cocoon. Development is direct and juveniles hatch with the definitive number of segments (Schmidt and Westheide 1972; Westheide 1990). Dwarf males in Dinophilus gyrociliatus are fully mature upon hatching. Nerillidae. Development in Nerillidae is direct, except in Paranerilla limicola that has planktonic larvae (Jouin and Swedmark 1965; 1971). In Nerilla antennata and other species the eggs are attached to the substratum, singly or in groups. Usually they are covered by a protecting envelope. Many species show a special kind of parental care also known in Syllidae: eggs are not released but attached to the posterior end of the females, where they pass the entire development until the juveniles are set free after having developed several chaetigers (Fig. 14.7A, B; Jouin 1967, 1968, Westheide 1990). So far this kind of brood care has been observed in species of Mesonerilla, Nerillidium and Nerillidopsis. If more than one developing egg is carried, they may be at different stages of development. In Mesonerilla

Problematic Annelid Groups

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Fig. 14.7. Development. A–B. Mesonerilla intermedia. SEM micrographs. A. Female with attached embryo dorsally covered by brood hood. B. Embryo of later stage with developing lateral antennae, median antenna and parapodia (arrowheads); arrow points to adhesive structure anterior to median antenna. C–F. Protodriloides symbioticus. Fertilization and first developing stages. C–D. Oocyte, formation of first polar body, nucleus of spermatozoon (arrow) and aster of oocyte chromosomes (arrowhead). E. Formation of male and female pronucleus and two polar bodies. F. First unequal cleavage leading to small AB and larger CD blastomere, beginning of second clevage; AB blastomere at top of figure. Abbreviations: bh, brood hood; e, embryo; fn, female pronucleus; la, lateral antennae; ma, median antenna; mn, male pronucleus; pb, polar bodies. A–B courtesy of M. C. M. Müller; C–F courtesy of C. Jouin-Toulmond.

intermedia an epidermal brood hood is formed, partly covering the developing eggs. Due to the presence of specialized cells in the attachment zone, a maternal contribution to embryonic nutrition has been suggested (Fransen 1983; Jouin 1968). Polygordiidae. A planktonic trochophore is formed in Polygordius (e.g. Fraipont 1887; Hay-Schmidt 1995). These larvae have a planktotrophic phase that may last several weeks. In certain species an exolarva is formed from this trochophore, which is characterized by serial addition of new segments posterior to the episphere. Other species possess an endolarva in which developing segments are folded up inside the body of the trochophore. At the end of the planktonic phase this larva undergoes metamorphosis during which large parts of the larval body are cast off. Investigations of structure and development of the nervous system in Polygordius lacteus indicate that the entire episphere, including the apical ganglion and the lateral nerves, the ventral nerve cord, the dorsal nerve and the oral nerve plexus, is retained in the adult (Hay-Schmidt 1995). In contrast to species of Protodrilida developing tentacles (palps?) are already present in early trochophores.

$$ Reproductive Biology and Phylogeny of Annelida Protodrilida. Within Protodrilida a planktonic larval stage is typical for Saccocirrus and Protodrilus whereas in Protodriloides development is direct, without a planktonic phase (e.g. Jägersten 1952; Jouin 1962, 1978-79; Sasaki and Brown 1983; Swedmark 1954). These trochophore-like larvae are characterized by an eversible ciliated foregut used for collecting food. In Saccocirrus species two lateral feeding appendages are formed proceeding from the prototroch. These appendages are absent in the group of species lacking a muscular ventral pharynx (Sasaki and Brown 1983). In S. uchadai metatrochophores metamorphose and ultimately comprise about eight segments. In all taxa palps develop in late metatrochophores or early juveniles after a few segments have been formed. In Protodriloides the development is similar in both species (Fig. 14.7E). After a period of 10–20 days in P. symbioticus juveniles with a few segments hatch from the cocoons but palps are visible as two anterior buds only. These stages are still lecitotrophic and 300–500 µm in length in P. symbioticus. Aphanoneura. In Aphanoneura egg deposition and early development have only been observed in Aeolosoma quarternarium (see Bunke 1967). Single eggs are laid within a colorless secretion that sticks the eggs to the substratum. Development is direct and juveniles hatch with six chaetigers. In Potamodrilus fluviatilis single eggs are deposited and hatching juveniles possess only one chaetiger. Parergodrilidae. Parergodrilidae have direct development and juveniles hatch with four chaetigers in both species. Single eggs are released through the female pores and deposited in cocoons that are attached to the substratum (Reisinger 1960; Purschke 1999). Cleavage has been observed in Parergodrilus heideri and it follows the pattern typical of polychaetes with large yolky eggs (Reisinger 1960). Hrabeiella. Development in Hrabeiella periglandulata most likely is direct; the smallest juveniles found possess five chaetigers (Rota 1998). No other observations on the development are available.

14.7 ACKNOWLEDGEMENTS I am grateful to the editors of this volume, Drs Greg Rouse and Fredrik Pleijel, for inviting me to write this contribution. My very cordial thanks are due to Dr. Claude Jouin-Toulmond for various suggestions, comments and discussions. I express my thanks to Professor Westheide, Dr Claude JouinToulmond, Dr Monika C. Müller, Dr Dieter Bunke, Dr Hennig von Nordheim, Inge Nienhüser and Klaus Rainer for unpublished information, material and micrographs. Thanks are also due to Anna Stein, Martina Biedermann and Janina Jördens for various kinds of assistance during preparation of the manuscript.

14.8 LITERATURE CITED Aiyar, R. G. and Alikunhi, K. H. 1944. On some archiannelids of the Madras coast. Proceedings of the National Institute of Science of India 10: 113-140.

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Åkesson, B. 1977. Parasite-host relationships and phylogenetic systematics. The taxonomic position of dinophilids. Pp. 19-28. In W. Sterrer and P. Ax (eds), The Meiofauna Species in Time and Space. Mikrofauna Meeresboden 61. Bleidorn, C., Vogt, L. and Bartolomaeus, T. 2003. New insights into polychaete phylogeny (Annelida) inferred from 18S rDNA sequences. Molecular Phylogenetics and Evolution 29: 279-288. Brinkhurst, R. O. and Nemec, A. F. L. 1987. A comparison of phenetic and phylogenetic methods applied to the systematics of Oligochaeta. Hydrobiologia 155: 65-74. Brown, R. 1981. Saccocirridae (Annelida: Archiannelida) from the central coast of New South Wales. Australian Journal of Marine and Freshwater Research 32: 439-456. Bunke, D. 1967. Zur Morphologie und Systematik der Aeolosomatidae Beddard 1895 and Potamodrilidae nov. fam. (Oligochaeta). Zoologische Jahrbücher für Systematik 94: 187-368. Bunke, D. 1985. Ultrastructure of the spermatozoon and spermiogenesis in the interstitial annelid Potamodrilus fluviatilis. Journal of Morphology 185: 203-216. Bunke, D. 1986. Ultrastructural investigations on the spermatozoon and its genesis in Aeolosoma litorale with considerations on the phylogenetic implications for the Aeolosomatidae (Annelida). Journal of Ultrastructural and Molecular Research 95: 113-130. Eckelbarger, K. 1988. Oogenesis and female gametes. Pp. 281-307. In W. Westheide and C. O. Hermans (eds) The Ultrastructure of Polychaeta. Mikrofauna Marina 4. Eibye-Jacobsen, D. and Kristensen, R. M. 1994. A new genus and species of Dorvilleidae (Annelida, Polychaeta) from Bermuda, with a phylogenetic analysis of Dorvilleidae, Iphitimidae and Dinophilidae. Zoologica Scripta 23: 107-131. Fauchald, K. and Rouse, G. W. 1997. Polychaete systematics: Past and present. Zoologica Scripta 26: 71-138. Ferraguti, M., Marotta, R. and Martin, P. 2002. The double sperm line in Isochaetides (Annelida, Clitellata, Tubificidae). Tissue and Cell 34: 305-314. Fraipont, J. 1887. Le genre Polygordius. Fauna und Flora des Golfes von Neapel 14: 1-125. Fransen, M. E. 1983. Fine structure of the brooding apparatus of the archiannelid Mesonerilla intermedia: Maternal connections to brooded eggs. Transactions of the American Microscopical Society 102: 25-37. Franzén, Å. 1977a. Sperm structure with regard to fertilization biology and phylogenetics. Verhandlungen der Deutschen Zoologischen Gesellschaft 1977: 123-138. Franzén, Å. 1977b. Ultrastructure of spermatids and spermatozoa in Archiannelida. Zoon 5: 97-105. Franzén, Å. and Sensenbaugh, T. 1984. Fine structure of spermiogenesis in the archiannelid Nerilla antennata Schmidt. Videnskabelige Meddelelser fra Dansk Naturhistorisk Forening 145: 23-36. Glasby, C. J., Hutchings, P. A., Fauchald, K., Paxton, H., Rouse, G. W., Watson Russel, C. and Wilson, R. S. 2000. Class Polychaeta. Pp. 1-296. In P. L., Beesly, G. J. B. Ross, and C. J. Glasby, (eds), Polychaetes & Allies: The Southern Synthesis. Fauna of Australia. Vol. 4A. Polychaeta, Myzostoma, Pogonophora, Echiura, Sipuncula. CSIRO Publishing, Melbourne, Australia. Gluzman, C. 1994. A fine structural study of the “spermatozeugmata” of Aeolosoma marcusi (Oligochaeta?). Comm. Biol. 12: 345-355. Gluzman, C. 1997. Sperm cells in Aeolosoma marcusi (Annelida, Oligochaeta). Biocell 21: 137-142.

$$

Reproductive Biology and Phylogeny of Annelida

Goodrich, E. S. 1901. On the structure and affinities of Saccocirrus. Quarterly Journal of Microscopical Science 44: 413-428. Hatschek, B. 1893. System der Anneliden, ein vorläufiger Bericht. Lotus 13: 123-126. Hay-Schmidt, A. 1995. The larval nervous system of Polygordius lacteus Scheinder [sic], 1868 (Polygordiidae, Polychaeta). Immunocytochemical data. Acta Zoologica 76: 121-140. Hempelmann, F. 1906. Zur Morphologie von Polygordius lacteus Schn. und Polygordius triestinus Wolterek, nov. spec. Zeitschrift für wissenschaftliche Zoologie 83: 527-618. Hermans, C. O. 1969. The systematic position of the Archiannelida. Systematic Zoology 18: 85-102. Hessling, R. and Purschke, G. 2000. Immunohistochemical (cLSM) and ultrastructural analysis of the central nervous system and sense organs in Aeolosoma hemprichi (Annelida, Aeolosomatidae). Zoomorphology 120: 65-78. Jägersten, G. 1943. Über den Bau des Kopulationsapparates und den Kopulationsmechanismus bei Dinophilus. Zoologiska Bidrag från Uppsala 22: 61-86. Jägersten, G. 1944. Zur Kenntnis der Morphologie, Encystierung und Taxonomie von Dinophilus. Kungliga Svenska Vetenskapsakademiens Handlinger 21: 1-90. Jägersten, G. 1952. Studies on the morphology, larval development and biology of Protodrilus. Zoologiska Bidrag från Uppsala 29: 425-511. Jamieson, B. G. M. and Rouse, G. W. 1989. The spermatozoa of the Polychaeta. (Annelida): An ultrastructural review. Biological Reviews 64: 93-157. Jördens, J., Struck, T., and Purschke, G. 2004. Phylogenetic inference regarding Parergodrilidae and Hrabeiella periglandulata (“Polychaeta”, Annelida) based on 18S rDNA, 28S rDNA and COI sequencens. Journal of Zoological Systematics and Evolutionary Research 42: 270-280. Jouin, C. 1962. Le développement larvaire de Protodrilus chaetifer Remane (Archiannélides). Comptes Rendus des séances de l’Académie des Sciences, Paris 255: 3065-3067. Jouin, C. 1966. Morphologie et anatomie comparée de Protodrilus chaetifer Remane et Protodrilus symbioticus Giard; création du nouveau genre Protodriloides (Archiannélides). Cahiers de Biologie Marine 7: 139-155. Jouin, C. 1967. Étude morphologique et anatomique de Nerillidopsis hyalina Jouin et de quelques Nerillidium Remane (Archiannélides Nerillidae). Archives de Zoologie Expérimentale et Générale 108: 97-110. Jouin, C. 1968. Sexualité et biologie de la reproduction chez Mesonerilla Remane et Meganerilla Boaden (Archiannélides Nerillidae). Cahiers de Biologie Marine 9: 3152. Jouin, C. 1970. Recherches sur les Protodrilidae (Archiannélides): I. Étude morphologique et systématique du genre Protodrilus. Cahiers de Biologie Marine 11: 367-434. Jouin, C. 1971. Status of the knowledge of the systematics and ecology of Archiannelida. Pp. 47-56. In N. C. Hulings (ed.), Proceedings of the First International Conference on Meiofauna. Smithsonian Contributions to Zoology 76. Jouin, C. 1975. Étude de quelques Archiannélides des côtes d´Afrique du sud; description de Saccocirrus heterochaetus n. sp. (Archiannélide, Saccocirridae). Cahiers de Biologie Marine 16: 97-110. Jouin, C. 1978-79. Spermatozoïde non flagellé et fécondation externe chez Protodriloides symbioticus (Giard) (Annélides, Polychètes, Archiannélides). Vie Milieu 28-29, 473-487.

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Jouin, C. and Rao, G. C. 1987. Morphological studies on some Polygordiidae and Saccocirridae (Polychaeta) from the Indian Ocean. Cahiers de Biologie Marine 28: 389-402. Jouin, C. and Swedmark, B. 1965. Paranerilla limnicola n. g., n. sp., Archiannélide Nerillidae du benthos vaseux marin. Cahiers de Biologie Marine 6: 201-218. Jouin-Toulmond, C. and Purschke, G. 2004. Ultrastructural study of the spermatozoa of Parenterodrilus taenioides (Protodrilida: “Polychaeta”) and its phylogenetic significance in the Protodrilida. Zoomorphology 123: 139-146. Karling, T. G. 1958. Zur Kenntnis von Stygocapitella subterranea Knöllner und Parergodrilus heideri Reisinger (Annelida). Arkiv för Zoologi 11: 307-342. Kristensen, R. M. and Eibye-Jacobsen, D. 1995. Ultrastructure of spermiogenesis and spermatozoa in Diurodrilus subterraneus (Polychaeta, Diurodrildae). Zoomorphology 115: 117-132. Kristensen, R. M. and Niilonen, T. 1982. Structural studies on Diurodrilus Remane (Diurodrilidae fam. n.) with description of Diurodrilus westheidei sp. n. from Arctic interstitial meiobenthos, W. Greenland. Zoologica Scripta 11: 1-12. Maganini, G. 1982. Reproduction in Nerilla antennata O. Schmidt (Archiannelida, Nerillidae): induction of spawning. Bolletino di Zoologica 49: 283-286. Marion, A. F. and Bobretzky, N. 1875. Étude des Annélides du golfe de Marseille. Annales des Sciences naturelles. Paris, Series 6, 2: 1-106. Marotta, R., Ferraguti, M. and Martin, P. 2003. Spermiogenesis and seminal receptacles in Aeolosoma singulare (Annelida, Polychaeta, Aeolosomatidae). Italian Journal of Zoology 70: 123-132. Martin, F. and Traut, W. 1987. The mode of sex determination in Dinophilus gyrociliatus (Archiannelida). International Journal of Invertebrate Reproduction and Development 11: 159-172. Mock, H. 1981. Zur Kenntnis von Diurodrilus subterraneus (Polychaeta, Dinophilidae) aus dem Sandhang der Nordseeinsel Sylt. Helgoländer Meeresuntersuchungen 34: 329-335. Müller, M. C. 1999. Das Nervensystem der Polychaeten: Immunhistochemische Untersuchungen an ausgewählten Taxa. Ph.D. Dissertation, University of Osnabrück, Germany. Müller, M. C. M. 2002. Aristonerilla: a new nerillid genus (Annelida: Polychaeta) with description of Aristonerilla (Micronerilla) brevis comb. nov. from a seawater aquarium. Cahiers de Biologie Marine 43: 131-139. Müller, M. C. M. and Westheide, W. 2002. Comparative analysis of the nervous system in presumptive progenetic dinophilid and dorvilleid polychaetes (Annelida) by immunohistochemistry and cLSM. Acta Zoologica 83: 33-48. Nordheim, H. von 1983. Systematics and ecology of Protodrilus helgolandicus sp. n., an interstitial polychaete (Protodrilidae) from subtidal sands off Helgoland, German Bight. Zoologica Scripta 12: 171-177. Nordheim, H. von 1984. Life histories of subtidal interstitial polychaetes of the families Polygordiidae, Protodrilidae, Nerillidae, Dinophilidae and Diurodrilidae from Helgoland (North Sea). Helgoländer Meeresuntersuchungen 38: 1-20. Nordheim, H. von 1989a. Six new species of Protodrilus (Annelida, Polychaeta) from Europe and New Zealand, with a concise presentation of the genus. Zoologica Scripta 18: 245-268. Nordheim, H. von 1989b. Vergleichende Ultrastrukturuntersuchungen der Eu- und Paraspermien von 13 Protodrilus-Arten (Polychaeta, Annelida) und ihre taxonomische und phylogenetische Bedeutung. Helgoländer Meeresuntersuchungen 43: 113-156.

