Reproductive Biology and Phylogeny of Annelida

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.

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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.

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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.

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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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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.

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%$ 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.

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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.

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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.

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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.

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%$ 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.

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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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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

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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,

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- 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

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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 <103 cells ml–1, >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 <20ppm. Expt. II: 2-cell to blastula. C. survivorship. D. duration of development. Two-cell embryos all died at salinities <20 ppm. Expt. III: Blastula to trochophore. E. survivorship; F. duration of development. Blastulae all died at salinities <25 ppm. Data are plotted as means ±SD of 3 replicate cultures (each started with 40 individuals). From Qiu, J.W. and Qian P.Y. 1997. Marine Ecology Progress Series 152: 79-88, Fig. 1.

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Fig. 7.3 Effects of food concentration, salinity and temperature on survivorship from trochophore to newly-settled juveniles of Hydroides elegans. Expt. IV. Trochophores all died at a salinity of 15 ppm. Data are plotted as means ±SD of 2 replicate cultures (each started with 20 individuals). From Qiu, J.W. and Qian P.Y. 1997. Marine Ecology Progress Series 152: 79-88, Fig. 2.

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&$ Reproductive Biology and Phylogeny of Annelida no such in-situ study has been done on polychaete larvae. Instead, laboratory set-ups are providing the surrogate of complex field experiments. Still, starting from purely observational studies, there have been done quite advanced experimental investigations that elucidate many aspects of polychaete larval trophic ecology (Gosselin and Qian 1997). The majority of polychaete trochophore larvae are lecithotrophic — only a few become planktotrophic particularly at later stages (Levin et al. 1991), but some are short lived and lecithotrophic, some are benthic or demersal, and a few develop completely into juveniles without ever having a freeswimming stage (Qian and Chia 1989). Within many annelid taxa, embryos and larvae obtain extraembryonic nutrition directly from the parent during development or from cells or fluids produced by the parent. During early development, nurse, follicle or test cells are known to provide food to embryos in some polychaete species. Planktotrophy. The larvae of marine invertebrates are generally characterized by their ability to show locomotion and to feed in the sea (Strathmann 1990). This ability has generated a wealth of diversity in larval form and function. Larvae that feed on external food sources (e.g. DOM, bacteria, phytoplankton, zooplankton) in the plankton are called planktotrophic. In general, planktotrophic development is widespread among both spiralian and oligomeric invertebrate phyla belonging to the Spiralia and Oligomera. This is considered to be the primitive form of larval nutrition in most phyla (Jägersten 1972), although this view has been challenged recently (Haszprunar et al. 1995, Rouse 1999). Polychaete trochophores are ciliated as larvae in most invertebrate species, the cilia serving for locomotion and, in species with feeding larvae, for food collection at the same time. They vary widely in the presence, size, and structure of the prototrochal and metatrochal ciliated bands, and other peroral cilia, and they may vary widely in their respective feeding mechanisms (Rouse 2000a, b). Some trochophore larvae capture particles between opposed ciliary bands by the action of both the compound prototrochal cilia and the metatrochal cilia. This requires the use of a prototroch to generate a current for locomotion and feeding, and the metatroch creates a beat in the opposite direction (Strathmann 1993). The ciliated food grove, made up of short cilia between the prototroch and metatroch, moves food to the mouth. The metatroch apparently helps to capture particles swept downstream by the prototroch and passes them into the food groove (Strathmann 1993). Others can capture particles when the metatroch is not beating, and some entirely lack a metatroch. There are several larval forms that appear to use opposed-band feeding and cannot be strictly classified as trochophores. They are very much similar to molluscan veligers (cf. Strathmann 1995). The Mitraria larvae of oweniid polychaetes catch particles between opposed bands of cilia, but the prototrochal cilia are simple (Emlet and Strathmann 1994). Trochophores of some families can capture particles much larger than the width of the food groove (Hermans 1978). The consequences of the morphological diversity

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of trochophores for feeding performance may be remarkable, but are largely unknown as yet (Strathmann 1995). Larval survival and growth vary with food quality and quantity (Qian and Chia 1991). Ontogenetic qualitative and quantitative changes in nutritional requirements have to be taken into consideration when evaluating the ecological consequences of feeding. In the Antarctic, polychaete as well as echinoderm and nemertean larvae selectively ingest bacteria and feed throughout the year on a relatively high and constant bacterial population (Pearse et al. 1991). Growth rates of detritus-fed polychaete larvae paralleled those of larvae fed at natural concentrations of phytoplankton (Qian and Chia 1990). In laboratory experiments, Gosselin and Qian (1997) showed that H. elegans larvae provided with bacteria as their sole particulate food source completed larval development, attached, and metamorphosed into healthy early juveniles (Fig. 7.4). In addition, several of these juveniles, when provided with phytoplankton after metamorphosis, developed to maturity and spawned viable offspring. Bacterial abundances measured in the Port Shelter, a bay in Hong Kong where the organisms for that study were collected, would not sustain maximum development rates of H. elegans larvae. However, bacterial abundances were sufficiently high and consistent over time to constitute a reliable food source. Consequently, starvation may not be a direct cause of larval mortality in H. elegans. The recruitment of this polychaete occurs throughout the year in Hong Kong waters, suggesting that spawning and successful larval development may be independent of phytoplankton availability, and that larvae largely rely on alternate food sources such as bacteria (Gosselin and Qian 1997). Maternally-derived nutrition. Those offspring relying on maternal sources of nutrition can obtain nourishment by: - lecithotrophy, which is the most common utilization of reserves (usually yolk) placed in the egg during oogenesis, - adelphophagy, which is feeding on eggs or siblings within the brood, or - translocation of nutrients directly from the parent during development. Many annelid taxa show lecithotrophy. Lecithotrophic development may take place in the water column, on the sea floor, or, in association with a parent. We consider direct development, in the absence of a larval stage or extraembryonic nutrition from the parent, to be a form of lecithotrophy. Some representatives with lecithotrophic development are present in most polychaete families. Lecithotrophy has been argued to be much less common in shallow-water environments of the tropics and subtropics (Thorson 1950). In such shallow-water habitats, the larval stages typically develop functional feeding structures and feed upon other members of the plankton. Advantages of planktotrophy may include a minimal investment in eggs for the parents. This in turn allows for a larger number of eggs and offspring from any given amount of stored yolk. However, many groups

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Fig. 7.4 A. Survivorship of larvae of Hydroides elegans. Expt. 2. B. Cumulative percentage of larvae in Expt. 2 that attached to the surface of the culture trays. On day 9, the remaining larvae were tested for competence with a 10-4 M solution of IBMX; values for day 10 are also cumulative, and include individuals attaching in the IBMX assay and those that had previously attached to the culture trays. From Gosselin, L.A. and Qian, P.Y. 1997. Marine Ecology Progress Series 161: 93-101, Fig. 5.

have apparently switched from planktotrophic to lecithotrophic development in the course of their evolution and vice-versa (see Rouse 1999, 2000a). The selective pressures responsible for such shifts in developmental mode are difficult to be traced convincingly, but lecithotrophic larvae are certainly independent of seasonal and spatial variations in phytoplankton concentration and also may benefit from more rapid development to the metamorphic stage (see Dahms and Qian 2004). In any event — once feeding larvae are lost from the life history of any particular species, they typically appear to be lost from that lineage forever in that particular form (Rouse 1999). This understanding helps to explain why there are so many structural and functional realizations of feeding larvae among marine invertebrates in general (Strathmann 1993) and the polychaetes in particular.