$$" Reproductive Biology and Phylogeny of Annelida Nordheim, H. von 1991a. Ultrastructure and functional morphology of the female reproductive organs in Protodrilus (Polychaeta, Annelida). Helgoländer Meeresuntersuchungen 45: 465-485. Nordheim, H. von 1991b. Ultrastructure and functional morphology of male genital organs and spermatophore formation in Protodrilus (Polychaeta, Annelida). Zoomorphology 111: 81-94. Orrhage, L. 1974. Über die Anatomie, Histologie und Verwandtschaft der Apistobranchidae (Polychaeta, Sedentaria) nebst Bemerkungen über die systematische Stellung der Archianneliden. Zeitschrift für Morphologie der Tiere 79: 1-45. Pierantoni, U. 1908. Protodrilus. Fauna und Flora des Golfes von Neapel 31: 1-226. Purschke, G. 1985a. Anatomy and ultrastructure of ventral pharyngeal organs and their phylogenetic importance in Polychaeta (Annelida). I. The pharynx of the Dinophilidae. Zoomorphology 105: 223-239. Purschke, G. 1985b. Anatomy and ultrastructure of ventral pharyngeal organs and their phylogenetic importance in Polychaeta (Annelida). II. The pharynx of the Nerillidae. Mikrofauna Marina 2: 23-60. Purschke, G. 1986. Ultrastructure of the nuchal organ in the interstitial polychaete Stygocapitella subterranea (Parergodrilidae). Zoologica Scripta 15: 13-20. Purschke, G. 1990a. Comparative electron microscopic investigation of the nuchal organs in Protodriloides, Protodrilus and Saccocirrus (Annelida, Polychaeta). Canadian Journal of Zoology 68, 325-338. Purschke, G. 1990b. Ultrastructure of the “statocysts” in Protodrilus species (Polychaeta): Reconstruction of the cellular organization with morphometric data from receptor cells. Zoomorphology 110: 91-104. Purschke, G. 1992. Ultrastructural investigations of presumed photoreceptive organs in two Saccocirrus species (Polychaeta: Saccocirridae). Journal of Morphology 211: 7-21. Purschke, G. 1993. Structure of the prostomial appendages and the central nervous system in the Protodrilida (Polychaeta). Zoomorphology 113: 1-20. Purschke, G. 1999. Terrestrial polychaetes—models for the evolution of the Clitellata (Annelida)? Hydrobiologia 406: 87-99. Purschke, G. 2000. Sense organs and the central nervous system in an enigmatic terrestrial polychaete, Hrabeiella periglandulata (Annelida)—implications for annelid evolution. Invertebrate Biology 119: 329-341. Purschke, G. 2002. Male genital organs, spermatogenesis and spermatozoa in the enigmatic terrestrial polychaete Parergodrilus heideri (Annelida, Parergodrilidae). Zoomorphology 121: 125-138. Purschke, G. 2003. Is Hrabeiella periglandulata (Annelida, “Polychaeta”) the sister group of Clitellata? Evidence from an ultrastructural analysis of the dorsal pharynx in H. periglandulata and Enchytraeus minutus (Annelida, Clitellata). Zoomorphology 122: 55-66. Purschke, G. and Hessling, R. 2002. Analysis of the central nervous system and sense organs in Potamodrilus fluviatilis (Annelida: Potamodrilidae). Zoologischer Anzeiger 241: 19-35. Purschke, G. and Jouin, C. 1988. Anatomy and ultrastructure of the ventral pharyngeal organs of Saccocirrus and Protodriloides with remarks on the phylogenetic relationships within the Protodrilida (Annelida, Polychaeta). Journal of Zoology 215: 405-432. Purschke, G. and Jouin-Toulmond, C. 1993. Ultrastructure of presumed ocelli in Parenterodrilus taenioides (Polychaeta, Protodrilidae) and their phylogenetic significance. Acta Zoologica 74: 247-256.

Problematic Annelid Groups

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Purschke, G. and Jouin-Toulmond, C. 1994. Ultrastructure of the sense organs and the central nervous system in Parenterodrilus taenioides and their phylogenetic significance in Protodrilida (Annelida, Polychaeta). Pp. 119-128. J. -C. Dauvin, L. Laubier and D. J. Reish (eds), Actes de la 4ème Conférence Internationale des Polychètes. Mémoires de Muséum Nationale d’Histoire Naturelle, Paris 162. Purschke, G. and Müller, M. C. 1996. Structure of prostomial photoreceptor-like sense organs in Protodriloides species (Polychaeta, Protodrilida). Cahiers de Biologie Marine 37: 205-219. Purschke, G. and Fursman, M. 2005. Spermatogenesis and spermatozoa in Stygocapitella subterranea (Annelida, Parergodrilidae), an enigmatic supralittoral polychaete. Zoomorphology 124: 137-148. Purschke, G., Hessling, R. and Westheide, W. 2000. The phylogenetic position of the Clitellata and Echiura—on the problematic assessment of absent characters. Journal of Zoological Systematics and Evolutionary Research 38: 165-173. Reisinger, E. 1925. Ein landbewohnender Archiannelide. Zeitschrift für Morphologie und Ökologie der Tiere 3: 197-254. Reisinger, E. 1960. Die Lösung des Parergodrilus-Problems. Zeitschrift für Morphologie und Ökologie der Tiere 48: 517-544. Riser, N. W. 1999. Description of a new species of dinophilid polychaete, with observation on other dinophilids and interstitial polychaetes in New England. Northeastern Naturalist 6: 211-220. Rota, E. 1997. First Italian record of the terrestrial polychaete Parergodrilus heideri Reisinger, with anatomical and ecological notes. Italian Journal of Zoology 64: 9196. Rota, E. 1998. Morphology and adaptations of Parergodrilus Reisinger and Hrabeiella Pizl & Chalupsky, two enigmatic soil-dwelling annelids. Italian Journal of Zoology 65: 75-84. Rota, E. and Lupetti, P. 1997. An ultrastructural investigation of Hrabeiella Pizl and Chalupsky, 1984 (Annelida). II. The spermatozoon. Tissue and Cell 29: 603-609. Rota, E., Martin, P. and Erséus, C. 2001. Soil-dwelling polychaetes: enigmatic as ever? Some hints on their phylogenetic relationships as suggested by a maximumparsimony analysis of 18S rRNA gene sequences. Contributions to Zoology 70: 127-138. Rouse, G. W. and Fauchald, K. 1997. Cladistics and polychaetes. Zoologica Scripta 26: 139-204. Rouse, G. W. and Jamieson, B. G. M. 1987. An ultrastructural study of the spermatozoa of the polychaetes Eurythoe complanata (Amphinomidae), Clymenella sp. and Micromaldane sp. (Maldanidae), with definition of sperm types in relation to reproductive biology. Journal of Submicroscopic Cytology 19: 573-584. Rouse, G. W. and Pleijel, F. 2001. Polychaetes. Oxford University Press, New York, USA. 354 pp. Sasaki, S. and Brown, R. 1983. Larval development of Saccocirrus uchidai from Hokkaido, Japan, and Saccocirus krusadensis from New South Wales, Australia (Archiannelida, Saccocirridae). Annotationes Zoologicae Japonenses 56: 299-314. Scharnofske, P. 1984. Anatomie, Ultrastruktur und Funktion der männlichen Geschlechtsorgane der Dinophilidae und Histriobdellidae (Annelida, Polychaeta). Ph.D. Dissertation, University of Göttingen, Germany. Scharnofske, P. 1986. Ultrastructure of sperm morphology of Trilobodrilus axi and T. heideri (Dinophilidae, Polychaeta) Helgoländer Meeresuntersuchungen 40: 419-430. Schmidt, P. and Westheide, W. 1972. Dinophilus gyrociliatus (Polychaeta). Nahrungsaufnahme und Fortpflanzung. Encyclopedia Cinematographica E 1750: 1-16.

$$$ Reproductive Biology and Phylogeny of Annelida Struck, T. 2003. Progenetische Evolution als Prinzip zur Entstehung neuer Arten innerhalb der Polychaeten am Beispiel der Dinophilidae/”Dorvilleidae” (“Polychaeta”, Annelida). Ph.D. Dissertation, University of Osnabrück, Germany. Struck, T. H. and Purschke, G. 2005. The sister group relationship of Aeolosomatidae and Potamodrilidae (Annelida, “Polychaeta”)—a molecular phylogenetic approach based on 18S rDNA and Cytochrome Oxidase I. Zoologischer Anzeiger 243: 281-293. Struck, T., Westheide, W. and Purschke, G. 2002a. Progenesis in Eunicida (“Polychaeta”, Annelida)—separate evolutionary events? Evidence from molecular data. Molecular Phylogenetics and Evolution 25: 190-199. Struck, T. Hessling, R. and Purschke, G. 2002b. The phylogenetic position of Aeolosomatidae and Parergodrilidae, two enigmatic oligochaete-like taxa of “Polychaeta”. Journal of Zoological Systematics and Evolutionary Research 40: 155-163. Struck, T. H., Halanych, K. M. and Purschke, G. (2005) Dinophilidae (Annelida) is not a progenetic Eunicida; evidence from 18S and 28S rDNA. Molecular Phylogenetics and Evolution (in press). Swedmark, B. 1954. Étude du développement lavaire et remarques sur la morphologie de Protodrilus symbioticus Giard (Archiannélides). Arkiv för Zoologi 6: 511-522. Timm, T. 1981. On the origin and evolution of aquatic Oligochaeta. Eesti NSV Teaduste Akadeemia Toimetised Bioloogia. 30: 174-181. Traut, W. 1969. Zur Sexualität von Dinophilus gyrociliatus (Archiannelida). II. Der Aufbau des Ovars und die Oogenese. Biologisches Zentralblatt 88: 695-714. Tzetlin, A. B. and Saphonov, M. V. 1992. Trochonerilla mobilis gen. et sp. n., a meiofaunal nerillid (Annelida, Polychaeta) from a marine aquarium in Moscow. Zoologica Scripta 21: 251-254. Westheide, W. 1971. Apharyngtus punicus nov. gen. nov. spec., ein aberranter Archiannelide aus dem Mesopsammal der tunesischen Mittelmeerküste. Mikrofauna des Meeresbodens 6: 1-19. Westheide, W. 1979. Ultrastruktur der Genitalorgane interstitieller Polychaeten. II. Männliche Kopulationsorgane mit intrazellulären Stilettstäben in einer Microphthalmus-Art. Zoologica Scripta 8: 111-118. Westheide, W. 1982. Ikosipodus carolinensis gen. et sp. n., an interstitial neotenic polychaete from North Carolina, USA, and its phylogenetic relationships within Dorvilleidae. Zoologica Scripta 11: 117-126. Westheide, W. 1985. The systematic position of the Dinophilidae and the archiannelid problem. Pp. 310-326. In S. C. Morris, J. D. George, R. Gibson and H. M. Platt (eds), The Origins and Relationships of Lower Invertebrates. The Systematics Association Special Volume 28. Clarendon Press, Oxford, UK. Westheide, W. 1988. Genital organs. Pp. 263-279. In W. Westheide and C. O. Hermans (eds) The Ultrastructure of Polychaeta. Mikrofauna Marina 4. Westheide, W. 1990. Polychaetes: Interstitial families. Synopsis of the British Fauna. No. 44. D. M. Kermack and R. S. K. Barnes (Series eds). Universal Book Services/Dr. W. Backhuys, Oegstgeest, The Nederlands, 152 pp. Westheide, W. and Purschke, G. 1996. Leptonerilla diplocirrata, a new genus and species of interstitial polychaetes from the island of Hainan, south China (Nerillidae). Proceedings of the Biological Society of Washington 109: 586-590. Westheide, W. and Riser, N. W. 1983. Morphology and phylogenetic relationships of the neotenic interstitial polychaete Apodotrocha progenerans n. gen., n. sp., Annelida. Zoomorphology 103: 67-87.

Problematic Annelid Groups

$

%$Westheide, W. and Schmidt, P. 1974. Trilobodrilus axi (Polychaeta). Nahrungsaufnahme und Fortpflanzung. Encyclopedia Cinematographica E 1955: 1-12. Windoffer, R. and Westheide, W. 1988. The nervous system of the male Dinophilus gyrociliatus (Annelida: Polychaeta). II. Electron microscopical reconstruction of nervous anatomy and effector cells. Journal of Comparative Neurology 272: 475488. Worsaae, K. 2005. Phylogeny of Nerillidae (Polychaeta, Annelida) as inferred from combined 18S rDNA and morphological data. Cladistics 21: 143-162. Worsaae, K. and Kristensen, R. M. 2003. A new species of Paranerilla (Polychaeta: Nerillidae) from northeast Greenland waters, Arctic Ocean. Cahiers de Biologie Marine 44: 23-39. Worsaae, K. and Müller, M. C. M. 2004. Nephridial and gonoduct distribution patterns in Nerillidae (Annelida: Polychaeta)—examined by tubulin staining and cLSM. Journal of Morphology 261: 259-269. Worsaae, K., Nygren, A., Rouse, G. W., Giribet, G., Persson, J., Sundberg, P. and Pleijel, F. 2005. Phylogenetic position of Nerillidae and Aberranta (Polychaeta, Annelida), analysed by direct optimization of combined molecular and morphological data. Zoologica Scripta 34: 313-328.

Index 16S ribosomal DNA 251 18S rRNA 5, 235, 236 28S rDNA 57, 237, 238, 253, 256, 257, 263, 264, 397

A ABd-b 124 Abdominal Cilia 157, 166 Aberranta 10, 13, 156, 640 Acanthobdella 235, 237, 248, 249, 250, 261, 350, 393, 404 Acanthobdellida 6, 7, 236, 244, 248, 249, 260, 261, 300, 379, 394, 423 Acanthobdellidae 244 Acanthobdellids 235, 252, 338, 339 Acanthodrilin Condition 274, 326, 327, 365 Acanthodrilinae 237, 261, 262, 263, 265, 281, 282, 325, 326, 327, 362, 363, 365 Acanthodrilus 273, 365 Accessory Glands 297, 298 Achaeta 248, 249, 251 Acholoe 452 Aciculae 9, 10, 475 Aciculata 9, 10, 13, 17, 46, 47, 48, 66, 154, 155, 156, 162, 164, 431, 640 Acoetidae 13, 155, 438 Acrocirridae 8, 11, 13, 48, 156, 497, 498, 500, 501, 502 Acrocirrus 48, 498, 500, 504, 508, 509 Acrosomal Vesicle 464, 651, 652, 653, 656 Acrosome 53, 55, 57, 58, 59, 60, 62, 252, 261, 265, 266, 267, 329, 331, 333, 335, 336, 337, 339, 340, 341, 342, 343, 345, 346, 348, 350, 351, 352, 354, 355, 356, 404, 434, 461, 462, 463, 464, 465, 466, 467, 508, 509, 515, 527, 530, 532, 533, 537, 538, 579, 580, 651, 653, 655, 656 — morphogenesis 336 — protube 404 — rod 336, 343, 346, 348, 352, 356, 464, 653 — tube 57, 58, 261, 265, 267, 333, 335, 336, 337, 349, 340, 342, 343, 345, 348, 350, 351, 352, 354, 355, 356, 404, 405, 655 Actin 333, 360, 367 Adelphophagia 607, 624, 626, 627

Adenodrilus 253, 303 Adhesive Gland 645 Adolescent Male Phase 82 Aeolosoma 52, 58, 59, 248, 249, 250, 340, 642, 655, 660 Aeolosomatidae 8, 12, 13, 52, 53, 56, 58, 59, 65, 143, 147, 157, 639, 647 African 240, 255, 258, 274, 398 Aglaophamus 442, 474 Agriodrilus 249, 301, 302 Ailoscolex 256, 316, 322, 323 Ailoscolecidae 255, 256, 258, 260, 261, 322, 323, 325 Ainudrilus 251 Akrotroch 142, 143, 148, 475, 480 Alae 319, 320, 321, 363 Albinaria 249 Albumen 240, 269, 368, 377, 408 Albumenotrophy 240, 269, 368, 408 Alciopa 48, 448, 461 Alciopidae 24, 25, 38, 39 Alciopina 48 Alciopini 39, 432, 466 Alentia 48, 452, 461 Aliolimnatis 402, 403 Allolobophora 239, 329, 331, 350, 351, 352, 361 Alluroididae 240, 245, 254, 255, 258, 260, 280, 281, 293, 304, 307, 309, 311, 312 Alluroidina 254, 260, 309 Alluroidoidea 243, 254, 309 Alma 256, 257, 270, 271, 274, 275, 278, 281, 319, 321, 363, 366 Almidae 244, 246, 255, 256, 257, 259, 261, 270, 275, 278, 280, 281, 291, 311, 319, 320, 366 Alminae 256, 257, 271, 319 Almoidea 256, 257, 261, 270 Alphadrilus 253 Alvinella 49 Alvinellidae 11, 13, 49, 56, 153, 156 Amblyosyllis 14, 455 Americobdella 396, 399, 401, 402 Americobdellidae 395, 401 Ampharetidae 11, 13, 24, 25, 49, 52, 144, 153, 156, 165 Amphicorina 50, 524, 526, 535, 539 Amphiduros 432

$% Reproductive Biology and Phylogeny of Annelida Amphiglena 50, 523, 529, 532, 535, 539, 551 Amphinomida 9, 13, 17, 46, 65, 154, 160, 431 Amphinomidae 9, 13, 46, 55, 144, 153, 154, 163, 165 Amphipolydora 571, 584, 626, 628, 629, 630 Amphisamytha 34, 49 Amphitrite 99, 165, 220 Amphitritides 249 Ampulla 284, 285, 287, 288, 307, 309, 312, 315, 326, 536, 537, 547 Amynthas 263, 266, 270, 288, 289, 331, 345, 352, 361 Anaitides 149, 163, 480 Anchoring Apparatus 58, 59, 333, 338, 509 Ancistrosyllis 452, 480 Annelid Cross 101 Annelida 3, 4, 5, 6, 7, 8, 12, 13, 16, 17, 23, 24, 32, 38, 39, 45, 46, 52, 53, 55, 57, 58, 62, 64, 65, 66, 77, 78, 82, 84, 93, 94, 95, 96, 97, 98, 99, 100, 101, 104, 107, 108, 109, 110, 114, 115, 116, 117, 120, 121, 122, 123, 124, 128, 130, 131, 132, 133, 141, 142, 143, 144, 146, 147, 151, 153, 154, 157, 158, 160, 166, 179, 180, 181, 182, 233, 235, 236, 237, 239, 247, 250, 331, 344, 360, 365, 393, 403, 406, 408, 412, 420, 421, 432, 433, 435, 436, 461, 463, 550, 574, 577, 639, 651, 653, 656 Annual Iteroparity 24 Annulus 651, 653 Antennae 9, 17, 472, 474, 475, 480, 482, 659 Antimetastatic 398 Aonidella 568, 569, 571, 575, 582 Aonides 566, 567, 568, 569, 571, 575, 576, 579, 582, 589 Aphanoneura 248, 249, 340, 641, 642, 647, 655, 658, 660 Apharyngtus 642 Aphelochaeta 498, 506, 507, 508, 511, 513, 515 Aphroditidae 13, 17, 24, 47, 56, 143, 155, 438 Aphroditiformia 10, 431, 467 Aphropharynx 498, 501, 512 Apical Organ 104, 141, 142 Apical Tuft 102, 103, 105, 106, 141, 142, 144, 148, 149, 472, 474, 480 Apistobranchidae 565, 568, 569 Apistobranchus 11, 13, 157, 565, 575 Apodotrocha 46, 58 Apomorphy 4, 5, 8, 9, 10, 253, 254, 266, 303, 327, 464, 521 Aporrectodea 351 Aquamegadrili 244, 255, 256, 257 Aquasperm 55, 56, 438, 439, 441, 443, 444, 445, 446, 447, 448, 449, 450, 451, 453, 454, 455, 456, 457, 458, 459, 460, 461, 462, 463, 464, 466, 467, 471, 532, 533, 571, 578, 579, 580, 651, 653, 656 Archenteron 368

Archiannelida 639 Architomy 498, 511, 512, 628, 629 Arctonoe 48, 452 Areco 319 Arenicola 46, 66, 99, 151, 161, 504, 510 Arenicolidae 8, 12, 13, 24, 46, 52, 151, 153, 154, 160, 161 Arhynchobdellida 292, 394 Armandia 46, 145, 158, 165 Articulata 4, 122 Asexual Reproduction 180, 296, 498, 506, 510, 511, 512, 550, 565, 566, 607, 626, 628, 628, 630, 631 Astacopsidrilus 239, 333, 343, 345 Asymmetric Cell Division 99 Asynchronous — cleavage 371 — development 511 — reproduction 507 Atherospio 568, 569, 571 Atokous 455, 457, 468, 511 Atria 261, 281, 296, 297, 298, 299, 300, 303, 309, 312, 400, 401, 402, 403 Atrium 269, 272, 284, 294, 295, 296, 297, 298, 299, 300, 301, 307, 311, 315, 399, 400, 401, 403, 648 Atypical — arrangement of testis-sacs 401 — cysts 333 — spermatozoa 352 Augeneriella 50, 54, 160 Aulodrilus 292 Aulophore 147, 151 Autapomorphy 303, 336 Autolytus 48, 455, 460, 465, 471, 480 Autosynthesis 32, 37, 529, 573 Axoneme 58, 60, 63, 333, 335, 338, 339, 341, 342, 345, 348, 350, 351, 355, 356, 461, 463, 464, 465, 466, 467, 527, 532, 651, 652, 653, 655, 656

B Bacescuella 290 Balantin Condition 326 Bandlet 117, 118, 119, 130, 371, 373, 374, 375, 410, 412, 415 Basal Body 267, 338, 339, 356, 360, 404, 466 Bathydrilus 251, 272, 292, 339 Bathynoe 84 Batracobdelloides 398 Begemius 263, 293 Behavior 68, 81, 84, 149, 180, 182, 190, 193, 195, 197, 198, 215, 217, 221, 363, 403, 465, 469, 582, 590, 595, 622 Bifid Chaetae 298 Biogeography 240, 398

Index Biramous 17, 472 Bispira 50, 550 Biwadrilidae 244, 255, 256, 260, 311, 313 Biwadriloidea 256, 260 Biwadrilus 256, 270, 271, 280, 281, 311, 313, 321 Blades 507, 515 Blast Cells 109, 113, 114, 117, 118, 119, 121, 130, 371, 373, 375, 376, 410, 411, 412, 415, 416, 417, 418, 419 Blastocoel 111, 377 Blastomeres 95, 96, 99, 100, 101, 102, 104, 105, 106, 107, 108, 109, 110, 111, 416, 626 Blastulae 184, 367 Blood feeding 394, 396, 398 Blood Vessels 24, 27, 377, 433, 437, 463, 469, 505, 510, 526, 573 Boccardia 189, 192, 571, 573, 574, 584, 612, 613, 618, 619, 620, 623, 626, 627 Boccardiella 49, 571, 579 Body Size 81, 83, 84, 85, 87, 89, 550, 657 Bollandia 455 Bonellia 52, 66 Bothrioneurum 290 Brachiopoda 5, 6 Brachyenteron 113, 115 Brachyury 113, 114, 115 Brada 501 Branchiobdella 37, 337 Branchiobdellida 3, 6, 7, 235, 236, 237, 240, 248, 249, 251, 252, 260, 261, 266, 267, 292, 293, 300, 338, 339, 350, 393, 394, 404 Branchiomma 50, 82, 529, 539, 550 Branchipolynoe 84, 85 Branchiura 249, 333 Brania 82, 87, 455, 460, 465 Brdu 120 Broadcast Spawning 57, 441, 442, 462, 468, 471, 527, 528, 532, 533, 535, 537, 582 Brood 78, 87, 89, 179, 187, 189, 190, 191, 192, 406, 453, 454, 461, 475, 480, 498, 508, 511, 523, 533, 534, 535, 536, 537, 540, 541, 542, 544, 545, 546, 547, 576, 586, 604, 605, 606, 658, 659 — care 498, 658 — protection 190, 508, 511, 576 Brooding 77, 83, 88, 151, 191, 406, 447, 448, 450, 456, 472, 474, 475, 482, 511, 513, 528, 531, 533, 535, 536, 537, 538, 539, 540, 541, 542, 543, 544, 546, 549, 550, 579, 586, 595, 604, 623, 624 Bythonomus 265, 266, 301, 337, 346, 347, 348, 358

C Cabira 452 Calamyzas 48, 455, 466

$%

Callidrilus 256, 281, 282, 319, 321 Cambarincola 248, 249, 250, 350 Cambrian 12, 17 Canadia 12 Canalipalpata 9, 10, 11, 13, 48, 49, 50, 51, 52, 156, 157, 162, 164 Caobangia 522, 524, 529, 535, 539 Capilloventridae 245, 252, 260, 296, 298, 343, 344 Capitella 27, 30, 31, 32, 33, 34, 35, 36, 46, 66, 85, 86, 94, 98, 99, 100, 107, 112, 120, 121, 123, 195, 202, 220, 221, 576 Capitellidae 8, 12, 13, 24, 46, 52, 146, 153, 154, 160, 161, 166, 435 Capitellides 46 Capitomastus 46, 82 Capitulum 265, 342, 345, 348 capricornia 47, 440, 462, 468 Carazziella 571, 619, 620 Carboniferous 12 Catastrophic Metamorphosis 141, 149, 152, 548 Cdc25 413, 416 Cell Ablations 93, 119, 408 Cell Fate 95, 96, 99, 101, 104, 106, 108, 111, 112, 113, 114, 132, 408, 411, 413, 415, 421 Cell Lineages 93, 99, 117, 368, 408 Cell Specification 108 Central Sheath 57, 339, 341, 343, 345, 346, 348 Centriole 59, 63, 267, 333, 335, 336, 337, 338, 340, 351, 360, 367, 434, 462, 463, 465, 509, 655, 656 Ceratonereis 443 Chaetae 4, 5, 9, 10, 11, 17, 81, 85, 102, 103, 105, 117, 145, 149, 151, 152, 158, 160, 182, 195, 196, 246, 251, 252, 253, 271, 272, 273, 281, 298, 309, 311, 316, 319, 321, 323, 324, 325, 362, 363, 365, 393, 469, 472, 474, 480, 482, 497, 502, 503, 511, 512, 514, 516, 548, 566, 576, 585, 587, 589, 592, 593, 594, 595, 596, 598, 599, 600, 601, 602, 604, 605, 606, 609, 611, 612, 614, 615, 616, 619, 620, 621, 622, 624, 625, 642, 647, 648, 658 Chaetal Sacs 373 Chaetopteridae 8, 10, 11, 13, 49, 143, 147, 148, 153, 157, 165, 521, 565, 567, 568, 569, 601, 603 Chaetopterus 49, 95, 96, 98, 99, 101, 102, 103, 106, 121, 123, 124, 125, 126, 127, 128, 129, 131, 146, 148, 165, 421, 601, 602, 603 Chaetosphaera 147, 149, 151 Chauvinelia 500, 504 Ch-en 121, 123 Chitinopoma 51, 66, 523, 530, 532, 533, 536, 541 Chloragocytes 37, 328 Chloritis 249, 250 Chondrichthyes 340



%$Reproductive Biology and Phylogeny of Annelida

Chone 165, 524, 526, 539 Chromatin 29, 63, 337, 355, 356, 357, 360, 404, 463, 656 Chromatoid Bodies 335 Chrysopetalidae 13, 17, 47, 56, 143, 153, 155, 163, 431, 432, 435, 438, 467, 472, 474 Chrysopetalum 47, 438, 472 Chv- hox1 124, 125, 126, 127, 128 Chv-hox2 98, 125, 127, 128 Chv-hox3 125, 126, 127 Chv-hox4 125, 127, 131 Chv-hox5 125, 126, 127, 132 Ciliated Food Groove 144, 146, 151, 165 Ciliated Funnel 355, 644 Circeis 527, 531, 533, 536, 544, 549 Cirratulida 501 Cirratuliformia 11, 13, 48, 49, 65, 156, 497, 500, 503, 506, 507, 515 Cirratulus 15, 498, 504, 507, 511, 514 Cirri 8, 9, 10, 97, 469, 472, 474, 475, 480, 482, 590, 593, 624, 630, 643 Cirriformia 27, 49, 65, 148, 165, 498, 511 Cirrophorus 569 Cistenides 49 Cladistic Analysis 56, 160, 250, 346, 501, 503, 522, 524 Cladogram 16, 17, 65, 241, 242, 396, 503, 537 Claspers 257, 274, 278, 319, 320, 321, 363, 366 Classification 7, 9, 181, 237, 240, 242, 245, 259, 260, 262, 263, 264, 267, 268, 393, 402, 435, 504, 524, 566, 567, 568, 569, 571, 573, 640 Cleavage 93, 95, 96, 97, 99, 100, 101, 102, 103, 104, 106, 107, 108, 111, 112, 117, 118, 121, 141, 240, 366, 367, 368, 369, 371, 377, 408, 409, 413, 415, 419, 472, 592, 659, 660 Clepsine 99 Clitellata 5, 6, 7, 8, 12, 13, 14, 24, 27, 37, 45, 46, 52, 56, 57, 58, 64, 65, 80, 88, 143, 146, 147, 154, 160, 179, 235, 236, 237, 239, 260, 261, 267, 288, 293, 339, 340, 393, 394, 435, 641, 642, 653 Clitellate Relationships 236 Clitellum 8, 57, 235, 240, 252, 255, 259, 261, 268, 269, 270, 271, 274, 275, 290, 291, 296, 298, 305, 306, 309, 311, 312, 315, 316, 319, 321, 322, 323, 324, 326, 328, 361, 362, 363, 399, 400, 406, 641 Closed Circulatory System 433 Clumped Spatial Distribution 83 Clymenella 46, 148, 161 Coagulation 398 Cocoon 64, 151, 160, 235, 240, 268, 269, 270, 275, 284, 288, 290, 315, 321, 327, 328, 329, 339, 349, 350, 361, 366, 368, 399, 406, 407, 408, 447, 582, 597, 645, 648, 656, 657, 658 Coeloblastulae 110 Coelom 4, 25, 26, 27, 36, 38, 39, 45, 53, 281, 302, 312, 315, 322, 325, 327, 328, 331, 333, 373,

435, 437, 461, 463, 468, 504, 507, 509, 510, 511, 513, 514, 516, 526, 528, 529, 530, 573, 574, 575, 576, 579, 604, 643, 650, 655 Coelomocytes 239, 529, 573 Coelomoducts 268, 377, 435, 504, 525 Coelomostomes 435 COI 57, 235, 237, 246, 247, 249, 250, 251, 340 Coition 361, 362, 365 Colinearity 124, 125, 128, 130, 420 Conducting (Vector) Tissue 400 Connectives 252, 253, 265, 416 Continuous Iteroparity 24 Copulation 25, 78, 81, 86, 87, 271, 274, 281, 284, 292, 311, 361, 363, 366, 398, 406, 440, 441, 444, 447, 450, 452, 457, 458, 462, 468, 469, 471, 578, 656, 657, 658 — gland 311 Copulatory Organ 58, 298, 299, 642, 643, 647, 649 Coralliodrilus 339, 345, 350 Cortical Alveoli 574, 576, 577, 578, 587, 598 Cortical Granules 328 Cossura 46, 569 Cossuridae 13, 46, 154, 500 Coupling Chaetae 363 Courtship 78, 80, 363 Covariation 550 Crassiclitellata 237, 240, 243, 245, 246, 247, 250, 253, 254, 255, 256, 258, 260, 269, 271, 274, 293, 311, 327, 329, 343, 368, 369 Cretaceous 13 Criodriliini 257 Criodrilinae 257, 291, 319, 321 Criodrilus 244, 246, 255, 256, 257, 259, 261, 270, 271, 281, 288, 290, 291, 292, 311, 321, 377 Cryptodrilus 282, 283, 352, 365 Ctenodrilida 501 Ctenodrilidae 12, 497, 500, 501, 502 Ctenodrilinae 11, 156, 498, 500, 501, 502, 512 Ctenodrilus 49, 65, 498, 501, 502, 503, 507, 508, 511, 512, 513, 514, 515, 516 Cuticle 17, 271, 306, 321, 406, 576, 587, 588, 592, 593, 595, 612, 616, 644, 646 Cyclin 413, 416 Cylicobdella 398 Cylicobdellidae 395, 403 Cysts 62, 63, 331, 333, 356, 357, 359, 360 Cytophore 27, 37, 53, 54, 58, 60, 62, 306, 328, 333, 335, 336, 351, 356, 357, 359, 360, 403, 404, 405, 509, 532, 533, 649, 651, 653, 655, 656