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Adelphophagy. Later in development, once feeding organs have formed, the larvae of some taxa ingest nearby eggs or fellow larvae prior to hatching or release from brood. This phenomenon, called adelphophagy, has been reported from several polychaete groups (Blake 1969, Simon 1967, Zajac 1984, Petch 1989). Nurse eggs are commonly produced by spionid polychaetes and serve as a source of extra-embryonic nutrition for developing young. Although the influence of nurse eggs on offspring development has been well documented, the origin of nurse eggs is not understood. Nurse eggs in the spionid Boccardia proboscidea appear to arise as do viable oocytes (Smith and Gibson 1999). After spawning, nurse eggs produce fertilization envelopes in this species indicating that development has been activated. Nurse eggs are also capable of producing polar bodies, which indicates completion of meiosis in those eggs. After activation, nurse egg cytoplasm becomes compartmentalized into small bodies that are ingested by developing larvae and nuclear DNA is lost. Hence, nurse egg production in B. proboscidea is an active developmental process and not simply an artifact of sperm limitation (Gibson 1997, Gibson et al. 1999). Osmotrophy. Our present understanding suggests that osmotrophy supplements nutrients obtained by other feeding modes. While there are no known cases where osmotrophy is the only food source for larvae developing from microlecithal eggs, it is possible that for some species utilization of DOM may be required as an additional food source to complete development. Some planktotrophic and/or lecithotrophic larvae may also take up dissolved organic matter (DOM), directly from seawater which represents a nutritional mode termed osmotrophy (Manahan 1990, Manahan and Wright 1991). There has been no convincing evidence yet that polychaete larvae use DOM as a nutritional source. Furthermore, it is unclear so far if DOM was taken up directly into the larval body or was taken up by bacteria that were then ingested by the larvae in their experiment because it is almost certain that bacteria would be present in their experimental setup. Certainly, there is a need to assess the role of bacteria in testing vessels when one intends to examine the DOM intake by larvae because bacteria associated with larvae alone can inoculate experimental vessels in short periods of time. Mixed development. Several nutritional modes may be adopted by a single species. The term “mixotrophy” has been allocated to species that are both autotrophic and heterotrophic. Several nutritional modes may occur either sequentially in a single individual or population, or simultaneously in different individuals, or in different populations (Levin et al. 1991, Qian and Chia 1993). Larval development with an early benthic phase and lecithotrophy, followed by a planktotrophic phase has been termed “mixed development” (Pechenik 1979). According to this author, mixed development is most common in polychaetes (as well as gastropods, nemerteans, and turbellarians). Mixed development combines the

' Reproductive Biology and Phylogeny of Annelida advantages of parental (or brood) protection with enhanced dispersal and habitat selection. Larval development in the spionid Polydora cornuta was found to include both planktotrophic and adelphophagic trophic modes (MacKay and Gibson 1999). Planktotrophic females produce broods in which all eggs develop and offspring hatch as small, 3-setiger larvae. Adelphophagic females produce broods in which 95% of the eggs are non-developing nurse eggs, which are ingested by the developing offspring. Some females switched between adelphophagy and planktotrophy either between successive broods or by delivering both types of capsules in a single brood. Nurse egg ingestion leads to the release of larvae of a wide range of sizes at hatching and a corresponding decrease in the duration of the planktonic phase. Adelphophagic females show differences in several life-history traits linked with reproductive effort (capsule volume, egg volume, number of nurse eggs, offspring size) and fecundity (number of larvae). Only brood size was positively influenced by female size. The lecithotrophic larvae of benthic adults may also develop adult structures in the plankton and commence feeding on plankton prior to settlement. This has been observed in several terebellid polychaete species (Levin and Greenblatt 1981). Shortages in trophic supplies may occur frequently under natural conditions. These led to several experimental approaches studying the limitations of polychaete larval growth to food limitations. Larvae of the polychaete Hydroides elegans were exposed to different feeding schedules to investigate the effects of starvation on metamorphosis and initial juvenile volume in a study by McEdward and Qian (2000). The timing of starvation and the total number of days of feeding had significant effects on the percentage of larvae that successfully completed metamorphosis and on the volume of the post-metamorphic juveniles. However, the duration of starvation did not influence metamorphosis or juvenile volume. Discontinuous feeding reduced the proportion of larvae undergoing metamorphosis and juvenile volume compared to larvae continuously fed for only 5 days, but had no effect on larvae that fed for 8 days. The larvae of H. elegans are remarkably tolerant of starvation but seem to require approximately 5 days of continuous feeding to achieve high levels of metamorphic success and large initial juvenile volume.

7.2.3 Site of Development The marine habitat itself influences several aspects of polychaete larval biology and ecology including sources of mortality, diet, morphological adaptations for feeding and locomotion, and behavior. Particular attention has been given to whether larvae develop in the water column or on/in soft- or hard-bottom substrata. To start with, there is a difference between development that takes place in the water, and benthic development which takes place on or within the seafloor. Among non-benthic larvae, we distinguish demersal larvae

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that develop in waters close to the seafloor from planktonic larvae occupying the remaining water column. Some polychaete larvae swim up towards the surface after being released from the egg-capsules and show a positive phototactic response. Then they quickly move into near-bottom waters and search for a suitable habitat (see Qian 1999). Whereas a demersal life habit may be most common in the deep sea, among encrusting species, or in patchy habitats, most larvae are planktonic and either planktotrophic or lecithotrophic. Benthic development. Polychaete benthic development may occur independent of the parent, attached to sediments, developing freely, or in direct association with a parent. A common form of development without parental care is the encapsulation of embryos for all or parts of the developmental period. Polychaetes have developed a vast array of mechanisms for containing and protecting embryos separate from the parent. These involve capsules that are commonly attached to hard or soft substrates, or tubes and burrows. Aparental encapsulation is welldeveloped in spionid polychaetes and arenicolids (Rouse 1999). Different modes of encapsulation offer protection from predators and physical hazards but are subjected to physiological constraints and can be energetically costly for the parents to produce (Pechenik 1979). Development in association with a parent can involve internal or external brooding. Brooding requires the presence of the parent for successful development, and is distinguished from encapsulated development. Polychaetes may brood larvae dorsally, laterally or ventrally, directly attached to the epidermis, in external pouches, beneath branchiae, or within modified setal structures such as opercular chambers (e.g. Spirorbinae and other Serpulidae) or elytral scales (Polynoidae, Sigalionidae) (Schroeder and Hermans 1975). Brooding is reported in males of some species, externally in the male tube of Hediste (syn. Nereis) acuminata (Weinberg et al. 1990). Pelagic development. Planktonic polychaete larvae spend different periods of time in the plankton before they become competent to settle and metamorphose. Recruitment to benthic habitats then depends on the availability of competent larvae that is affected by abiotic parameters as the hydrodynamics (water flow and flow velocity) (cf. Butman 1987), and biotic parameters such as the presence and densities of adult populations (Davis et al. 1989), adult reproductive cycles, output (Roughgarden et al. 1988), and larval mortality (Rumrill 1990, Dahms et al. 2004b).