D D-quadrant 95, 96, 97, 100, 101, 102, 104, 106, 107, 109, 112, 117, 369, 371, 410, 417, 418 Deformation Movement 367, 369 Delaya 253

Index Demonax 50, 539 Dendrobaena 329, 351, 369 Dentatisyllis 456 Dero 248, 249, 250, 377 Desdemona 524, 526 Desmogaster 304, 305 Deuterostomia 93 Development 4, 6, 23, 25, 28, 39, 45, 53, 58, 60, 62, 63, 93, 94, 95, 96, 102, 104, 105, 106, 108, 109, 110, 114, 116, 117, 118, 119, 120, 121, 122, 123, 125, 126, 128, 130, 131, 132, 141, 142, 143, 146, 149, 151, 154, 156, 158, 160, 166, 179, 180, 181, 182, 183, 184, 186, 187, 188, 189, 190, 191, 192, 193, 194, 197, 210, 218, 240, 252, 265, 269, 270, 273, 282, 284, 293, 294, 297, 313, 328, 331, 336, 359, 366, 367, 368, 369, 371, 373, 374, 376, 377, 378, 396, 406, 408, 409, 410, 412, 413, 415, 418, 419, 420, 422, 423, 436, 437, 438, 440, 442, 444, 445, 446, 448, 450, 452, 454, 456, 458, 460, 461, 462, 463, 469, 471, 472, 473, 474, 475, 480, 482, 497, 498, 504, 505, 508, 509, 510, 511, 513, 514, 516, 528, 530, 533, 535, 537, 538, 540, 542, 544, 546, 548, 549, 565, 566, 569, 571, 574, 576, 577, 578, 582, 583, 584, 586, 587, 588, 589, 594, 595, 597, 598, 599, 600, 601, 602, 604, 605, 606, 607, 608, 610, 611, 612, 613, 614, 616, 617, 619, 621, 622, 623, 624, 625, 626, 627, 628, 631, 658, 659, 660 Diaphorodrilus 327 Dichogaster 263, 265, 292, 326, 327 Dichogastrini 264, 265 Dichotomous Gland 284, 307, 309 Didymogaster 263, 265, 281 Digaster 240, 263, 265, 282, 283, 289, 352, 365 Dimorphism 85, 292, 504, 505, 512 Dinophilidae 58, 340, 639, 640, 642, 651, 656, 658 Dinophilus 42, 46, 58, 85, 642, 651, 656, 658 Diopatra 31, 34, 47 Diplocardia 254 Diplotesticulata 244, 252, 253, 293, 302, 351 Diplotrema 263, 273, 282, 326 Dipolydora 571, 584, 585, 586, 614, 617, 620, 621, 622, 626, 628, 629 Direct 23, 25, 32, 37, 94, 95, 109, 117, 121, 151, 180, 182, 187, 191, 192, 209, 215, 218, 246, 271, 288, 290, 369, 420, 421, 468, 471, 498, 507, 508, 511, 512, 575, 576, 586, 598, 619, 623, 624, 625, 656, 657, 658, 660 Disomidae 171, 567 Dispio 569, 571, 575, 576, 582, 587, 588, 589 Ditrupa 541 Diurodrilidae 58, 639, 643, 656 Diurodrilus 46, 57, 58, 59, 640, 643, 651 DNA 5, 123, 189, 251, 259, 260, 261, 304, 356, 357, 359, 360, 393, 396

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Dodecaceria 498, 500, 501, 506, 509, 510, 511, 514 Dorsal 112, 113, 114, 135 Dorsal Organs 616, 657 Dorvillea 163 Dorvilleidae 9, 13, 46, 58, 87, 143, 144, 153, 154, 158, 163, 640 Dorydrilidae 245, 252, 260, 296 Dorydrilus 252, 296 Doublet 339, 350 Downstream 151, 153, 160, 161, 162, 163, 165, 166, 186 Downstream Larval Feeding 153, 160, 163 Drawida 284, 293 Drilocrius 256, 257, 319, 320, 321 Drosophila 93, 94, 98, 112, 115, 119, 120, 121, 122, 124, 128, 131, 132, 419, 421, 422 Dwarf Males 56, 526, 528, 642, 650, 656, 658 Dysponetus 47, 438, 467

E Echiura 4, 5, 6, 7, 13, 52, 53, 56, 64, 65, 144, 145, 151, 153, 157, 160, 163, 501, 502 Ect-aquasperm 55, 56, 438, 439, 441, 443, 444, 445, 446, 447, 449, 453, 454, 455, 457, 458, 459, 461, 462, 463, 464, 466, 467, 471, 532, 571, 578, 579, 580, 651, 653, 656 Ectoderm 99, 107, 109, 110, 111, 112, 113, 115, 117, 118, 121, 122, 123, 125, 129, 367, 369, 371, 373, 376, 377, 408, 410, 415, 418 Ectoteloblast 112, 369, 374 Edysozoa 93 Egg 14, 23, 33, 36, 38, 39, 55, 78, 79, 80, 82, 85, 88, 95, 96, 97, 106, 149, 183, 187, 189, 190, 191, 192, 288, 307, 316, 319, 324, 325, 327, 328, 329, 331, 350, 367, 368, 369, 371, 406, 408, 437, 444, 445, 446, 450, 455, 460, 461, 464, 475, 480, 508, 511, 512, 513, 529, 530, 536, 537, 538, 540, 542, 544, 546, 547, 566, 568, 571, 574, 575, 576, 577, 578, 582, 584, 585, 586, 587, 588, 589, 592, 593, 594, 595, 597, 598, 606, 607, 608, 612, 613, 618, 621, 622, 623, 624, 626, 627, 657, 658, 660 — cortex 95, 367 — envelope 39, 464, 530, 571, 575, 576, 577, 578, 587, 588, 592, 593, 594, 595 — exchange 78, 80 — mass 445, 446, 450, 455, 460, 511, 513, 582, 586 — size 85, 192, 480, 512, 529, 530 — trading 78, 80 Eisenia 5, 27, 29, 37, 117, 246, 248, 249, 250, 269, 321, 328, 329, 331, 351, 369, 373, 377, 410 Eiseniella 351, 361 Emboly 368, 369 Embryogenesis 93, 117, 129, 240, 367, 406, 408, 419, 420, 422 Embryology 6, 95, 108, 109, 110, 367, 368, 441

$%" Reproductive Biology and Phylogeny of Annelida Embryos 87, 88, 93, 96, 99, 101, 102, 103, 104, 106, 108, 110, 111, 115, 118, 119, 120, 161, 184, 186, 191, 269, 271, 329, 368, 369, 374, 376, 377, 408, 409, 415, 419, 421, 422, 448, 457, 478, 479, 511, 523, 531, 536, 575, 592, 607, 608, 612, 613, 622, 623, 626, 627 Enantiodrilus 254, 303, 312 Enchytraeidae 237, 240, 244, 245, 246, 247, 248, 250, 260, 261, 266, 277, 288, 296, 297, 298, 328, 336, 340, 343, 361, 368, 373 Enchytraeus 27, 37, 160, 167, 239, 248, 266, 297, 328, 329, 331, 341, 361, 368 Encounter Predators 153, 155, 156, 157, 166, 474 Endocytosis 32, 37, 328, 529 Endodermal 111, 369, 371, 374, 408, 410, 421, 525 Endolarva 158, 659 Endomyzostoma 474 Endonuclear Canal 335, 340, 350 Engrailed (En) 95, 118, 120, 121, 122, 413, 416, 419, 422 Enhancer of Split 413, 422 Enipo 453, 461 Ent-aquasperm 55, 56, 448, 449, 450, 451, 453, 455, 456, 458, 460, 467, 533, 578, 579 Environmental Stimuli 507 Eoclitellata 260, 261 Epiboly 110, 111, 368 Epidermis 63, 113, 118, 191, 235, 240, 268, 271, 284, 295, 305, 306, 321, 322, 367, 373, 375, 412, 421, 535, 643, 646, 648, 649, 656, 657 Epididymides 316 Epigamia 456, 480 Epigamous 433, 465, 469, 471 Epirodrilus 293 Episphere 142, 143, 152, 158, 549, 659 Epithelium 27, 111, 112, 113, 269, 281, 285, 289, 290, 292, 307, 309, 311, 329, 355, 371, 373, 376, 377, 378, 408, 409, 410, 412, 413, 415, 417, 418, 420, 507, 526, 574, 644 Epitokes 88, 433, 435, 504, 510, 511, 512 Epitoky 88, 433, 437, 438, 440, 442, 444, 446, 448, 450, 452, 454, 456, 458, 460, 464, 468, 469, 471, 507, 510, 511, 512, 623 Epitrachys 393 Erpobdella 248, 249, 399 Erpobdellidae 395, 401, 403 Erpobdelliformes 292, 395, 396, 401, 403, 404, 406 Erpochaete 149 Errantia 7, 431 Esophageal Diverticula 325 Eteone 449, 480 Eteoninae 432 Euchone 50, 524, 526, 539 Euclitellata 235 Euclymene 86 Eudiplotrema 327

Eudrilidae 240, 244, 256, 258, 259, 260, 261, 263, 268, 270, 273, 277, 282, 290, 312, 315, 335, 343, 349, 350 Eudrilus 256, 258, 259, 277, 331, 333, 335, 336, 337, 338, 340, 349, 350 Euhirudinea 236, 260, 300 Eukerria 263, 276, 325, 326, 361, 363, 365 Eulalia 48, 449, 461, 466, 467 Eulepethidae 13, 155 Eumida 449, 466, 480 Eunice 148, 163 Eunicid Polychaetes 235 Eunicida 9, 12, 13, 17, 39, 46, 47, 66, 147, 154, 155, 431 Eunicidae 9, 13, 39, 47, 143, 144, 148, 151, 153, 154, 158, 163 Euphrosinidae 9, 13, 154 Eupolygaster 293 Euprostate 282 Eurasian 399 Euratella 26 European 398, 399, 508, 510, 528, 598, 605 Eurythoe 46, 55, 65, 163 Euspermatozoa 292, 340, 345, 350, 353, 354, 355, 356, 357, 359, 361, 653, 657 Eusyllinae 471 Eusyllis 456 Eutyphoeus 362, 363 Eve 120, 121, 413, 416, 419, 420 Even-skipped 413, 416, 422 Eversible Pharynx 153 Exogone 48, 86, 456, 479 Exogoninae 82, 471 Exolarva 152, 158, 659 External Fertilization 53, 56, 63, 64, 78, 87, 179, 193, 467, 508, 509, 510, 532, 533, 550, 551, 657 Extinction 399 Extraovarian 25, 27, 32, 38, 327, 437, 443, 445, 446, 449, 456, 459, 460, 461, 529, 573, 574, 649, 650 Extraovarian Oogenesis 25, 38, 461, 573, 574 Eyes 99, 102, 103, 104, 106, 160, 416, 420, 433, 468, 511, 512, 516, 549, 567, 587, 588, 589, 590, 592, 593, 595, 596, 597, 599, 607, 609, 611, 612, 614, 615, 616, 618, 619, 622, 623 Eyespots 152, 395, 396, 510, 511, 512, 514, 516, 549

F Fabricia 34, 50, 524 Fabriciinae 50, 524, 526, 528, 532, 535, 539 Fabricinuda 51 Fabriciola 51 Fabrisabella 524 Facultative Change of Gender 86 Fate Maps 99, 109, 119, 367

Index Fauveliopsidae 11, 13, 156, 497, 498, 500, 501, 502, 510 Fauveliopsis 498, 501, 504, 506, 510 Feeding Appendages 660 Female 14, 15, 25, 27, 32, 38, 55, 56, 60, 62, 77, 78, 81, 82, 83, 84, 85, 86, 87, 88, 160, 190, 235, 268, 270, 271, 274, 277, 280, 288, 295, 296, 298, 303 309, 312, 315, 316, 318, 319, 322, 325, 327, 350, 399, 400, 401, 402, 403, 437, 462, 467, 468, 471, 472, 475, 477, 478, 479, 480, 506, 510, 514, 526, 529, 535, 537, 538, 540, 542, 544, 546, 548, 573, 574, 575, 579, 580, 584, 585, 586, 605, 606, 625, 627, 643, 644, 645, 646, 647, 648, 649, 650, 656, 657, 659, 660 — ducts 268, 295, 643, 648 — function 78, 86 — organs 471, 643, 645, 647, 648, 649 — pores 235, 274, 277, 296, 298, 309, 312, 315, 318, 319, 322, 325, 327, 526, 660 Fertilization 23, 24, 25, 53, 55, 56, 57, 60, 62, 63, 64, 65, 66, 78, 80, 81, 82, 86, 87, 95, 96, 179, 180, 184, 189,193, 265, 268, 277, 284, 290, 296, 313, 315, 329, 339, 343, 350, 352, 353, 356, 361, 366, 398, 401, 406, 408, 461, 462, 464, 467, 468, 469, 508, 509, 510, 511, 528, 529, 531, 532, 533, 535, 537, 548, 550, 551, 566, 576, 578, 581, 582, 586, 595, 601, 627, 656, 657, 658, 659 — cone 366 Ficopomatus 526, 537, 541, 548 Filograna 526, 527, 529, 534, 536, 540, 550 Filograninae 51, 525, 540, 541 Filter House 166 Fitness 78, 82, 84, 87, 180, 183 Flabelliderma 165, 499, 505, 510, 516 Flabelligella 500, 504 Flabelligera 499, 501, 502, 505, 509, 515, 516 Flabelligeridae 8, 11, 13, 49, 52, 153, 156, 165, 497, 499, 500, 501, 502, 505 Flagellum 53, 58, 60, 63, 266, 333, 335, 338, 339, 340, 346, 348, 351, 352, 354, 355, 360, 404, 405, 463, 466, 508, 509, 515, 533, 579, 651, 653, 655 Flame Cell 377 Fletcherodrilus 14, 266, 281, 345, 352 Floriprotis 532, 536, 541 Flota 11, 497, 502 Follicle Cell 27, 36, 37 Foregut 17, 111, 113, 114, 115, 121, 127, 128, 131, 409, 412, 416, 418, 422, 660 Fossil 12, 13, 393 Fragmentation 358, 359, 360, 510, 628, 630 Freshwater 123, 235, 237, 239, 251, 256, 296, 343, 353, 371, 373, 394, 396, 398, 400 Fridericia 248, 249, 297 Frogs 396 Frontal Organ 143

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Functions of the Cytophore 333 Funnel 294, 297, 307, 315, 322, 355, 378, 435, 436, 536, 643, 644

G Galathealinum 530 Galeolaria 51, 66, 165, 527, 531, 532, 535, 537, 541, 548 Gametogenesis 403, 433, 437, 480, 497, 511, 566, 586 Gastrolepidia 84 Gastrotroch 142, 146, 148, 151, 596, 609, 615 Gastrula 110, 115, 141, 369, 377 Gastrulation 108, 109, 110, 111, 112, 113, 114, 115, 116, 369, 371, 373, 376, 377 Gattyana 453 Gemmiparity 433, 465, 471 Gene Expression 94, 98, 108, 122, 124, 128, 129, 130, 131, 413, 419, 420 Genital Chaetae 85, 271, 273, 281, 311, 316, 319, 321, 363, 365, 647 Genital Papillae 312, 315, 471, 504, 505 Genital Pores 271, 291, 362, 504 Geofdyneia 285, 286 Germinal Bands 110, 111, 112, 113, 118, 130, 409, 412, 413, 415, 417, 419 Germinal Epithelium 27, 507, 526 Germinal Plate 111, 113, 118, 128, 131, 409, 412, 415, 417, 419, 420 Gland Cells 300, 307, 309, 311, 643, 647, 648, 656 Glossiphonia 29, 110, 399 Glossiphoniidae 394, 396, 398, 400, 401, 403, 406 Glossoscolecidae 244, 254, 256, 257, 258, 259, 260, 261, 263, 270, 273, 275, 279, 285, 291, 303, 312, 313, 320, 321, 324, 366 Glossoscolecini 258 Glossoscolecoidea 259, 260 Glossoscolex 281, 285, 312 Glycera 163, 438, 439, 468, 472 Glyceridae 13, 24, 25, 143, 147, 153, 155, 163, 432, 433, 435, 436, 468, 472 Glyceriformia 432, 438, 439, 440 Glycinde 439 Glycogen 29, 289, 335, 338, 339, 340, 348, 350, 351, 352, 356, 435 Glycymeris 248, 250 Glyphidrilocrius 257, 321 Glyphidrilus 254, 256, 271, 319, 320, 321, 363 Goblet Cells 305, 306 Golgi 29, 32, 33, 34, 36, 37, 282, 288, 333, 335, 336, 337, 404, 653 Gonad 23, 116, 300, 378, 507, 510, 526 Gondwanan 398, 399 Goniada 439

$%$ Reproductive Biology and Phylogeny of Annelida Goniadella 439 Goniadidae 13, 143, 153, 155, 436 Gonochorism 24, 77, 80, 83, 84, 85, 86, 88, 467, 468, 469, 471, 504, 513, 528, 529, 550, 642, 643, 645, 657 Gonoduct 301, 378, 505 Gonopore 398, 399, 403, 505, 526 Goosecoid 113, 114, 115 Gordiodrilus 285 Grooved Palps 8, 9, 10, 11, 12, 497, 501, 503 Ground Water 639 Growth Hormone 507 Growth Zone 4, 109, 117, 120, 125, 127, 132, 369, 631 Grubeosyllis 48, 82, 87, 460 Guamerins 398 Gyptis 432, 440, 460, 462

H Haemadipsidae 395, 396, 402 Haementeria 394, 396, 398, 400, 408 Haemopidae 395, 396, 402 Haemopis 396, 398, 399, 401 Hairy 121, 413, 416, 422 Halosydna 453 Hamingia 52, 56 Haploscoloplos 46 Haplosyllis 456 Haplotaxida 242, 243, 253, 265, 302 Haplotaxidae 242, 244, 245, 246, 252, 253, 254, 260, 265, 269, 273, 303, 305, 348, 351 Haplotaxis 238, 251, 253, 254, 265, 303, 331, 333, 336, 337, 338, 345, 348, 351 Harmothoe 32, 48, 85, 149, 163, 340, 436, 453, 454, 461, 467, 475 Hartmaniellidae 9, 13, 154 Hastirogaster 293 Hb 98, 107, 112, 113, 120, 121, 417 Hedgehog 113, 116, 121, 132, 413, 416, 422 Hediste 82, 85, 191, 437, 443, 464, 469, 474 Helix 113, 248, 250, 404, 413 — loop-helix 113, 413 Helobdella 78, 82, 84, 88, 94, 95, 98, 101, 111, 115, 116, 122, 126, 128, 129, 180, 398, 400, 402, 408, 409, 410, 411, 413, 415, 416, 417, 418, 420 Hemiclepsis 248, 249 Hemipodia 440, 472 Hemisegmental 412, 415 Hemocoel 27 Hermaphrodite 82, 86, 288, 361, 466, 514, 515, 536, 648 Hermaphroditic 24, 27, 77, 78, 79, 80, 81, 82, 83, 84, 86, 87, 268, 328, 466, 467, 469, 504, 511, 642, 643, 645, 647, 657, 658 Hermione 438

Heronidrilus 251 Hes 120, 121, 413, 416 Hesione 468 Hesionidae 13, 47, 56, 143, 147, 153, 155, 163, 431, 432, 435, 440, 441, 468, 469, 472 Hesionides 47, 431, 446, 466, 469 Hesiospina 462 Heterochaeta 294, 353, 360, 512 Heterochaetella 253 Heterodrilus 251 Heteronereis 435, 470 Heteronomicity 127 Heteropodarke 432 Heteroporodrilus 263, 274, 280, 282, 283, 289, 365 Heterospio 11, 157, 500, 567, 569, 571, 572, 575, 582 Heterosynthesis 32, 328, 529, 573 Hirudinea 8, 81, 236, 237, 248, 249, 250, 260 Hirudinida 3, 6, 7, 45, 179, 237, 260, 261, 393, 394, 408 Hirudinidae 395, 396, 401, 402 Hirudo 94, 125, 128, 129, 302, 394, 395, 400, 401, 407, 408, 416, 417, 418, 420 Histriobdella 47 Histriobdellidae 9, 13, 47, 142, 143, 147, 153, 154 Hologynus 253, 302 Holonephridia 263 Homeobox 98, 107, 116, 123, 124, 125 Homeodomain 122, 123, 413 Homology 4, 5, 38, 64, 122, 147, 153, 163, 328, 475, 568 Homoplasy 311, 346, 361 Hoplochaetella 273, 326 Hormogaster 259, 270, 323, 345, 351, 366 Hormogastridae 244, 256, 258, 259, 261, 270, 273, 316, 323, 351 Hox 95, 123, 124, 125, 126, 127, 128, 129, 130, 131, 132, 413, 416, 417, 420, 421, 422 — genes 123, 124, 125, 126, 127, 128, 130, 131, 132, 420, 421, 422 Hrabeiella 12, 52, 57, 66, 157, 639, 641, 648, 649, 651, 652, 655, 656, 658, 660 Hr-nos 107 Hro-dl 113, 114, 416, 419, 421 Hro-sna 113, 114, 418 Hro-twi 98, 113, 114, 413, 418 Hr-wnta 107 Htr-lox22 113, 114, 115 Hunchback 98, 107, 112, 113, 121, 132, 413, 417, 422 Hyalinoecia 47, 66 Hydroides 51, 104, 106, 120, 183, 184, 185, 188, 190, 193, 194, 196, 198, 199, 200, 201, 202, 203, 205, 207, 210, 211, 212, 213, 214, 215, 216, 217, 218, 219, 220, 221, 222, 526, 529, 530, 532, 535, 537, 541, 542, 548 Hydrothermal Vents 525, 548

Index Hypodermic Impregnation 25, 81, 85, 290 Hypodermic Injection 466, 469, 656 Hypoplectrus 78 Hyposphere 143, 152, 158, 549 Hystricosoma 642

I Ichthyotomus 13, 432 Idanthyrsus 49, 523, 538 Inanidrilus 251, 273 Inbreeding 85 Incertae Sedis 8, 9, 10, 13, 52, 57, 156, 157, 253, 447, 639 Indirect Development 95, 514 Insemination 80, 81, 271, 290, 361, 362, 398 In-situ Hybridization 94, 125, 126, 128 Insulodrilus 251, 343 Intercellular Bridges 25, 27, 28, 37, 328, 329, 436, 437, 461 Internal Fertilization 57, 63, 64, 66, 80, 86, 277, 296, 313, 315, 343, 350, 398, 461, 510, 529, 535, 548, 656, 657, 658 Interpolation of the Mitochondria 57, 339 Interstitial 25, 85, 87, 464, 466, 469, 639 Intracellular Tracers 118 Intraovarian 25, 27, 32, 35, 38, 327, 436, 437, 441, 442, 453, 530, 573, 649, 650, 651 — oogenesis 25, 27, 32, 38, 437, 573 Introsperm 55, 56, 339, 440, 441, 442, 446, 447, 448, 452, 455, 457, 458, 464, 468, 508, 571, 579, 580, 651, 652 Iospilidae 13 Isochaetides 353, 355 Iteroparous 504, 529

J Janua 82, 210, 217, 527, 534, 537, 544 Janus Monsters 96 Jasmineira 50, 524, 526 Jelly Masses 504, 511 Josephella 525, 550 Juvenile 17, 86, 95, 116, 149, 152, 158, 160, 180, 181, 183, 190, 193, 194, 195, 196, 202, 409, 438, 448, 452, 477, 514, 548, 571, 588, 599, 600, 602, 603, 606, 610, 611, 616, 617, 627, 640, 656

K Kayarmacia 326 Kefersteinia 32, 437 Kinbergonuphis 47 Kinkaidiana 346, 348 Komarekiona 255, 256, 257, 269, 261, 322, 323

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Komarekionidae 244, 255, 256, 258, 259, 260, 261, 322 Krenedrilus 272 Krohnia 48, 449, 466 Kynotidae 244, 256, 257, 261, 273, 281, 316, 318

L Lacydonia 13, 432 Laeonereis 443 Laetmonice 47 Lagisca 454 Lamellibrachia 35 Lamprodrilus 249 Lanice 5, 104, 248 Laonice 566, 567, 568, 571, 575, 579, 582, 589, 590, 591, 598 Laonicinae 566, 567, 568, 569 Laonome 532, 539 Larvae 60, 88, 102, 103, 105, 106, 114, 115, 119, 120, 123, 126, 127, 141, 143, 144, 145, 146, 147, 148, 149, 151, 152, 153, 154, 155, 158, 160, 161, 163, 165, 166, 179, 180, 181, 182, 183, 186, 187, 188, 189, 190, 191, 192, 193, 195, 196, 197, 198, 199, 200, 201, 202, 203, 204, 206, 209, 210, 215, 216, 217, 218, 219, 220, 221, 222, 438, 440, 449, 452, 461, 472, 474, 475, 480, 482, 511, 512, 513, 514, 515, 516, 523, 528, 530, 531, 533, 534, 535, 536, 537, 547, 548, 549, 550, 551, 565, 567, 575, 582, 585, 586, 587, 589, 590, 591, 592, 593, 594, 595, 596, 597, 598, 599, 600, 601, 602, 603, 604, 605, 606, 607, 610, 612, 613, 614, 615, 617, 618, 619, 620, 621, 622, 623, 624, 625, 626, 627, 630, 632, 658, 659, 660 — chaetae 152, 158, 182, 195, 196, 472, 474, 516, 589, 593, 594, 602, 604, 606, 609, 615, 616, 622 — characters 8, 569, 571, 640 — leeding 149, 151, 153, 154, 158, 160, 163, 165, 166 Lateral Organs 646, 647, 657 Laubieriopsis 498, 501, 504, 508, 510, 513 Laubierpholoe 48, 434, 447, 461, 475, 477 Laurasian 398, 399 Lecithotrophy 95, 145, 149, 151, 153, 154, 155, 158, 160, 165, 166, 182, 186, 187, 188, 189, 190, 191, 192, 193, 328, 376, 438, 439, 440, 441, 444, 446, 447, 448, 455, 457, 459, 460, 474, 475, 480, 498, 511, 514, 528, 530, 536, 537, 539, 540, 541, 542, 543, 544, 545, 546, 547, 549, 575, 586, 587, 588, 604, 605, 606, 607, 608, 626, 627 Leech 3, 7, 8, 23, 25, 27, 29, 37, 39, 78, 81, 82, 84, 87, 88, 93, 94, 95, 96, 98, 99, 101, 102, 107, 108, 110, 111, 112, 114, 115, 116, 118,

$%& Reproductive Biology and Phylogeny of Annelida 119, 120, 121, 123, 124, 125, 126, 128, 129, 130, 131, 132, 151, 179, 180, 235, 236, 237, 240, 244, 250, 251, 252, 253, 293, 301, 302, 337, 338, 339, 346, 350, 393, 394, 395, 396, 397, 398, 399, 400, 401, 402, 403, 404, 405, 406, 407, 408, 409, 410, 411, 412, 413, 415, 416, 418, 419, 420, 421, 422 Leitoscoloplos 32, 148, 161 Le-msx 98, 107, 113, 413, 417, 419, 420, 421 Lepidonotus 48, 66, 434, 454, 461 Leptonereis 444 Limen 265, 340, 343, 345, 346, 348, 351, 352 Limited Circulatory System 433 Limnic 639 Limnodriloides 251, 265, 272, 333, 336, 343, 353, 355 Limnodrilus 251, 273 Lindaspio 571, 575, 582 Linta 396, 398, 399 Lizardia 47, 440, 462, 468 Local Mate Competition 84, 85, 86 Long-headed Sperm 508, 566 Longosomidae 567 Lopadorhynchidae 13, 142, 143, 147, 155, 432, 442 Lopadorhynchus 442 Lophotrochozoa 93, 99, 122, 132 Lox 113 Lox1 125, 128, 130 Lox2 125, 128, 130, 131, 417, 421 Lox3 116, 418, 421, 422 Lox4 125, 128, 130, 417, 421 Lox5 125, 128, 417, 420, 421 Lox6 125, 126, 128, 130, 132, 416, 420 Lox7 125, 128, 416, 420 Lox10 116, 417, 420, 421 Lox18 125, 130, 416, 420 Lox20 125, 126, 128, 130, 417, 420, 421 Lumbricidae 52, 80, 244, 245, 246, 248, 250, 256, 257, 258, 259, 260, 263, 265, 270, 273, 274, 281, 290, 315, 317, 318, 329, 331, 351, 352, 361, 362, 363, 369 Lumbricillus 239, 265, 266, 288, 340, 342 Lumbricoidea 255, 257, 258, 259, 260, 319 Lumbriculata 244, 252, 253, 260, 293, 300 Lumbriculida 6, 236, 242, 252, 266, 394 Lumbriculidae 236, 237, 242, 244, 245, 248, 249, 250, 251, 252, 253, 260, 265, 266, 267, 273, 277, 293, 296, 300, 301, 304, 336, 346, 347, 350, 358, 368, 410 Lumbriculus 247, 248, 249, 250 Lumbricus 78, 80, 81, 110, 246, 248, 249, 315, 317, 318, 319, 329, 331, 333, 336, 345, 351, 352, 362, 363, 366 Lumbrineridae 9, 13, 47, 143, 144, 153, 154, 163 Lumbrineris 47, 163 Lutodrilidae 244, 255, 256, 257, 259, 261, 319, 320

Lutodrilus 255, 259, 261, 292, 293, 319, 320, 321, 363 Lycodrilus 252, 296 Lygdaminae 522 Lygdamis 538 Lzf2 98, 112, 113, 121, 413, 417, 419, 420, 421, 422

M Macellicephala 467 Macrobdella 398, 401 Macrochaeta 48, 498, 500, 504, 506, 508, 509, 515 Macromeres 99, 100, 101, 104, 108, 109, 121, 367, 369, 408, 409, 410, 412, 416, 417 Magelona 49, 165, 575, 583, 602, 604, 605 Magelonidae 11, 13, 49, 147, 153, 157, 165, 166, 565, 568, 569 Malabarinae 325 Malacoceros 15, 566, 567, 568, 571, 575, 579, 582, 586 Malagasy 395, 398, 399 Maldanidae 8, 12, 13, 24, 46, 148, 153, 154, 160, 161 Male 55, 59, 77, 78, 80, 81, 82, 83, 84, 85, 86, 87, 88, 191, 240, 242, 245, 252, 254, 257, 258, 259, 262, 265, 268, 270, 271, 272, 273, 274, 275, 276, 281, 282, 284, 285, 288, 293, 294, 295, 296, 297, 298, 299, 300, 301, 302, 303, 304, 305, 307, 309, 311, 312, 313, 315, 316, 318, 319, 321, 322, 323, 324, 325, 326, 327, 337, 361, 362, 363, 365, 399, 400, 401, 402, 403, 404, 461, 462, 467, 468, 469, 471, 472, 479, 505, 506, 509, 510, 511, 514, 515, 526, 529, 535, 578, 581, 583, 584, 642, 643, 644, 645, 646, 647, 648, 649, 650, 658, 659 — combat 83, 85 — competition 83, 84, 85 — function 78, 80, 82, 86 — organs 469, 471, 642, 643, 647, 648 — pores 242, 252, 254, 257, 258, 262, 265, 268, 270, 271, 272, 273, 274, 276, 293, 294, 296, 297, 298, 299, 302, 305, 309, 311, 312, 315, 316, 318, 319, 321, 322, 323, 324, 325, 326, 327, 361, 365, 505, 526 — terminalia 282, 326 Manayunkia 51, 54, 55, 524 Manchette 63, 333, 335, 337, 357, 463 Mantle Cell 377 Map Kinase 106, 107, 133 Marenzelleria 49, 571, 573, 575, 576, 577, 578, 579, 580, 582, 586 Marifugia 542 Marphysa 47, 151 Marsupiobdella 396, 398 Mastotermes 360 Mate Choice 83 Maternal Determinants 95, 96