7.2.4 Developmental Variation at Generic and Species Level A full range of developmental types are present among the Polychaeta, including pure broadcast spawners showing lecithotrophic and planktotrophic development, brooding in capsules and viviparity (i.e. the release of free living juveniles). There are some invertebrate genera where species exhibit contrasting patterns of larval development. Some of the most striking examples of variation among congeners are seen in

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polychaetes (Blake 1969, Grassle and Grassle 1976). Species in the spionid polychaete Streblospio provide a good example of the range of developmental modes possible in a body brooder. These include: - laying of small (60-70 µm) eggs within brood pouches (Streblospio benedicti) or beneath dorsal branchiae (Levin and Bridges 1995) (Streblospio spp.) and release of long-lived, 3-setiger planktotrophic larvae by both forms (Levin 1984, Rice 1991) - development from larger eggs (120 µm) within brood pouches and release of short-lived, lecithotrophic, 9-setiger larvae (S. benedicti) - direct development from large (200 µm) eggs atop intersegmental, dorsal grooves and release of 14-setiger, crawl-away juveniles (S. shrubsolii — Cazaux 1985). Despite the high incidence of congeners with different development modes, the occurrence of multiple larval development modes exhibited by offspring of the same or different individuals within a single species is extremely rare. Most early reports of this phenomenon have proved to involve cryptic or sibling species. Production of both planktotrophic and lecithotrophic larvae by members of the same species has been confirmed in spionid polychaetes (Blake and Kudenov 1981, Levin 1984, Petch 1989). It may occur in capitellid polychaetes as well (Qian and Chia 1992). Among spionid polychaetes, poecilogony can result from adelphophagy or from eggs of varying size. In several species of the Polydora-Boccardia complex, larvae may emerge from capsules within a worm tube as 3-setiger, planktotrophic larvae, or as juveniles, well fed on nurse eggs and/or siblings (Blake and Kudenov 1981, Zajac 1984, Petch 1989). Individual females of the spionid Streblospio benedicti brood either 70 µm eggs that develop as planktotrophic larvae spending several weeks in the plankton or 120 to 200 µm eggs that develop as lecithotrophic larvae with an abbreviated planktonic period (Levin 1984). The best documented poecilogonous species (one species produce more than one type of larval offspring, often differing in terms of how they feed (or whether they feed) and habitat (planktonic vs. benthic)) is the spionid Boccardia proboscidea (Chia et al. 1996, Gibson 1997, Gibson et al. 1999). This species produces both planktotrophic and adelphophagic larvae, which feed on phytoplankton and yolky eggs, respectively. Egg size, egg number, and duration of the planktonic period showed high heritability in a population of S. benedicti in North Carolina (Levin et al. 1991). Sometimes, the relative period of time polychaete larvae spend in a brood structure versus in the plankton is variable, without a change in the nutritional mode (Schroeder and Hermans 1975).

7.2.5 Consequences of Developmental Modes The consequences of different developmental modes are almost certainly far reaching, but remain poorly understood. Since the publication of Thorson’s (1950) often-cited synthesis, the idea that population-level properties, such as recruitment fluctuations and stability of benthic

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population size are dependent on development mode and fecundity, has achieved the status of a paradigm. Jägersten (1972) in his version of the Gastraea theory argued that the occurrence of free-spawning small eggs, external fertilization and subsequent indirect, planktotrophic development represents a uniformly plesiomorphic condition. Although his Bilaterogastraea theory has been widely ignored (see Rouse 1999), Jägersten’s scenario about life history evolution still represents the paradigm for the evolution of marine invertebrate reproduction in general (Wray 1995). This has been criticized as well (e.g. Haszprunar et al. 1995), but the combination of the scenarios outlined above for a hypothetical ancestral reproductive mode and the hypothesis proposed by Strathmann (1993) that feeding larvae tend to be lost rather than gained, may have led to the continued acceptance of Jägersten’s assumptions. However, there have been few reliable tests of these ideas (cf. Levin and Bridges 1995). Levin and Huggett (1990) assembled the published data on population variability in marine benthic invertebrates to assess whether planktotrophy and lecithotrophy conferred different levels of variation. However, the analyses of several sets of data revealed no evidence that a particular developmental mode confers greater or lesser population stability for the poecilogonous polychaete Streblospio benedicti when comparing its planktotrophic and lecithotrophic populations (Levin and Huggett 1990).

7.2.6 Settlement and Metamorphosis 7.2.6.1 Introduction A critical stage in the larval life occurs at competence, a physiological state wherein dispersing larvae are able to attach and subsequently metamorphose into sedentary juvenile stage (Chia and Rice 1978). The terms of “settlement and metamorphosis” describe separate phenomena. Settlement is defined as the behavioral performance when pelagic larvae descend from the plankton to the benthos, and move upon the substratum with or without attaching to it. Settlement is reversible: a larva can swim up again from the substrate to resettle at another location. Metamorphosis is basically a morphogenetic event. For all larvae, metamorphosis includes the loss of larva-specific organs and the development of juvenile or adultspecific organs (see Hydroides elegans as a good example in Fig. 7.5). For sessile marine invertebrates, a firm and often permanent attachment may represent the first stage of metamorphosis. 7.2.6.2 Settlement Rodriguez et al. (1993) define settlement as a process with two phases. The first is a behavioral searching phase, and the second a phase of permanent residence of attachment to a substrate, which triggers metamorphosis and where morphogenetic events take place. However, metamorphosis can commence even before larval settlement or right after larval attachment to the substratum (Fenaux and Pedrotti 1988). The two different processes,

'" Reproductive Biology and Phylogeny of Annelida

Fig. 7.5 Postembryonic development of Hydroides elegans. A. early trochophore larva. B. competent larva. C. metamorphosing larva (0.5 h after the onset of attachment). D. metamorphosing larva (6 h after the onset of attachment). E. metamorphosing larva (9 h after the onset of attachment). F. young juvenile (24 h after the onset of attachment) (photoshop file of Professor Qian).

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larval settlement and larval metamorphosis in marine invertebrates may require different environmental cues (see below and Chia 1989). Larvae of some invertebrates settle and metamorphose in response to a specific cue from a unique source (Burke 1983, Rodriguez et al. 1993). Others respond to cues originating from several sources (Pawlik 1992 — and see below). Settlement. Settlement is the process by which a planktonic larva or juvenile moves to the substratum and locates a site at which to begin its benthic life. Polychaete larvae may be able to exert some control over their vertical distribution by constant swimming or by adjusting sinking rates (for instance by erecting larval chaetae). Once on the bottom, larvae of many species can actively accept or reject particular settlement sites on the basis of a variety of chemical and physical cues (Qian 1999). However, few natural settlement cues (positive or negative) have been completely isolated and identified, and data on signal transduction pathways involved in settlement behavior are still limited (cf. Hadfield and Paul 2001). At this place we will not consider the increasing evidence on artificial inducers for metamorphosis. These are possibly important for the understanding of developmental aspects and internal metamorphic signaling systems (Dahms et al. 2004b). However, artificial inducers do not contribute much to our understanding of the chemical ecology of invertebrate recruitment in their natural environment. Settlement in soft bottoms. Local current patterns, flow velocity, and, particularly, near-bottom flow dynamics play very important roles in the retention and settlement of invertebrate larvae in general (Hannan 1981). Polychaete larval settlement appears to be a very dynamic process. As said, larvae can reject one site and select another for settlement. Such active site selection is often mediated by environmental cues (e.g. Woodin 1986, Pawlik 1992 — for reviews). Larvae of many marine polychaetes require specific cues to settle and metamorphose. The role of organic constituents of sediments in the specific recognition of appropriate habitats for settlement by polychaete larvae has early been recognized (Butman 1987, Butman and Grassle 1992, Butman et al. 1988a, b, Grassle and Butman 1989). Settlement by infaunal polychaete larvae and other infaunal macro-invertebrates is dependent on such organic material (Snelgrove et al. 1999). Cohen and Pechenik (1999) concluded from settlement experiments with larvae of Capitella sp. I that the cue is probably organic and bound to fine particles. This allows for the possibility that living bacteria are important for the fact that much of the organic enrichment of marine soft bottoms is undoubtedly due to bacteria. Biggers and Laufer (1992, 1996) studying the same species, found that metamorphosis was stimulated by insect juvenile hormones and related compounds. Also, chemicals with a similar activity as juvenile hormones could be detected in marine sediments that stimulated settlement. Gregarious settlement. As a consequence of larvae choosing to settle in response to the presence of adults, juveniles, or recent recruits of the same species — have been reported for several phyla (Knight-Jones 1951), but in