Index Mating 77, 78, 80, 81, 82, 83, 84, 85, 86, 87, 88, 361, 363, 365, 366, 403, 406, 438, 440, 442, 444, 446, 448, 450, 452, 454, 456, 458, 460, 465, 467, 468, 469, 471, 510, 533, 581, 656, 658 — conflict 78 — group Size 82 — system 77, 80, 83, 85, 87, 88 Mediomastus 161, 166 Megadrile 244, 246, 254, 273, 319, 351 Megalomma 539 Meganerilla 643 Megascolecidae 14, 238, 244, 246, 248, 254, 256, 258, 261, 262, 263, 265, 266, 270, 273, 274, 275, 280, 281, 282, 283, 285, 286, 288, 289, 304, 307, 312, 323, 325, 326, 352, 362, 363, 365 Megascolecin Condition 327 Megascolecinae 237, 246, 248, 249, 261, 263, 264, 265, 273, 277, 282, 283, 286, 289, 325, 327, 362, 365 Megascolecini 264, 265 Megascolecoidea 255, 258, 261, 325 Meiofauna 78, 82, 639 Meiosis 29, 96, 189, 357, 360, 366, 367, 369, 403, 408, 548 Meiotic Apparatus 367 Meiotic Division 327, 333, 335, 650 Melinna 165 Meniscotroch 142, 143, 145, 149, 155, 163, 166, 472, 474, 475, 480 Mercierella 32 Meronephridia 263, 265 Mesenchytraeus 297, 298, 340 Mesobdella 399, 402, 403 Mesoblasts 371, 373 Mesochaetopterus 49, 602, 603 Mesoderm 99, 107, 109, 111, 112, 113, 114, 115, 116, 117, 121, 122, 123, 127, 128, 131, 132, 367, 371, 375, 376, 408, 410, 412, 415, 421, 422 Mesodermal Segmentation 375 Mesolecithal 368 Mesonerilla 650, 658, 659 Mesoteloblasts 109, 111, 373, 374, 410 Mesotroch 142, 146, 148, 151, 601, 602 Metabonellia 151 Metagynophora 243, 244, 245, 253, 254, 260, 277, 281, 293, 298, 303, 304 Metalaeospira 51, 536, 545, 546 Metamerism 4 Metamorphosis 95, 117, 125, 127, 141, 145, 149, 152, 158, 160, 179, 180, 182, 187, 190, 193, 195, 196, 197, 198, 199, 202, 203, 204, 205, 210, 215, 216, 217, 218, 221, 222, 368, 482, 511, 522, 548, 588, 595, 596, 597, 598, 599, 601, 602, 604, 605, 606, 608, 616, 617, 619, 659 Metanephridia 17, 268, 377, 435, 436, 468

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Metanephromixia 435, 436 Metaphase 27, 327, 335, 367, 369, 408 Metaphire 288, 331, 362, 365 Metaprostates 282, 312, 326 Metataxis 253 Metatroch 113, 142, 143, 144, 145, 146, 147, 148, 149, 151, 153, 158, 163, 165, 166, 186, 480, 516, 549 Metatrochophore 127, 147, 151, 438, 472, 474, 475, 480, 481, 549, 660 Metavermilia 536, 542 Metazoa 5, 32, 38, 39, 160 Methanoaricia 26, 46 Michalakus 312, 315, 316 Microchaetidae 244, 256, 257, 258, 259, 260, 270, 273, 281, 315, 316, 350 Microchaetus 255, 259, 281, 293, 311, 350, 351 Microdriles 240, 269, 270, 273, 274, 277, 281, 298, 327, 329, 333, 368 Microfilaments 96, 367 Micromaldane 25, 46, 340 Micromeres 99, 101, 104, 107, 109, 113, 114, 367, 369, 373, 374, 408, 409, 410, 411, 412, 413, 415, 416, 417, 418, 419 Micronereis 444, 469 Microphthalmus 47, 431, 447, 466, 469, 647 Micropodarke 462 Microprotula 536, 540 Microscolecin Condition 326 Microscolex 281, 285, 288, 326, 362, 365 Microspio 566, 569, 571, 575, 579, 583 Microtubuli 465, 466 Midgut 109, 110, 111, 113, 115, 116, 121, 127, 128, 131, 367, 369, 373, 376, 377, 409, 410, 416, 417, 418, 422 Midpiece 53, 57, 58, 59, 60, 245, 265, 266, 267, 333, 335, 337, 338, 340, 342, 343, 345, 346, 348, 350, 351, 352, 354, 355, 464, 465, 509, 532, 533, 579, 651, 653, 655, 656 Mitochondria 29, 31, 32, 33, 34, 37, 53, 55, 57, 58, 59, 60, 62, 63, 261, 265, 267, 288, 328, 333, 335, 337, 338, 339, 340, 343, 345, 346, 348, 351, 352, 354, 355, 356, 360, 367, 369, 371, 404, 408, 435, 461, 462, 463, 464, 465, 466, 508, 509, 532, 533, 550, 579, 580, 651, 653, 655, 656 Mitochondrial Cytochrome Oxidase Subunit I 246 Mitraria 149, 152, 158, 186, 528, 548 Mixonephridia 435, 436 Modified Sperm 53, 56, 57 Molecular Analysis 236, 244, 250, 253, 254, 255, 256, 259, 261, 304, 325, 431, 502, 642 Molecular Evidence 3, 6, 122, 236, 251, 261 Molecular Studies 5, 7, 108, 120, 236, 244, 246, 258, 296, 296, 413, 422, 641, 642 Mollusca 5, 6, 64, 143, 147, 161, 248, 249, 360, 365

$& Reproductive Biology and Phylogeny of Annelida Moniligaster 284, 285, 293, 304, 305, 306, 307, 309 Moniligastrida 242, 243, 254, 304 Moniligastridae 240, 242, 245, 254, 260, 269, 273, 281, 284, 304, 305, 306, 307, 309 Monophyly 3, 4, 5, 6, 7, 8, 9, 10, 57, 65, 236, 237, 240, 243, 244, 250, 251, 253, 254, 255, 256, 258, 259, 263, 265, 266, 267, 304, 340, 394, 396, 432, 433, 497, 501, 502, 524, 525, 639, 640, 641, 642 Monopylephorus 273, 294, 339, 343 Morphocladistic Analysis 240, 241, 242, 243, 244, 245, 246, 251, 252, 253, 254, 255, 256, 257, 258, 261 Morphogenesis 107, 116, 121, 122, 141, 181, 335, 336, 337, 357, 374, 375, 409, 410, 422, 627, 628 Morulae 53, 60, 306, 307, 331, 333, 463, 509, 533, 651 Markov Chain Monte Carlo (MCMC) 246, 247 Msx 98, 107, 113, 413, 417, 419, 420, 421 Mucous 26, 27, 78, 80, 86, 87, 153, 157, 206, 209, 288, 305, 449, 450, 471, 480, 582, 584, 586, 595, 604 — bag 449, 450, 471 Multivesicular Bodies 328 Myrianida 456, 457, 465, 470, 471 Mytilus 249, 250 Myxicola 50, 524, 540 Myxicolinae 524 Myzostoma 34, 47, 62, 63, 442, 463, 474 Myzostomida 10, 13, 47, 61, 62, 143, 144, 153, 155, 432, 442, 468, 474

N Naiades 48, 450, 466 Naididae + Tubificidae 237 Naidinae 248, 249, 250, 260, 296, 299, 346 Naineris 46 Nais 346 Namalycastis 444 Nanos 98, 107, 112, 113, 132, 413, 418, 419, 421, 422 Narapa 251, 300 Narapidae 251, 252, 260, 299, 300 Natatory Capillaries 511 Nautiliniellidae 13, 155, 431 Neanthes 47, 82, 86, 88, 249, 444, 464, 469, 474 Nectochaete 149, 151, 439, 446, 474, 475, 481, 587, 588, 589 Nectohelmis 642 Nectosoma 149 Negative Feedback 507 Nematogenia 270, 326, 348 Nematomorphs 339

Nemertea 5 Neoclitellata 260, 261 Neodexiospira 51, 530, 537, 545, 549 Neoleanira 454 Neomenia 161 Nephridia 11, 118, 128, 252, 265, 327, 371, 377, 412, 416, 417, 421, 435, 436, 468, 504, 505, 510, 516, 525, 581, 583, 586, 645, 647, 648 Nephridioblast 377 Nephridiopores 373, 505, 583, 584, 585 Nephroblast 377 Nephromixia 435 Nephrostome 435, 574, 583 Nephtyidae 13, 24, 47, 143, 153, 155, 163, 432, 433, 436, 442, 443, 468, 474 Nephtys 47, 163, 442, 443, 462, 468, 474 Nereididae 13, 24, 25, 38, 47, 143, 153, 155, 158, 160, 163, 431, 433, 436, 443, 444, 445, 446, 464, 469, 474 Nereidiformia 10, 431, 446, 447, 469 Nereimyra 440 Nereis 47, 82, 86, 88, 99, 102, 106, 124, 125, 126, 127, 191, 248, 340, 445, 464, 469, 474 Nerilla 14, 48, 644, 651, 657, 658 Nerillidae 10, 13, 48, 87, 156, 639, 640, 643, 650, 651, 657, 658 Nerillidium 643, 653, 655, 658 Nerillidopsis 658 Nerininae 566, 567, 569, 571, 572, 582, 591, 624 Netrin 413, 418, 420 Neurochaetal Spines 507 Neurons 114, 123, 126, 130, 132, 412, 416, 417, 418, 420, 421 Neurotroch 142, 146, 148, 149, 166, 480, 513, 549, 590, 592, 594, 596, 597, 599, 600, 609, 611, 615, 616, 618, 619, 622, 623 Nicolea 26, 49 Nk-2 113, 116, 413, 417, 422 Nos 98, 107, 112, 113, 132, 413, 415, 418, 419, 421 Notaulax 50 Nothria 163 Notophyllum 432, 450 Notoscolex 265, 282, 283, 365 Nototroch 142, 146, 149, 611 Novafabricia 51 Nuage 29, 31 Nuchal Organs 4, 5, 8, 9, 11, 17, 151, 480, 511, 589, 592, 609, 611, 615, 616, 619, 641 Nuclear Cone 461 Nuclear Fragmentation 360 Nuclear Morphogenesis 337, 357 Nucleoli 29 Nucleus 28, 29, 31, 33, 36, 55, 57, 58, 59, 60, 62, 63, 96, 118, 265, 266, 267, 307, 311, 322, 331, 333, 335, 336, 337, 338, 339, 340, 341,

Index 342, 343, 345, 346, 348, 350, 351, 352, 354, 355, 356, 360, 404, 405, 434, 462, 463, 464, 465, 466, 467, 508, 509, 515, 527, 532, 533, 579, 580, 598, 644, 650, 651, 652, 653, 655, 656, 659 Nurse Cells 25, 27, 28, 29, 37, 38, 39, 329, 403, 461, 530, 575, 650 Nvi-post1 125, 126, 127

O O or P Fate 410 Ocnerodrilidae 237, 240, 244, 258, 259, 261, 263, 270, 273, 276, 281, 282, 285, 287, 312, 325, 326, 348, 361, 365 Ocnerodrilinae 263, 325, 363 Ocnerodriloidea 261 Ocnerodrilus 281, 288, 311 Ocotogonadal 303 Octochaetidae 262, 263 Odontosyllis 457, 471 Oenonidae 9, 13, 155 Olavius 251 Oligobrachia 530, 550 Oligochaeta 3, 6, 7, 8, 179, 235, 236, 237, 240, 245, 250, 255, 257, 263, 266, 274, 275, 279, 280, 284, 288, 291, 297, 301, 302, 303, 311, 313, 316, 318, 320, 324, 331, 339, 344, 366, 368, 394 Oligochaete 37, 78, 94, 96, 98, 99, 107, 109, 110, 235, 236, 240, 244, 245, 246, 263, 265, 267, 268, 270, 274, 290, 293, 296, 298, 328, 333, 337, 338, 339, 340, 341, 346, 348, 349, 350, 352, 353, 358, 360, 361, 368, 371, 373, 376, 393, 394, 400, 408, 410 Oligolecithal 368 Onuphidae 9, 13, 24, 25, 39, 47, 143, 144, 153, 155, 163, 640 Onuphis 47 Oocyte 23, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 39, 78, 95, 184, 269, 307, 309, 327, 328, 329, 331, 367, 369, 403, 413, 417, 418, 437, 510, 573, 575, 578, 646, 647, 648, 650, 659 Oogenesis 23, 24, 25, 26, 27, 29, 32, 37, 38, 39, 104, 187, 309, 327, 328, 331, 360, 403, 436, 437, 438, 440, 442, 444, 446, 448, 450, 452, 454, 456, 458, 460, 461, 507, 508, 529, 530, 565, 573, 574, 576, 627, 649, 650, 651 Oogonia 26, 27, 37, 328, 329, 331, 333, 403, 461, 651 Oogonial Polyplasts 27 Oogonium 27, 328, 329 Oolemma 29, 34, 329 Ooplasmic Rearrangement 96 Oosthuizobdella 398 Ophelia 148, 149, 161, 165, 575, 585, 608, 620 Opheliidae 11, 13, 24, 46, 52, 144, 145, 153, 154, 160, 161, 165, 641

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Ophiodromus 163, 441, 462, 472 Ophryotrocha 14, 25, 46, 78, 79, 80, 82, 83, 84, 85, 86, 87 Opisthodrilus 270, 312 Opisthopora 242, 243, 245, 252, 254, 260, 281, 307 Opistocystidae 245, 252, 260, 296 Opposed-band 141, 144, 145, 146, 147, 151, 153, 160, 163, 165, 166, 186 — feeding 141, 145, 153, 163, 165, 166, 186 — trochophore 144, 147, 160 Oral Brush 142, 146, 149, 155, 156, 163, 166, 474, 475 Orbiniidae 13, 24, 46, 56, 142, 143, 144, 148, 153, 154, 161, 235, 641 Ordovician 12 Organogenesis 369, 376, 420 Origin of Ectoderm and Mesoderm 371 Orthochromatic Mucous Cells 305 Orthodenticle 113, 413, 418 Osedax 15, 52, 526, 528, 529, 530, 547 Otx 113, 114, 115, 418, 420, 422 Ovaries 23, 24, 25, 27, 30, 32, 36, 38, 39, 245, 253, 254, 268, 274, 293, 294, 296, 297, 298, 300, 302, 303, 304, 307, 309, 311, 312, 315, 316, 319, 322, 324, 325, 327, 329, 373, 400, 401, 417, 437, 461, 468, 504, 505, 507, 508, 513, 526, 529, 530, 573, 574, 604, 643, 647, 648, 649, 651 Oviducal Funnels 307, 322 Oviduct 295, 296, 315, 316, 402, 468, 526, 644, 647, 648 Ovisac 27, 37, 268, 315, 316, 327, 328, 331, 504, 508, 526, 648, 649 Owenia 49, 141, 145, 149, 165, 523, 525, 528, 530, 538, 548 Oweniidae 8, 10, 13, 25, 49, 144, 145, 149, 153, 157, 158, 160, 165, 196, 521, 522, 523, 526, 528, 529, 530, 533, 538, 550 Oxyptychus 398, 401 Ozobranchidae 394, 400, 401

P Pachytene 29 Paddle-shaped Claspers 321 Pair Bonds 88 Paleanotus 438 Paleozoic 399 Palp 8, 10, 599, 600, 616, 618, 619, 624, 631, 646 Palpata 8, 9, 13, 161, 162, 164 Panthalis 438 Paradexiospira (Spirorbides) 533 Parafabricia 51 Parahox 413, 418, 422 Paralacydonia 13, 155, 163, 432, 447 Paralaeospira 51, 536 Paralvinella 49, 66, 249

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Reproductive Biology and Phylogeny of Annelida

Paranais 267, 346 Paranaitis 432 Paranerilla 657, 658 Paraonidae 11, 13, 64, 154, 568 Paraphyletic 6, 7, 8, 10, 17, 236, 253, 255, 256, 258, 259, 261, 262, 266, 267, 303, 340, 394, 396, 398, 500, 501, 502, 522, 524, 525, 569 Parapionosyllis 457 Parapodia 8, 17, 24, 126, 127, 129, 131, 147, 149, 433, 435, 468, 469, 471, 475, 480, 600, 611, 614, 616, 619, 647, 659 Paraprionospio 571, 582 Paraprotis 51, 85, 527, 531, 533, 536, 542, 549 Paraprotula 536, 542 Parascolelepis 571, 575, 579, 580, 582, 586, 589, 593, 596, 597, 598 Paraspermatid 357, 359, 360 Paraspermatozoa 60, 292, 340, 350, 353, 355, 356, 357, 359, 360, 361, 653, 657 Paratomy 498, 512, 514, 550, 628, 629, 631, 647 Parental Care 87, 88, 180, 191, 406, 438, 440, 442, 444, 446, 448, 450, 452, 454, 456, 458, 460, 658 Parenterodrilus 52, 641, 645, 646, 647, 652, 653, 657 Parergodrilidae 12, 13, 52, 56, 57, 66, 157, 501, 639, 641, 648, 651, 655, 658, 660 Parergodrilus 52, 57, 66, 85, 648, 652, 655, 658, 660 Paronuphis 29 Parsimony 5, 236, 237, 238, 240, 242, 255, 257, 262, 263, 264, 265, 267, 298, 355, 568 Parvidrilidae 251, 252, 260, 261, 298, 299 Parvidrilus 251, 298, 299 Patagoniobdella 398, 401 Pd-bra 113, 115 Pd-gsc 113, 115 Pd-otx 113, 114 Pectinaria 149, 165 Pectinariidae 11, 13, 24, 25, 49, 52, 147, 149, 153, 156, 160, 165, 166, 521, 522 Pectinodrilus 345 Pelagobia 442 Pelodrilus 253, 302, 348 Penes 261, 271, 296, 298, 299, 361, 363, 462, 466, 468, 469 Penial Bulb 298, 649 Penial Chaetae 271, 272, 273, 309, 311, 325, 362, 363 Penis 273, 274, 277, 298, 311, 362, 363, 398, 399, 402, 403, 406, 643, 644, 647, 656 Perforatorium 336, 342, 343, 345, 404, 656 Perinereis 88, 445, 446, 464 Periodicity 266, 421, 507 Perionychella 247, 248, 250 Perionychini 264 Perionyx 263 Peristodrilus 272