'$ Reproductive Biology and Phylogeny of Annelida particular for tube-dwelling polychaetes (Burke 1986). Sabellariid and serpulid polychaetes often settle gregariously and form colonies, sometimes producing large mounds, which are characteristic for these tube-living worms. Larvae of these species are known to settle on the tubes made of mucus-cemented sand-grains of adult conspecifics (Toonen and Pawlik 1994, 2001). Studies on Phragmatopoma lapidosa californica, Sabellaria alveolata, Spirobranchus polycerus, Hydroides dianthus, and H. ezoensis have found adult-associated compounds to be responsible for larval settlement leading to adult aggregations (Marsden 1991, Pawlik 1992). All these representatives are sessile after metamorphosis, and in some species factors released from adult organisms have been hypothesized as stimuli for larval settlement. However, not all aggregations of sessile marine invertebrates are mediated through intraspecific chemical cues. Asexual propagations for example can produce such masses in the serpulid Salmacina spp. Other dense aggregations may arise through a settlement response to an abundant benthic cue as in another serpulid, Hydroides elegans (Beckmann et al. 1999). 7.2.6.3 Metamorphosis Anatomical changes. Metamorphosis, when segmented larvae acquire adult structures and lose purely larval structures, is a striking change in most polychaetes. In most polychaetes, cellular larval structures like the prototroch are histolyzed and presumably resorbed. In a few larvae that have a more striking metamorphosis (e.g. Polygordiidae, Oweniidae) these structures are shed and may then be ingested by the new juvenile (Wilson 1932). Larval chaetae are simply shed. Metamorphosis may occur before or after larvae have settled to the bottom. If it occurs in the plankton, and if the juvenile remains there for some time, the resulting stage may be called a secondary larva. For example, the segmented larvae of some terebellids undergo metamorphosis in the plankton, where they remain for days to weeks, living in mucus tubes and feeding with the tentacles of the adult architecture (Bhaud et al. 1990). Segmented larvae of many polynoids also metamorphose in the plankton and remain there as juveniles for days to weeks. Eventually, these secondary larvae settle and start the benthic existence of juveniles. Habitat transition. Metamorphosis may result in the immediate termination of a planktonic existence, but more often the elongated, metamorphosing larvae remain planktonic for varying lengths of time. The metamorphosing stages of spionids, sabellariids, and oweniids even possess greatly enlarged erectile anterior setae that serve as flotation or protective devices (Bhaud and Cazaux 1990). Delayed metamorphosis. The degree to which the dispersing individual selects an appropriate habitat for the sedentary phase of the life cycle obviously determines the probability that a given individual will indeed survive to reproductive maturity (Pechenik 1990). Among marine invertebrates with free-living larvae, selective pressures acting against random metamorphosis to the adult form and habitat must have been substantial. The dispersing larvae of many species are highly selective

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about where they will metamorphose and whether they can delay their metamorphosis for varying periods of time (up to many months for some polychaete species — cf. Qian and Pechenik 1998) if they fail to encounter the appropriate cues. The cues that subsequently trigger metamorphosis are often associated with some components of the adult environment: the adult food source or prey species, for example, or quite commonly, adults of the same species. Conditions and extend to which larvae actually delay their metamorphosis in the field and the conditions under which they delay their metamorphosis are difficult to explore. However, it became clear that there are unanticipated costs associated with the delay of metamorphosis in terms of postmetamorphic survival or growth. In these cases, the adaptive benefits of prolonging the competent period of larval development may not be realized in practice (Bhaud et al. 1990).

7.2.7 Cues for Settlement and Metamorphosis 7.2.7.1 Introduction It still remains mostly unexplored whether one or more cues are involved in the transition from a planktonic larval form to a benthic life. For a number of species, larvae are brought to their prospective settling substrate by ontogenetic changes in their tropistic behavior; becoming either positively geotactic or negatively phototactic, which brings polychaete larvae towards the sea-bottom (Thorson 1946). In most cases, metamorphosis occurs after larvae are already close to appropriate settlement substrata and thus able to detect surface cues, which may be physical or chemical, or both (Chia and Rice 1978, Marsden et al. 1990). The interaction between competent larvae and the substratum are the prime determinants of settlement at small spatial scales as well as for postsettlement mortality. This interplay can be affected by physical, chemical or biological parameters. The latter may comprise the natural communities in general, and naturally inducing or inhibiting chemical compounds released by conspecific individuals, sympatric species, prey species, or biofilms (Qian 1999). Prevailing among invertebrate larvae are examples where a single inducer is known to affect the entire recruitment process. Larval settlement in an individual species can be controlled by a single cue or a mixture of cues. Examples include prosobranchs (McGee and Targett 1989), nudibranchs (Lambert et al. 1997), echinoids (Pearce and Scheibling 1991), and ascidians (Young and Braithwaite 1980). Other invertebrate larvae demand separate cues for settlement and metamorphosis. Barnacles (Clare and Matsumura 2000) and oysters (Tamburri et al. 1992) are good examples. For marine polychaetes, cues can originate from conspecific individuals or sympatric organisms, microbial films, food items, or the habitat. Although a variety of chemicals, including proteins, free fatty acids, polysaccharides, neurotransmitters, and inorganic ions, have been

'& Reproductive Biology and Phylogeny of Annelida suggested in inducing larval settlement and/or metamorphosis, only a few cues operating in situ have been isolated and structurally identified (reviewed by Qian 1999). Hydroides elegans is one of the most investigated biofouling species in terms of larval response to bacterially derived settlement cues. It is a tubebuilding polychaete worm that occurs widely in tropical and sub-tropical waters. Its congeneric species such as H. dianthus, H. ezoensis and H. norvegia occur in colder waters, but are less well studied (Toonen and Pawlik 1996, Okamoto et al. 1998, Watanabe et al. 1998). 7.2.7.2 External non-chemical cues Water currents and flow dynamics may determine both, vertical and horizontal distribution of larvae in the water column as well as the number of competent larvae transported to settling sites (Banse 1986, Le Tourneux and Bourget 1988, Pawlik and Mense 1994). At large spatial scales, marine polychaete larvae have been shown in the field to move like passive particles (e.g. Bhaud and Cazaux 1990). At small spatial scales, after larvae have descended to the sea bottom, both, near-bottom hydrodynamics and active larval substratum selection determines the success of larval settlement (Walters et al. 1997, Qian 1999). These larvae can actively select settlement sites by horizontal swimming between available habitats effectively only in still water or in a laminar flow (Snelgrove et al. 1993). Pawlik and Butman (1993) conducted settlement experiments with larvae of Phragmatopoma lapidosa in turbulent flume flows (near-surface velocities of 5, 10, 15, 20, 25, 30 and 35 cm s–1) over a hydrodynamically smooth bed. Boundary shear velocities spanned the critical shear velocity for initiation of particle motion and for suspended-load transport of passive larval mimics. Larvae were allowed to pass over a sediment array with two treatments: 1) sand produced by adults that is a natural inducer of metamorphosis and, 2) non-inductive sand. They found that at intermediate flows where numbers of metamorphosed juveniles and the total number of larvae and juveniles in the array were maximal, larvae tumbled along the flume bottom, as did the passive larval mimics. At slower flow velocity, larvae actively left the bottom and swam into the water, passing over the array. At fastest flow velocity, hydrodynamics may have reduced larval settlement, because larvae, like the mimics, were eroded from the bed and carried as suspended load over the array or because enhanced turbulent mixing distributed larvae more evenly in the water, thus reducing their concentration close to the sea bed. Once reaching the substratum, behavioural responses to chemical cues were found to be ultimately responsible for metamorphosis (Chia and Rice 1978). Qian et al. (1999) found that velocity and Reynolds number have very strong effects on larval settlement patterns and settling rates of Hydroides elegans in pipe systems. PVC tubes, on which biofilm had been allowed to develop for 48 hours, were used to studying attachment of H. elegans. In all but low flows, the flow velocity and Reynolds number were poorly correlated with