Peristomium 4, 9, 10, 113, 114, 129, 144, 376, 408, 521, 535, 595, 609, 618, 623 Perkinsiana 50, 535, 540, 551 Petitia 48, 457, 464 Phagocytosis 462 Phalacrostemma 538 Phallodrilinae 251, 267, 292, 345 Pharynx 9, 57, 153, 253, 376, 377, 589, 590, 592, 593, 594, 596, 609, 616, 621, 660 Pheretima 247, 250, 271, 277, 326, 352, 360 Pherusa 499, 501, 505, 508, 510, 514 Pholoe 48, 163, 436, 447, 448, 475 Pholoidae 13, 25, 38, 48, 143, 146, 153, 155, 163, 436, 447, 448, 475 Pholoides 448 Phoronida 6 Phragmatopoma 32, 34, 50, 196, 198, 199, 216, 217, 218, 221, 222, 529, 530, 538, 548 Phreodrilidae 244, 245, 246, 251, 252, 260, 296, 313, 331, 343 Phreodrilus 265, 333, 343, 345 Phreoryctidae 254 Phyllochaetopterus 601, 602, 603, 628, 629 Phyllodoce 31, 34, 249, 432, 450, 471, 480, 481 Phyllodocida 9, 10, 12, 13, 17, 38, 47, 48, 53, 64, 66, 143, 149, 153, 155, 156, 160, 163, 431, 432, 433, 435, 436, 438, 461, 466, 467, 472, 480, 482 Phyllodocidae 13, 24, 25, 38, 39, 48, 143, 149, 153, 156, 163, 432, 433, 435, 436, 448, 449, 450, 451, 471, 475 Phylogenetics 254, 301, 399 Phylogeny 1, 3, 38, 66, 93, 109, 110, 160, 235, 237, 239, 245, 255, 257, 265, 267, 311, 331, 340, 346, 348, 393, 394, 397, 398, 431, 432, 482, 500, 521, 524, 566, 571, 573, 586, 639, 640 Phylogeny of the Oligochaete Families 245 Pickfordia 327 Piercing Chaetae 272 Pilargidae 13, 48, 143, 156, 431, 452 Pileolaria 51, 530, 536, 537, 547, 549 Pionosyllis 457 Piromis 510 Piscicolidae 394, 396, 400, 401, 406 Pisione 48, 163, 452, 461, 471, 482 Pisionidae 13, 48, 143, 147, 156, 160, 163, 431, 436, 452, 471, 482 Pisionidens 452, 471, 482 Placobdella 396, 398, 400, 407 Placobdelloides 395, 398 Placostegus 532, 542 Planktonic Larva 195 Planktotrophic 95, 146, 149, 151, 153, 158, 163, 165, 179, 186, 188, 189, 190, 191, 192, 193, 328, 438, 439, 440, 441, 442, 446, 447, 448, 450, 452, 453, 454, 455, 459, 472, 474, 475, 480, 498, 508, 528, 530, 536, 537, 538, 539,

Index 540, 541, 542, 543, 548, 549, 551, 604, 605, 607, 618, 619, 621, 626, 627, 659 — development 151, 179, 186, 191, 193, 475, 508, 605 — eggs 328 Planktotrophy 151, 154, 156, 158, 160, 186, 187, 190, 193 Platelet 38, 39, 398 Platynereis 47, 66, 82, 94, 96, 106, 113, 114, 115, 123, 163, 437, 446, 464, 469, 474 Plesiomorphy 4, 5, 27, 38, 56, 64, 65, 66, 116, 147, 153, 160, 165, 179, 193, 240, 241, 254, 261, 265, 274, 281, 293, 296, 298, 303, 304, 309, 311, 325, 327, 337, 340, 341, 343, 348, 351, 368, 396, 403, 408, 410, 412, 482, 532, 535, 537, 550, 551, 576, 639 Plotohelmis 48, 451 Podarke 99, 473 Podarkeopsis 460, 462 Poecilochaetidae 565, 567, 568, 569, 572 Poecilochaetus 11, 149, 157, 565, 567, 568, 569, 571, 572, 573, 575, 578, 582 Poecilogony 192, 498, 604, 607, 622, 624, 626, 627, 628 Poeobiidae 11, 497, 501, 502 Poeobius 13, 497, 499, 500, 501, 502, 503, 505, 508, 509, 510 Pogonophora 4, 5, 6, 7, 10, 58, 60, 61, 62, 521, 522, 525 Polar Bodies 96, 189, 367, 369, 371, 408, 659 Polar Lobe 100, 102, 103 Polarity in the Primary Oocyte 367 Pole Plasm 96, 367, 371 Polybrachia 530 Polychaetes 3, 5, 7, 8, 9, 10, 11, 12, 13, 17, 23, 24, 25, 27, 29, 32, 37, 38, 39, 52, 53, 55, 56, 57, 63, 64, 78, 82, 83, 84, 85, 86, 87, 88, 95, 96, 98, 102, 104, 106, 108, 109, 110, 111, 114, 115, 117, 119, 120, 123, 124, 125, 127, 130, 131, 132, 142, 143, 144, 145, 146, 151, 160, 166, 179, 180, 182, 183, 186, 188, 189, 191, 192, 195, 196, 197, 199, 210, 235, 236, 237, 248, 249, 250, 268, 293, 298, 328, 329, 339, 340, 367, 368, 432, 433, 435, 436, 464, 466, 471, 497, 501, 508, 509, 521, 522, 525, 551, 565, 567, 568, 569, 571, 576, 581, 584, 602, 623, 624, 629, 632, 639, 640, 641, 649, 660 Polydora 31, 34, 49, 84, 86, 166, 190, 192, 221, 566, 567, 568, 569, 571, 573, 574, 575, 576, 577, 579, 580, 581, 583, 584, 585, 586, 612, 618, 620, 621, 622, 626, 629, 631 Polydorella 571, 629, 630, 631, 632 Polygordiidae 11, 12, 13, 52, 142, 144, 157, 158, 160, 165, 196, 639, 645, 653, 657, 659 Polygordius 52, 99, 152, 158, 165, 641, 645, 650, 653, 659 Polynoe 454

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Polynoidae 13, 24, 48, 52, 87, 143, 146, 149, 153, 156, 163, 191, 452, 453, 454, 475, 508 Polyophthalmus 29 Polytoreutus 292 Pomatoceros 158, 525, 526, 529, 530, 537, 542, 548 Pomatoleios 51, 530, 532, 542 Pontodora 13 Pontodrilus 246, 247, 248, 249, 250 Pontoscolex 246, 256, 257, 258, 270, 273 Population Density 83, 87, 536 Potamodrilidae 12, 13, 59, 65, 639, 641, 647 Potamodrilus 12, 52, 58, 59, 65, 157, 340, 642, 648, 655, 658, 660 Potamothrix 271, 272 Precopulatory Behavior 363 Premeiotic Doubling 360, 361 Presegmental 144, 146, 475, 602 Primary Oocyte 307, 309, 327, 328, 329, 367, 369 Primary Spermatocytes 333, 335, 510 Primitive Sperm 53, 55, 56, 57, 337, 578 Primordial Germ Cells (Protogonia) 331 Prionospio 49, 66, 566, 567, 568, 569, 571, 575, 579, 580, 582 Pristina 123 Proacrosome 336 — Cap 404 — Vesicle 404, 405 Probe Excess Titration 125 Proboscipedia 128 Proboscis 10, 88, 113, 116, 303, 304, 394, 396, 412, 415, 417, 422, 472, 474, 480 Proceraea 457, 458 Procerastea 458 Progenetic 640 Progenetically 640 Pronucleus 408, 510, 659 Propappidae 246, 253, 260, 296, 297, 298, 300 Propappus 246, 251, 253, 297, 298 Propheretima 263, 277 Prostate gland 240, 261, 262, 263, 273, 274, 276, 281, 282, 283, 285, 294, 295, 296, 297, 298, 304, 307, 309, 311, 312, 315, 316, 318, 319, 322, 323, 324, 325, 326, 327, 361, 362, 365, 378, 648, 649, 658 Prostate-like Glands 309, 316, 319, 322, 323 Prostomium 4, 10, 17, 112, 114, 129, 152, 303, 304, 374, 376, 406, 417, 420, 466, 469, 480, 521, 566, 588, 589, 590, 591, 593, 595, 596, 597, 602, 604, 607, 609, 611, 615, 616, 618, 619, 622, 623, 624, 645 Protandric 83, 468, 507, 512, 513, 514, 515, 528 — hermaphroditism 528 Protandry 82, 83, 268 Protodrilida 11, 52, 56, 157, 639, 641, 645, 653, 657, 659, 660

$&" Reproductive Biology and Phylogeny of Annelida Protodrilidae 11, 12, 13, 146, 153, 160, 646, 652 Protodriloides 52, 641, 645, 653, 655, 657, 659, 660 Protodriloididae 11, 12, 13, 153, 160 Protodrilus 52, 165, 360, 641, 645, 646, 653, 655, 657, 660 Protogyny 83 Protolaeospira 51, 533, 536, 546 Protonephridia 377, 435, 436, 468 Protonephromixia 435, 436 Prototroch 104, 109, 113, 126, 127, 142, 143, 144, 145, 146, 147, 148, 149, 151, 153, 155, 158, 163, 165, 166, 186, 196, 368, 474, 475, 480, 516, 548, 549, 550, 587, 588, 589, 590, 591, 592, 593, 594, 595, 596, 597, 598, 599, 600, 602, 606, 607, 609, 612, 613, 614, 615, 616, 618, 619, 622, 623, 660 Protrochophore 147, 151 Protuberodrilus 272 Protula 532, 536, 540, 549 Provisional Epithelium 111, 112, 409, 410, 412, 413, 415, 417, 420 Psamathe 32, 437, 441 Psammodrilidae 12, 13, 153, 157 Psammolyce 48, 434, 454, 461 Psammoryctides 271, 272, 353 Pseudatherospio 568, 571 Pseudochitinopoma 542 Pseudocopulation 443, 448, 468, 471, 657 Pseudoeurythoe 26 Pseudofabricia 51 Pseudofabriciola 51 Pseudomystides 451 Pseudopodia 462 Pseudopolydora 49, 199, 571, 580, 581, 584, 620, 622, 626, 627 Pseudopotamilla 15, 50, 532 Pseudovermilia 536, 542 Pygidium 4, 129, 468, 587, 589, 590, 593, 599, 602, 609, 611, 614, 615, 616, 619, 620, 621 Pygmaeodrilus 274, 285, 287, 326 Pygospio 566, 567, 569, 571, 575, 579, 581, 583, 584, 586, 606, 607, 608, 610, 626, 627, 628, 629, 630 Pygospiopsis 568, 569, 571, 572, 575, 623, 624, 625, 632

Q Questa 46, 235 Questidae 13, 46, 64, 153, 154, 641

R Ramex 49 Randiellata 245, 260, 293, 294 Randiellidae 241, 245, 260, 273, 293, 300 Random Mating 83, 84, 85

Raphidrilus 498, 501, 507, 508, 511, 512, 513 rDNA 57, 236, 237, 238, 239, 240, 246, 250, 253, 256, 257, 263, 264, 298, 340, 397, 502, 641, 642 Reciprocity 78, 80 Regeneration 116, 123, 296, 628, 630, 631 Rel 98, 113, 413 Reproductive Stylet 507, 512 Reproductive Success 77, 78, 80, 82, 83, 85 Reproductive System 8, 252, 268, 293, 294, 298, 300, 302, 303, 304, 305, 309, 313, 315, 350, 399, 400, 433, 464, 504, 510, 525, 526, 573, 578, 642 Rer 29, 32, 33, 36, 37, 328, 333, 335 Respiratory Auricles 395 Rheomorpha 642 Rhizodrilus 265, 272, 273, 292, 340, 346, 350 Rhododrilus 262, 326 Rhodopsis 529, 533, 536, 540, 549, 550 Rhyacodrilinae 250, 251, 267, 292, 294 Rhyacodrilus 272, 292, 294, 295, 342, 343 Rhynchelmis 248, 249, 250, 267, 301, 302, 346, 348, 368, 376, 377 Rhynchobdellida 292, 394, 396, 403 Rhynchonerella 48, 451, 461 Rhynchospio 571, 575, 582 Ribosomes 328, 335, 408 Ridgeia 52, 60, 62, 149, 533, 548, 550 Riftia 52, 60, 62, 530, 533, 548, 550 Righiella 309, 311 RNA 94, 107, 240, 329, 413 Rod 253, 265, 336, 337, 340, 342, 343, 345, 346, 348, 350, 351, 352, 356, 462, 464, 651, 653, 655, 656 Romanchella 51, 523, 536, 546 Rosettes 53, 322 Rostraria 145, 149, 151 Rynchonerella 26

S Sabella 34, 50, 532, 535, 540, 550 Sabellaria 50, 102, 103, 105, 106, 145, 165, 196, 199, 528, 538, 539 Sabellariidae 10, 11, 13, 24, 49, 144, 145, 153, 157, 160, 165, 199, 521, 522, 523, 525, 526, 528, 529, 530, 533, 538, 539, 548, 550 Sabellariinae 522 Sabellida 10, 13, 49, 50, 51, 52, 53, 56, 58, 66, 157, 521, 522, 523, 526, 538, 550 Sabellidae 10, 13, 24, 25, 37, 50, 53, 54, 56, 87, 142, 146, 147, 153, 157, 158, 160, 165, 521, 522, 523, 524, 525, 526, 528, 532, 533, 535, 539, 540, 550 Sabellinae 50, 53, 524, 528, 532, 535, 539, 540, 550 Sabellonga 522 Saccocirridae 11, 13, 144, 145, 151, 153, 160, 165

Index Saccocirrus 52, 145, 165, 641, 644, 645, 647, 652, 653, 655, 657, 660 Salifidae 395, 401, 403 Salmacina 51, 54, 86, 196, 526, 527, 529, 533, 534, 536, 541, 549, 550 Salvatoria 458, 465 Scaleworms 10, 431, 432, 436, 467, 475, 477, 482 Scalibregmatidae 13, 46, 154, 160 Schizogami 456 Schizogamous Epitoky 433 Scissiparity 433, 471 Sclerolinum 529, 550 Scolecida 8, 11, 12, 13, 46, 64, 66, 154 Scolecolepides 571, 575, 582, 586 Scolecolepis 165 Scolelepis 566, 567, 568, 571, 575, 579, 582, 589, 591, 595, 596, 598 Scoloplos 14, 99, 109, 110 Secondary Spermatocytes 333, 464 Secondary Tube 253, 265, 335, 336, 340, 342, 343, 345, 346, 348, 351, 352 Sedentaria 7, 522 Segment 4, 11, 83, 116, 118, 119, 120, 121, 122, 123, 126, 127, 128, 130, 132, 145, 148, 242, 252, 253, 254, 258, 259, 265, 270, 272, 273, 274, 277, 280, 281, 283, 284, 285, 286, 288, 293, 294, 295, 296, 297, 298, 299, 300, 302, 303, 304, 307, 309, 311, 312, 315, 316, 318, 319, 322, 323, 324, 325, 326, 327, 331, 361, 362, 363, 371, 373, 376, 406, 409, 412, 416, 417, 418, 419, 421, 435, 437, 462, 467, 468, 469, 474, 475, 480, 482, 497, 504, 505, 507, 525, 526, 549, 565, 573, 580, 587, 590, 592, 593, 596, 599, 600, 609, 611, 614, 615, 616, 619, 620, 621, 623, 643, 645, 647, 648 — polarity 11, 120, 122, 132, 419 Segmental Organs 9, 11, 433, 435, 436, 467, 468, 471, 500, 502, 504, 525, 526, 535, 536 Segmentation 4, 24, 114, 116, 117, 120, 121, 122, 123, 125, 126, 127, 128, 131, 147, 160, 305, 373, 375, 376, 419, 422, 601, 602, 612, 630, 645 — genes 120, 121, 122, 422 Segmentation of the Ectoderm 373 Self-fertilisation 536 SEM 144, 369, 480, 514, 531, 534, 625, 646, 659 Semelparity 24 Seminal Funnels 302, 303, 304, 316, 366 Seminal Groove 274, 277, 326, 362, 363 Seminal Receptacle 401, 581, 584, 647, 648 Seminal Vesicle 304, 307, 464, 507, 512, 647, 648, 649, 655 Semiscolescidae 395 Semiscolex 398 Semivermilia 543 Sensillae 416, 420 Septate Junctions 284, 353, 355