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patterns of larval settlement (Fig. 7.6). A hydrodynamic measure, which is theoretically independent of the size of the tube, the so called ‘velocity gradient’, was well correlated with the highest settlement of H. elegans. Comparisons of results from field and laboratory experiments reveal slight differences, however. Settlement of H. elegans was offset to higher shear values in the tubes of smaller diameters (Fig. 7.7). H. elegans larvae had the narrowest range of settlement in relation to flow as settlement of this species was the highest from 8 to 20 s–1 and hardly occurred above 200 s-1 (Qian et al. 2000). Factors that apparently affect larval settlement of marine polychaetes include surface roughness, thickness of the deposit on the substratum, wet angles, type of substratum, degree of tidal exposure, tolerance of wave energy, phototaxis, and availability of settlement sites (Woodin et al. 1993). Dean (1981), for example, reported for the serpulid polychaete Hydroides dianthus that it settles more pronounced on bare (rough) surfaces than on mimic panels (smooth surfaces). Larvae of the terebellid Eupolymnia nebulosa settled more pronounced in the presence of a stable support where sediment elements are fine enough to be manipulated and used for tubebuilding. With limitation of space for settlement and growth, adults of the spionid Pseudopolydora paucibranchia can inhibit the larval settlement of Armendia sp. (Tamaki 1985). Adults of Hydroides elegans are eliminating any settling larvae within the range of their tentacle length (Bryan et al. 1997). 7.2.7.3 External chemical cues Conspecific cues. Representatives of the Sabellariidae build tubes of sand grains they cement. Gregarious sabellariids often live together in extensive colonies of these tubes by recruiting larvae from the plankton. Evidence that a chemical cue in the tube cement triggers larval settlement was first reported for British populations of Sabellaria alveolata by Wilson (1968, 1974). Larvae of Phragmatopoma lapidosa californica, a sand tube-building worm from California, settle and metamorphose preferentially on anterior portions of sand tubes of conspecific adults (Jensen and Morse 1984). Glass beads that conspecifics had previously been used to build tubes were also highly inductive. The stimulus for metamorphosis was inactivated by boiling. However, the chemical nature of the settlement cue from adult tubes that should stimulate settlement and metamorphosis in this species has been controversial (Jensen and Morse 1984). Marine polychaete worms often settle gregariously and form colonies, sometimes producing large mounds. Larvae of these species are known to settle on the tubes of adult conspecifics. Phragmatopoma californica, commonly found along the coast of California, forms reefs of amassed sand tubes, and its larvae are induced to settle and metamorphose by the organic extract of natural sand tubes (Pawlik 1986). An active fraction of the extract contained a mixture of fatty acids ranging from C14 to C22, in which eicosapentaenoid (20:5), palmitic (16:0), and palmitoleic acids (16:1) were predominant. In fact, palmitoleic, linoleic (18:2), arachidonic (20:4), and eicosapentaenoic acids induced larval settlement (Pawlik and Faulkner

 Reproductive Biology and Phylogeny of Annelida

Fig. 7.6 Settlement of Hydroides elegans in 5 and 10 mm diameter tubes in the laboratory, with laboratory-reared larvae. Mean percentage settlement ±SD is plotted against mean flow velocity (upper panel), Reynolds number (middle panel), and velocity gradient (lower panel). From Qian, P.Y., Rittschoff, D., Sreedhar, B., and Chia, F.S. 1999. Marine Ecology Progress Series 191: 141-151, Fig. 5.

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Fig. 7.7 Settlement of Hydroides elegans in 7 types of 10 mm diameter tubes: A. in the field. B. In the laboratory. Mean percentage of settlement or mean number of larvae settled ±SD is plotted against mean flow velocity and mean velocity gradient. The velocity gradient was calculated according to Crisp (1955) and Qian et al. (1999), and was rounded up to the nearest integrate number. From Qian, P.Y., Rittschoff, D., and Sreedhar, B. 2000. Marine Ecology Progress Series 207: 109-121, Fig. 2.



Reproductive Biology and Phylogeny of Annelida

1986). However, some controversial reports have been published (Jensen et al. 1990). These fatty acids are likely to act as “juvenile hormone (JH)-active compounds” as suggested by Biggers and Laufer (1992). Methyl farnesoate is an endogeneous JH-active compound in the annelid Capitella sp. 1. Larvae of the Japanese serpulid Hydroides ezoensis were also reported to settle and metamorphose in response to the tube extract, from which a new monoacylglycerol, 1-(4Z, 7Z, 10Z, 13Z, 16Z, 19Z)-docosahexaenoyl glycerol has been obtained as an active component (Watanabe et al. 1998); the synthetic sample induced larval metamorphosis at 3 × 10–6 to 3 × 10–5 M. However, high percentages of larval mortality were observed at concentrations higher than 3 × 10–5 M. Again, these compounds are likely to mimic JH-active compounds. Conspecific associated compounds were tested in a study by Bryan et al. (1998), where conspecific adult homogenates induced 39 and 82% of Hydroides elegans larvae to settle and metamorphose within a period of 2 and 4 days, respectively (Fig. 7.8). Homogenates of the adult tube alone did not induce settlement, indicating that the inducer originates from the worm. Extraction and assays on crushed adult homogenates revealed that the inductive compounds from adults are smaller than 10000 Daltons. Cues from biofilms. Biofilms fundamentally affect settlement for many invertebrate larvae (Wieczorek and Todd 1998) and the prevalence of biofilms on marine substrate surfaces may provide habitat signals for settling marine organisms in general (Johnson et al. 1997) and for polychaete settling stages in particular (Beckmann et al. 1999). These biofilms usually originate as adsorbed organic molecules, which may serve as a stimulus for the attachment of bacteria, benthic diatoms, fungi, and

Fig. 7.8 Percent larval metamorphosis of Hydroides elegans in response to conspecific worm homogenates assayed at 3 concentrations after 2 and 4 days incubation. Control is filtered seawater (FSW). Data plotted are means ±SD of 5 replicates. From Bryan, J.P., Qian, P.Y., Kreider J.L., and Chia F.S. 1997. Marine Ecology Progress Series 146: 81-90, Fig. 4.

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unicellular eukaryotes (Baier 1983). Marine bioorganic films (natural biofilms) are heterogeneous in composition and highly variable over time. Most research has focused on the role of bacteria in biofilms in enhancing the acceptability of surfaces for settlement of invertebrate larvae, and extensive literature exists on this topic and hard bottom settlers (cf. Holmstrøm and Kjelleberg 2000). Larvae of Hydroides elegans can settle well in response to biofilms (Fig. 7.9). This effect is not mediated by the polarity of the biofilm surface (Qian 1999, Qian et al. 2000). If a biofilm is not detected, larvae arrest developmentally and remain in the water column up to several months. Hadfield et al. (1994) observed a positive correlation between larval settlement and bacterial density in biofilms, and proposed bacteria as inducers for larval settlement in H. elegans. The Qian group has tested bacterial effects of over 200 bacterial isolates originated from natural biofilms, and epibiotic bacterial communities of sponges and the green alga Ulva reticulata. The effects of bacterial strains on larval settlement of H. elegans can be neutral, inductive or inhibitive (Dobretsov and Qian 2002, Lau et al. 2002, Lee and Qian 2003). Lau and Qian (1997) isolated 12 bacterial strains from natural biofilms and conducted larval settlement bioassays with Hydroides elegans. The results indicated that different bacterial strains caused different levels of

Fig. 7.9 Metamorphosis of Hydroides elegans larvae in response to bacterial density of biofilm. Histogram indicates the increase in bacterial density of biofilm over the biofilm age and line drawing indicates the percentages of metamorphosis of larvae. Data plotted are the mean ±SD of 6 replicates. From Qian, P.Y. 1999. Hydrobiologia 402: 239-253, Fig. 3.

" Reproductive Biology and Phylogeny of Annelida settlement in this species. An enumeration of bacterial density on the test substrata revealed that responses of larvae were not due to differences in bacterial density. Bacterial strains that consistently induced settlement were used for multispecies biofilm assays, revealing that both, qualitative and quantitative changes in species composition affected larval settlement. Changes in either bacterial species or the ratio of bacterial species in multispecies bacterial films also affected H. elegans larval settlement (Fig. 7.10). When two bacteria with similar effects were combined, the resulting biofilms were not different from the corresponding monospecies films. When two bacteria with different effects were mixed, the effects were antagonistic and dependent on the bacterial species and their ratios when mixed. An integration of results from the antibiotic assays together with those from the assays of larval settlement on bacterial films revealed that phlorotannins and related compounds were inhibitory to some of the bacterial species that induced high levels of H. elegans larval settlement. However, some of the bacteria that induced low levels of H. elegans larval settlement were resistant to these compounds. Bacterial effects do not show any taxon-specific pattern — larval settlement can be induced by bacterial isolates having distant phylogenetic relationships, while closely related isolates may evoke different larval responses. For instance, inductive isolates are widely distributed in the four major phylogenetic branches of the bacteria, namely Cytophaga-FlexibacterBacteroides, Gram-positive, alpha-Proteobacteria and gammaProteobacteria. Moreover, all possible effects on larval settlement can be observed in isolates of some genera such as Alteromonas and Bacillus. A similar study by Unabia and Hadfield (1999) reported that larval settlement did not correlate with the biochemical characteristics (e.g. substrate utilization and enzymatic capabilities) of the bacteria under investigation, without that they have been identified, however. 7.2.7.4 Cues from bacteria Johnson et al. (1997) hypothesized that the induction of invertebrate settlement and metamorphosis by bacteria may be far more widespread than expected. According to these authors ubiquitous settlers may respond to a number of bacterial species in biofilms which are common to many substrata. Specialized settlers, on the other hand, settle in response to bacteria that are specifically adapted to those surfaces, rather than to metabolites of living substrata themselves. Further laboratory experiments of the Qian group (e.g. Lau et al. 2003) showed that: - inductive bacterial strains were more active in stationary phase than in log phase. - settlement inducers were associated with cell surfaces as the culture broth of bacteria was inactive. - dead bacteria or biofilms treated with antibiotics were not active. Using the bacterium Roseobacter littoralis as a model strain, Lau et al. (2003) investigated the effect of bacterial extracellular polymers

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Fig. 7.10 Metamorphosis of Hydroides elegans in response to multispecies bacterial films after 48 h of incubation. The numbers on the x-axis correspond to the identities of the bacteria. Monospecies bacterial films were used as controls. Multispecies bacterial films were developed from mixing suspensions of 2 bacterial species in a 1:1 ratio. Data that are significantly different are indicated by different letters above the bars. From Lau, S.C.K. and Qian, P.Y. 1997. Marine Ecology Progress Series 159: 219-227, Fig. 3.

$ Reproductive Biology and Phylogeny of Annelida (exopolymers) on the larval settlement of H. elegans. Bioassays with exopolymer fractions dissociated from bacterial films evoked the initial stages of the larval settlement process, (i.e. larvae slowed down, secreted a mucous thread and crawled over the surface). This response is typical of larvae that encounter an attractive bacterial film. In contrast, bioassays with exopolymers in association with UV-irradiated, metabolically inactive bacterial films evoked complete settlement. However, the percentage of responding larvae was negatively correlated with the magnitude of UVdosage (Fig. 7.11). Since UV energy crosslink both intra- and extracellular pertinacious components, it could not be distinguished whether the decrease in larval settlement was due to a modification of pertinacious components of exopolymers or due to the elimination of cellular activity. Nevertheless, the results ascribe bacterial exopolymers the role of an indicator of substratum suitability and provide evidence that the polysaccharide moiety of exopolymers does not complement this effect. These laboratory results were consistent with field experiments conducted by Keough and Raimondi (1995) where larvae of 19 sessile invertebrate taxa were exposed to a variety of chemical cues including bioorganic films developed in the laboratory and kept under field conditions for 7 days. These experiments show that under field conditions, polychaete larvae settle at very different rates onto surfaces differing only in the composition of the microbial communities. Recent investigations on the nature of bacteria-derived inductive cues for larval settlement in H. elegans suggested bacterial metabolites as a cue and bacterial exopolymers as a facilitator (Harder et al. 2002a). The role of bacterial metabolites as settlement cues was first suggested by using the highly inductive bacteria Roseobacter sp. and Rhodovulum sp. as model strains, which showed a strong correlation between larval settlement and metabolic activity of bacterial films (Lau and Qian 2001). By using a comprehensive bioassay-guided isolation and gas-chromatographic analysis of bacterial metabolites, Harder et al. (2002b) isolated a mixture of waterborne bacterial metabolites having strong inductive effects on larval settlement of H. elegans. These metabolites were dominated by compounds of low to medium polarity such as alkalized alkanes, alkenes, and alkylated hydroxyketones in the volatility range facilitated by n-heptane and neicosan (Fig. 7.12). There is strong good evidence that the inductive metabolites require bacterial exopolymers as a vehicle to convey their effect to larvae of H. elegans, otherwise remain inactive (Harder et al. 2002b). In bioassays of this investigation, the metabolites only elicit larval settlement when adsorbed to charcoal, suggesting that adsorption to a suitable substratum is essential for their activity or sufficient concentration is required in order to be active (Fig. 7.13). Larvae did not respond to waterborne metabolites when prevented from contacting the bacterial film surface. These results indicate that an association of the chemical signal with a sorbent-like substratum may be an essential cofactor for the expression of biological activity. This

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Fig. 7.11 A. The percentage of larval settlement of Hydroides elegans after 4 h in response to films of Roseobacter litoralis that received different dosages of UV radiation. B. The percentage of respiring cells in films of R. litoralis after UV-irradiation. “Positive control” represents bacterial film without UVirradiation. “Negative control” represents a sterile microscope coverslip. Data are representative of five repeated experiments. The percentage of larval settlement is expressed as mean ±SD of five replicates; the percentage of respiring bacterial cells as mean ±SD of three replicates. Data that are significantly different at α=0.05 according to a Tukey’s test are indicated by different letters above the bars (P<0.0001, one-way ANOVA). From Lau, S.C.K., Harder, T., and Qian, P.-Y. 2003. Biofouling 19(3): 197-204, Fig. 1.