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Sequential Hermaphroditism 77, 82, 83 Serpula 51, 532, 543, 548, 549 Serpulidae 10, 12, 13, 24, 25, 51, 52, 53, 54, 87, 144, 146, 147, 153, 157, 160, 165, 191, 521, 522, 523, 524, 525, 526, 527, 528, 531, 532, 533, 534, 537, 540, 541, 542, 543, 544, 545, 546, 547, 549 Serpulinae 51, 523, 525, 528, 541, 542, 543, 549, 550 Sex Allocation 77, 78, 84 Sex Change 86, 87 Sex Determination 85, 86, 650 Sex Ratio 83, 84, 85, 86, 87, 88 Sexual Biology 398 Sexual Dimorphism 85, 504, 505, 512 Sexual Selection 82, 83, 85 Sheath Cells 437 Shizogamy 471 Sib Competition 78 Sibling Species 39, 192, 437, 576, 582, 612, 615, 627 Siboglinidae 6, 10, 13, 52, 56, 58, 60, 61, 146, 149, 153, 157, 165, 521, 522, 525, 526, 529, 530, 533, 547, 549, 550 Siboglinum 52, 60, 61, 165, 529, 530, 533, 537, 550 Sigalion 48, 454, 461 Sigalionidae 13, 48, 143, 146, 156, 163, 191, 454, 455, 475 Sigambra 48, 464 Sige 451 Simultaneous Hermaphroditism 77, 78, 468, 528, 529 Sinohesione 441, 468 Sipuncula 5, 6, 64, 147, 161, 502 Sipunculus 161 Sirsoe 47, 437, 441, 462 Skeletal Elements 647 Slavina 346 Smithsonidrilus 251, 355 Smooth Endoplasmic Reticulum 335 Snail 88, 112, 113, 114, 413, 418, 422 Social Interactions 83 Somite Formation 376 South American 251, 398, 401 Sparganophilidae 244, 255, 259, 260, 273, 275, 280, 319, 322, 324, 348 Sparganophiloidea 256, 258, 260 Sparganophilus 255, 256, 257, 259, 261, 270, 271, 275, 281, 282, 303, 319, 321, 324, 345, 348, 351 Spenceriella 263, 270, 274, 280, 352, 362, 363, 365 Sperm 8, 23, 37, 38, 45, 46, 53, 55, 56, 57, 58, 59, 60, 61, 62, 63, 64, 65, 66, 77, 78, 80, 81, 82, 83, 84, 85, 87, 88, 189, 240, 246, 252, 261, 265, 266, 267, 268, 270, 271, 272, 273, 274, 285, 288, 289, 290, 292, 296, 298, 307, 309,

$&$ Reproductive Biology and Phylogeny of Annelida 311, 313, 315, 322, 226, 327, 328, 335, 337, 338, 339, 340, 341, 343, 345, 346, 348, 350, 351, 352, 353, 355, 356, 357, 358, 359, 361, 366, 378, 393, 398, 399, 400, 401, 402, 403, 404, 406, 434, 438, 440, 442, 444, 446, 448, 450, 452, 454, 456, 458, 460, 461, 462, 463, 464, 465, 466, 467, 468, 469, 471, 472, 498, 500, 507, 508, 509, 510, 511, 512, 515, 524, 525, 526, 527, 528, 529, 530, 532, 533, 535, 537, 538, 540, 542, 544, 546, 548, 550, 565, 566, 571, 578, 579, 581, 582, 583, 584, 586, 626, 627, 643, 646, 648, 651, 653, 655, 656, 657, 658 — digestion 80, 81 — entry 366 — storage organs 81 — transfer 64, 81, 271, 272, 288, 461, 532, 578, 583, 656, 657 Spermathecae 55, 56, 57, 64, 80, 268, 270, 271, 273, 281, 284, 299, 300, 304, 307, 311, 312, 313, 315, 316, 319, 322, 323, 325, 326, 349, 353, 361, 366, 462, 533, 535 Spermathecal — Ampulla 307, 309, 315 — Atrium 269, 284, 307, 315 — Chaetae 271, 272, 363 — Duct 284, 285, 307, 309 — Pores 246, 271, 272, 277, 280, 281, 296, 298, 309, 312, 315, 316, 319, 322, 323, 325, 326, 327, 361, 362 Spermatids 45, 53, 54, 58, 60, 62, 293, 306, 307, 331, 331, 335, 337, 338, 339, 357, 360, 361, 434, 462, 463, 466, 467, 505, 509, 526, 527, 528, 530, 532, 533, 548, 649, 651, 653, 655, 656 Spermatocyte 53, 335, 361 Spermatogenesis 37, 45, 52, 309, 328, 331, 335, 349, 350, 352, 357, 360, 361, 366, 393, 403, 405, 461, 464, 466, 508, 509, 532, 578, 579, 649, 651, 655 Spermatogonia 45, 52, 53, 293, 331, 333, 360, 361, 403, 464, 509, 655 Spermatophore 60, 62, 63, 81, 290, 292, 315, 398, 406, 581, 583, 646, 647, 657 Spermatozeugmata 60, 62, 292, 352, 353, 355, 356, 360, 529, 533, 548, 655, 657 Spermatozoa 45, 53, 63, 266, 267, 268, 274, 284, 285, 287, 288, 290, 292, 293, 307, 311, 331, 333, 336, 337, 338, 339, 340, 346, 348, 349, 351, 352, 353, 360, 361, 366, 404, 405, 406, 509, 510, 580, 641, 642, 643, 644, 646, 647, 648, 650, 651, 652, 653, 655, 656, 657, 658 Spermiocysts 62, 463, 468 Spermioducts 642, 643, 644, 645, 647, 648 Spermiogenesis 45, 53, 57, 58, 60, 62, 63, 331, 333, 337, 338, 351, 357, 358, 359, 360, 361, 463, 530, 532, 533, 565, 579, 642, 651, 653 Sphaerodoridae 13, 24, 25, 38, 156, 158, 160, 432, 455, 480

Sphaerodoropsis 480 Sphaerodorum 437, 455, 480 Sphaerosyllis 48, 82, 458, 464, 478 Spindle 100, 102, 271, 359, 529 Spinther 10, 13, 16, 156 Spio 566, 567, 569, 571, 573, 575, 576, 577, 579, 583, 586, 626 Spiochaetopterus 601, 602, 603 Spionida 10, 11, 13, 49, 66, 157, 160, 521, 565, 566, 568, 641 Spionidae 11, 12, 13, 24, 37, 49, 56, 149, 153, 157, 165, 166, 565, 566, 567, 568, 569, 571, 572, 575, 579, 580, 584, 586, 591, 624 Spioninae 566, 567, 568, 569, 571, 572, 579, 585, 624, 628 Spiophanes 566, 667, 568, 569, 571, 575, 579, 580, 582, 590, 591, 592, 593, 594 Spiral Cleavage 97, 99, 100, 112, 368, 369 Spirobranchus 51, 144, 196, 532, 534, 543, 548 Spirorbidae 521, 522 Spirorbinae 51, 182, 191, 523, 524, 525, 527, 531, 534, 536, 543, 544, 545, 546, 547, Spirorbis 51, 82, 221, 533, 536, 537, 543, 544, 549 Stereoblastula 111 Sternaspidae 497, 501, 502 Sternaspis 11, 13, 143, 147, 153, 156, 499, 502, 505, 506, 508, 510, 513, 514, Sthenelais 163, 455, 475 Stolons 87, 433, 465, 470, 471, 480, 628, 629, 631 Stomodeum 109, 111, 113, 114, 115, 376 Stratiodrilus 47, 66 Streblosoma 49, 66 Streblospio 32, 33, 35, 49, 192, 193, 571, 573, 575, 576, 577, 579, 580, 583, 584, 604, 606, 623, 626 Stygocapitella 57, 648, 652, 656, 658 Stylaria 124, 267, 294, 346, 369 Stylet 507, 512, 647 Subacrosomal Space 60, 348, 350, 351, 465, 509, 532, 533 Sucker 240, 321, 393, 395, 412, 417, 421 Swarming 438, 439, 441, 443, 444, 445, 446, 456, 457, 458, 459, 468, 469, 471 Syllidae 13, 24, 48, 52, 56, 87, 143, 146, 153, 156, 158, 160, 163, 431, 433, 435, 455, 456, 457, 458, 459, 460, 471, 480, 658 Syllides 458 Syllidia 411, 462 Syllis 84, 87, 163, 458, 459, 461, 465, 480 Symbiotic Bacteria 525 Symplesiomorphy 251, 253, 296 Synapomorphg 9, 11, 254, 255, 261, 304, 340, 393, 394, 396, 524, 640, 655 Synaptonemal Complex 28 Syncytium 53, 376 Syngenodrilidae 254, 255, 260, 304, 309 Syngenodrilus 255

Index

T Tagmosis 127 Tannic Acid 338, 354 Taxonomy 8, 242, 277, 399, 601 Taylorpholoe 48, 448, 461, 475 Tectidrilus 251 Teeth 394, 596 Teloblast Mother Cell 111 Teloblastogenesis 369, 373 Teloblasts 96, 109, 111, 112, 113, 114, 117, 118, 120, 121, 125, 131, 367, 371, 373, 374, 376, 377, 408, 410, 411, 412, 415, 416, 417, 419 Teloplasm 95, 96, 97, 107, 111, 112, 408, 409, 410, 413, 415, 417, 418 Telotroch 142, 146, 148, 149, 158, 474, 480, 516, 548, 587, 588, 589, 590, 592, 593, 594, 595, 596, 597, 599, 606, 609, 611, 612, 613, 614, 615, 616, 617, 618, 619, 621, 622, 623, 624 Terebella 26 Terebellida 10, 11, 13, 521, 522 Terebellidae 11, 13, 24, 25, 49, 52, 147, 147, 158, 165 Terebellides 15, 165 Terebelliformia 11, 13, 49, 66, 156, 157 Terebrasabella 50, 82, 535, 540 Terrestrial 3, 12, 57, 85, 237, 244, 246, 254, 255, 256, 257, 269, 273, 322, 395, 396, 398, 401, 402, 639 Terrimegadrili 244, 255, 273 Testes 27, 38, 45, 52, 53, 60, 243, 245, 246, 252, 253, 254, 268, 273, 280, 293, 294, 296, 297, 298, 299, 300, 301, 302, 303, 304, 305, 307, 309, 311, 312, 315, 318, 319, 322, 324, 325, 333, 373, 463, 468, 526, 647, 648, 649, 655 Testis 227, 280, 281, 293, 294, 295, 298, 300, 302, 303, 304, 305, 306, 307, 309, 311, 312, 315, 318, 319, 322, 331, 504, 505, 509, 510, 526, 642, 649 Testis-sac 306, 307, 399, 400, 401, 403 Tetrads 53, 401, 434, 463, 464, 527, 530, 532, 651, 653 Tetragon Fibers 339, 341, 343, 345, 346, 348, 351, 352, 356 Thalassodrilides 251, 355 Thalassodrilus 343 Tharyx 498, 504 Thecacysts 290, 292 Theromyzon 94, 95, 99, 350, 400, 405, 408, 416 Thrombin 398 Tiguassu 244, 253, 303, 304 Tiguassuidae 242, 253, 260, 303, 304 Tomopteridae 13, 24, 48, 142, 147, 156, 158, 432, 436, 460 Tomopteris 25, 27, 48, 460, 467 Torrea 48, 451, 466 Torresiella 327, 361

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Travisia 46 Triassic 13 Trichobranchidae 13, 158, 165 Trichodrilus 248 Trilobodrilus 46, 642, 643, 651 Triplet 338 Tripolydora 49, 66, 571, 580, 583 Trochoblasts 99, 143, 158 Trochochaeta 11, 157, 565, 567, 568, 569, 571, 572, 575, 582, 589, 598, 599, 600, 623, 624 Trochochaetidae 565, 567, 568, 569, 572 Trochophore 104, 109, 113, 125, 126, 141, 144, 145, 147, 149, 151, 160, 181, 183, 184, 185, 186, 194, 368, 406, 439, 441, 447, 452, 472, 474, 475, 480, 511, 528, 534, 548, 549, 550, 588, 595, 597, 612, 659, 660 Trophosome 526, 550 Trypanosyllis 84, 459, 460 Tubercula Pubertatis 274, 309, 312, 315, 316, 322, 323, 324, 363 Tubifex 88, 96, 98, 99, 102, 107, 109, 110, 112, 117, 248, 249, 251, 328, 329, 337, 345, 352, 353, 354, 355, 356, 357, 358, 359, 360, 366, 367, 368, 369, 371, 373, 374, 375, 376, 377, 378, 410 — siblings 251 Tubificata 244, 245, 260, 293, 294 Tubificida 245, 246, 265, 266, 294 Tubificidae 237, 243, 244, 245, 246, 248, 249, 250, 251, 260, 267, 280, 294, 295, 296, 304, 343, 346, 368, 378 Tubificidae + Naididae 244 Tubificina 294 Tubificoides 251, 294, 353 Tumak 259, 312, 316 Tumakidae 259, 260, 312 Turtles 394, 396, 406 Twi 98, 113, 114, 413, 418 Twist 98, 112, 113, 413, 418 Tylorrhynchus 47, 446, 464, 469 Typhloscolecidae 13, 436 Typosyllis 48, 87, 461

U Uncispio 11, 567, 568, 569, 571, 572, 575, 582 Uncispionidae 11, 565, 567, 568, 569, 572 Uncopherusa 568 Urechis 6, 52

V Vagina 398, 400, 403 Vanadis 33, 48, 451, 466 Vas Deferens 294, 295, 307, 322, 504 Vasa Deferentia 254, 273, 281, 284, 296, 298, 300, 302, 311, 312, 315, 319, 322, 324, 325, 326, 365, 468

$&& Reproductive Biology and Phylogeny of Annelida Vestimentifera 4, 5, 6, 10, 58, 60, 61, 62, 149, 521, 523, 525, 526, 529, 547 Vestimentiferan Pogonophorans 267 Viktoriella 438 Villiersia 253 Vitelline Envelope 328 Vitellogenesis 23, 25, 27, 29, 32, 37, 327, 328, 329, 331, 437, 461, 507, 508, 510, 529, 573, 649 Vitellogenic 23, 26, 27, 29, 30, 31, 32, 34, 35, 36, 37, 39, 328, 507, 530, 643, 648, 650, 651 — mechanisms 23, 39 — oocytes 26, 27, 32, 507, 530, 643, 648, 651 Viviparous 445, 456, 459, 475, 508, 511, 571, 604

Y Yolk 29, 31, 32, 33, 34, 35, 36, 37, 38, 39, 93, 95, 96, 101, 109, 110, 149, 187, 240, 269, 270, 328, 367, 368, 369, 371, 375, 377, 408, 409, 410, 411, 412, 415, 437, 529, 549, 573, 595, 607, 608, 612, 614, 626, 627, 650 — bodies 29, 32, 34, 37, 39

Z

W Whitmania 398 Wnt 98, 108, 413, 415, 418, 419, 420, 422 — genes 108, 422

X Xandaros 568

Xerobdella 399 Xerobdellidae 396, 399, 402 Xironodrilus 249 Xironogiton 248, 249 Xlox 116, 418

Zeppelina 501 Zinc-finger 413 Zipper 111, 346, 352, 356 Zona Pellucida 328, 331, 350 Zonula Collaris 328, 331, 357 Zygote 24, 97, 100, 112, 134, 135, 361, 408, 413, 472, 549 Zygotene 29