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Fig. 7.12 Gas chromatograms of chloroform extracts of bacterial metabolites of Roseobacter sp., Vibrio sp., Rhodovulum sp., and Micrococcus sp. GC conditions (Hewlett-Packard 5890 Series II): 30 m fused silica capillary column (DB-5), 0.25 mm ID; temperature program: 40°C for 5 min, 40-150°C at 10°C/min, 150°C for 2 min. From Harder, T., Lam, C., and Qian, P.-Y. 2002a. Marine Ecology Progress Series 229: 105-112, Fig. 4.

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Fig. 7.13 Larval settlement in response to charcoal- and glass-adsorbed chloroform extracts of bacterial metabolites. Data are mean percentages ±SD of 5 replicates of larval settlement after 12 h. *Statistically significant difference (Student’s t test, α=0.05). Harder, T., Lau, S. C. K., Dahms, H.-U., and Qian, P. -Y. 2002b. Journal of Chemical Ecology 28(10): 2029-2043, Fig. 3.

concept is supported by the pattern of larval settlement in compartmentized test vessels, in which larvae only settle in the bacterial film-bearing compartment, but not in the unfilmed compartment that can readily receive waterborne metabolites from bacterial films (Harder et al. 2002b). Bacterial exopolymers, which are highly sorptive to bacterial metabolites (Decho 1990), seem to be of essential importance for the inductive effect of bacterial metabolites. Exopolymers are high molecular weight mucous secretions of bacteria, which exist either as capsules that closely surround cells or loose-slime matrices. Exopolymers do not directly evoke settlement response in larvae of H. elegans (Lau et al. 2003), but may retain bacterial metabolites on bacterial cell surfaces and thus render the induction of larval settlement for bacterial metabolites. This hypothesis is in accordance with a similar suggestion on the role of bacterial exopolymers in mediating larval settlement in oysters (Decho and Zimmer-Faust 1996). Again, bacterial exopolymers are suggested to play a key role for this phenomenon (Harder et al. 2002b). Provided that exopolymers mediate larval settlement by binding inductive bacterial metabolites to cell surfaces, the lack of a settlement response of certain bacterial films may be related to the properties of exopolymers in these bacteria. It is well known that the quantity and quality of exopolymers may vary drastically among strains or even within the same strain under different conditions (Sutherland 1985, Decho 1990). Moreover, bacteria lacking exopolymers are also common (Cowen 1992). Hence, the effect of bacteria on larval settlement are likely be affected by the possession, quantity, and quality of exopolymers. Exopolymers are proposed to be information-rich in nature and are direct cues for larval settlement of other invertebrates, mainly due to the vast diversity of exopolymer conformation given by the broad arrangement

 Reproductive Biology and Phylogeny of Annelida of monosaccharide units and the additional non-carbohydrate constituents. Whilst Szewzyk et al. (1991) suggested that exopolymers of Pseudomonas sp. induced settlement of the tunicate Ciona intestinalis by passively entrapping the larvae, Kirchman et al. (1982a) proposed a lectin-binding model for larval settlement of the spirorbid serpulid Janua brasiliensis on films of the Gram-negative bacterium Deleya marina. It was suggested that larval settlement was mediated by the binding of lectin molecules on the surface of larvae to the specific arrangement of sugars in exopolymers. In contrast to Hydroides elegans, this effect is independent of bacterial viability. Railkin and Chikadze (1999) proposed a two-way lectin-carbohydrate model, suggesting that larval settlement of the hydroid Gonothyraea loveni was mediated by lectin molecules present on both the larval surface and microbial film. An interesting case of polytypic cue dependence is reported for Hydroides dianthus (Toonen and Pawlik 1994). In sibling cohorts of this species, 10 to 20% settle in response to common biofilms, while the remaining portion of the population depend on cues from living postsettlement H. dianthus. Toonen and Pawlik (1996) propose that this duality provides, in each cohort, a group of colonizing larvae that settle onto surfaces characterized by biofilms. Once established, the founders serve as a stimulus for recruitment of the larger portion of larvae, leading to the buildup of compact colonies of H. dianthus on submerged surfaces (see gregarious settlement). This polytypic response may provide initial founder communities and subsequent dense aggregating communities in this species. Studies of H. ezoensis in Japan, assumed that dense aggregations of this species in the field must be induced by conspecific adults (Okamoto et al. 1998; Watanabe et al. 1998). Compounds from worm homogenates, however, were not tested for bacterial biofilms. Therefore, it remains to be shown that settlement of H. ezoensis in the field is strictly induced by adult worm metabolites or attracted by the subsequent development of bacterial films. 7.2.7.5 Cues from diatoms Only a few studies have addressed the role of marine benthic diatoms in mediating the settlement response of invertebrate larvae. Until recently, the role of diatoms as potential mediators of larval settlement has largely been investigated with grazing invertebrate species intended for mass aquaculture and human consumption, such as abalones (Slattery 1992, Bryan et al. 1998), sea urchins and sea cucumbers (Itô and Kitamura 1997). A commonality among these organisms is their post-larval dietary requirement for benthic diatoms. A positive correlation of the abundance of diatoms in biofilms on the mediating effect on larval attachment has been demonstrated in these organisms (Slattery 1992). The cues involved in the mediation of the larval settlement response have not been identified but were hypothesized to be algal-derived food odors. As for the Polychaeta, the role of benthic diatoms as non-dietary mediators of marine invertebrate larval attachment and metamorphosis

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have been comprehensively investigated in Hydroides elegans. A variety of diatoms were isolated from natural biofilms and separately investigated in still-water larval settlement assays by (Harder et al. 2002a). The statistical evaluation of test results with 32 unialgal films in comparison to controls of natural biofilm and clean glass substratum categorized the settlement response towards individual diatom species into 3 distinct groups, i.e. the same as towards a natural biofilm, the same as towards clean glass surface, and between these extremes (Fig. 7.14). Out of 32 diatom strains under investigation, 13 fell into the inductive categories. These 13 strains comprised 7 genera, indicating that the inductive effect on larval settlement by diatoms was not genus-specific (Harder et al. 2002a). However, an unambiguous correlation of these results to the presence of diatoms was confounded by the inherent experimental difficulties of performing larval bioassays under axenic conditions, i.e. in the absence of bacteria. Although the abundance of bacteria in diatom bioassays was significantly lower than in natural biofilms, a mere quantitative evaluation of the impact of the bacterial contamination would have neglected the significant qualitative effect of some bacteria on the induction of larval settlement in H. elegans. To decouple the larval response to the bacterial component from that to nonaxenic diatom films, the bacterial communities associated with diatom cultures were separated by filtration to create diatom-free microbial films, representing the respective bacterial contaminants of each culture. Under the assumption that the separated bacteria reflected the community structure in mixed diatom-bacteria films, a comparison of the 2 film types allowed a qualitative evaluation of non-axenic, unialgal films on the induction of larval settlement. Whilst the observed bacterial abundance in diatom-free microbial films and non-axenic diatom films was statistically the same, larval settlement towards diatom films was significantly higher than to microbial control films, indicating that the induction of larval settlement was due to qualitative characteristics of diatoms and not confounded by concomitant bacteria (Lam et al. 2003) (Fig. 7.15). In a follow-up study, Lam et al. (2005a) investigated the origin and nature of diatom-derived larval settlement cues in more detail. A model system included 4 diatoms, two of which being inductive (Achnanthes sp. and Nitzschia constricta), two were non-inductive (Amphora tennerima and Nitzschia frustulum) on larval settlement of H. elegans. It was shown that the activity of the diatom-derived settlement cue was cell surface-associated and independent of the viability of diatom cells (Fig. 7.16). The qualitative differences of the diatom EPS samples were further analyzed by bioassayguided gel-chromatography (Fig. 7.17). The bioactive EPS fraction of Achnanthes sp. mainly consisted of large macromolecules, while the bioactive EPS fraction of N. constricta consisted of both large and small macromolecules. Due to their sensitivity to changes in environmental parameters, diatoms may serve as small scale models of substratum suitability for larval settlement of benthic marine invertebrates, analogous to the reported role of bacteria in marine biofilms.

Fig. 7.14 Mean percentage of larval settlement of Hydroides elegans after 24 h in response to 32 individual monospecific diatom films. A clean glass slip represented the negative control ©; a glass slip filmed with natural biofilm represented the positive control (NBF). Bars denoted with symbols are indicating the assignment to one of the following statistical categories (multiple pairwise comparison of settlement rates on individual diatom-filmed slips with both controls; Dunnett’s test, α=0.05; oneway ANOVA, p<0.001): #, no effect (same percentage of larval settlement as in the negative control); *, inductive (same percentage of larval settlement as in the positive control); @, weakly inductive (percentage of larval settlement significantly higher and lower than in the negative and positive control, respectively). Data plotted are means ±SD of 5 replicates. From Harder, T., Lau, S.C.K., Dahms, H.-U. and Qian, P.Y. 2002b. Journal of Chemical Ecology 28(10): 2029-2043, Fig. 1.

 Reproductive Biology and Phylogeny of Annelida

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Fig. 7.15 A. Mean percentage of larval settlement of Hydroides elegans after 24 h in response to filmed (black) and unfilmed (shaded) surfaces in comparison to controls of marine biofilms (MBF) and filtered seawater (FSW). Significant differences between larval settlement rates on filmed and unfilmed surfaces are asterisked (Student’s t-test, α=0.05). Data plotted are means ±SD of 5 replicates. B. Mean density of diatoms on filmed and unfilmed surfaces. NC: Nitzschia constricta; A: Achnanthes sp. Significant differences between diatom densities are asterisked (Student’s t-test, α=0.05). Data plotted are means ±SD of 5 replicates based on counts of 5 fields of view at 100 × magnification. NA = not applicable. C. Mean density of bacteria on filmed and unfilmed surfaces. Significant differences between bacterial densities are asterisked (Student’s t-test, α=0.05). Data plotted are means ±SD of 3 replicates based on counts of 5 fields of view at 1250 × magnification. From Lam, C., Harder, T. and Qian, P.Y. 2003. Marine Ecology Progress Series 263: 83-92, Fig. 4.

" Reproductive Biology and Phylogeny of Annelida

Fig. 7.16 Mean percentage of larval settlement of Hydroides elegans after 24 h in response to live and dead diatom films in comparison to control of filtered seawater (FSW). Data that are significantly different at α = 0.05 according to a Tukey’s test are indicated by different letters above the bars. Data plotted are mean percentages ±SD of 5 replicates. From Lam, C., Harder, T. and Qian, P.Y. 2003. Marine Ecology Progress Series 263: 83-92, Fig. 4.

Fig. 7.17 Bioassay-guided size fractionation of Achnanthes sp., and the effect of individual, phytagelimmobilized size fractions (F1-4) on larval settlement of the polychaete Hydroides elegans (after 5 h) in comparison to a control (c) of pure phytagel (Phytagel). Data with significant differences are indicated by different letters (Tukey’s test, α= 0.05). Data plotted are mean percentages of 5 replicates). From Lam, C., Harder, T., Qian, P.Y. 2005a. Marine Ecology Progress Series, Fig. 4.

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Lam et al. (2005a) found a clear density-dependent effect of the diatom surface occupation on the induction of larval settlement. In the case of Achnanthes sp., even the lowest surface occupation of 1.8% induced larval settlement significantly higher than the control of filtered seawater. In an effort to evaluate the effect of algal extracellular polymers (EPS) as potential chemical mediators of larval settlement, it was shown that immobilized EPS-samples of inductive diatoms triggered settlement of H. elegans larvae, in contrast to EPS originating from non-inductive diatoms. These preliminary results indicate that the chemical signal perceived by surface-exploring larvae of H. elegans may be encrypted in the sequence and/or constitution of extracellular biopolymers, such as proteins and carbohydrates. 7.2.7.6 Environmental impact on bioactivity of biofilms As diatoms and bacteria in biofilms are very sensitive to changes in environmental parameters (reviewed by Wieczorek and Todd 1998), the quantity and quality of such biofilm components can reflect small-scale variation of local environmental conditions as well as suitability of the habitat for larval settlement as marine invertebrate larvae are shown to distinguish between biofilms of varying characteristics (Wieczorek et al. 1995 and references therein). This concept has been recently put under test in Qian’s laboratory. Qian et al. (2003) found that two barnacle species can clearly distinguish biofilms developed in subtidal and intertidal areas and larval settlement is not mediated by bacterial or diatom density in the biofilm or biomass of biofilm but rather by the species composition of bacteria in the biofilms. Laboratory experiments further show that larvae of two barnacle species and Hydroides elegans can distinguish the biofilms developed under different temperature and salinity conditions, although responses of three invertebrate species to the biofilm are different (Lau et al. 2005). In addition, as a direct follow-up study, Lam et al. (2005b), found that the production of EPS by benthic diatom as well as bioactivity of EPS was affected by the culture conditions of diatoms. The diatoms EPS obtained from high temperature treatments were more inductive to larval settlement than the low temperature treatments, irrespective of the salinity setting (Fig. 7.18). These results provide first-hand evidence that environmental conditions can shape the biological components of biofilms and their attractiveness to settling larvae, in turn affect recruitment and distribution patterns of marine invertebrates. 7.2.7.7 Partially characterized chemical cues Chemical inducers for the settlement or metamorphosis of marine invertebrate larvae in general have been comprehensively reviewed during the last decade (e.g. Pawlik 1992, Hadfield and Paul 2001). We may emphasize in this context that the behavioral components of settlement vs. metamorphosis, may be affected by different cues — also by different chemical cues (Qian 1999). As with natural antifoulants, there are few documented examples of natural and ecologically relevant inducers of settlement: i.e. characterized

$ Reproductive Biology and Phylogeny of Annelida

Fig. 7.18. A and C. Mean percentage of larval settlement of Hydroides elegans after 24 h in response to diatom films of Achnanthes sp. (A) and Nitzschia constricta (C) at four combinations of two different temperatures and salinities. Filtered seawater (FSW) was used as a negative control. Data that differed significantly (Tukey’s test, α = 0.05) are indicated by different letters above the bars. Data are mean percentages ±SD of 5 replicates. B. and D.: Mean diatom density of Achnanthes sp. (B) and Nitzschia constricta (D) for 4 films grown under 4 combinations of temperatures and salinities. Data are mean percentages ±SD of 3 replicates. From Lam, C., Harder, T., Qian P.Y. 2005b. Marine Ecology Progress Series, Fig. 2.

cues which induce settlement, and have been quantified and measured at in situ concentrations against relevant target organisms. There are many examples, however, of chemical compounds being critical for the benthic transition of polychaete larvae (Gee 1964). Such natural cues are watersoluble (polar) peptides and carbohydrates in particular. Fatty acids. Fatty acids have been implicated in the settlement and metamorphosis of the sabellariid Phragmatopoma lapidosa californica, which settles in response to cues deposited in the tube sand cement (Pawlik 1986, 1990). Pawlik and Faulkner (1986, 1988) showed that certain free fatty acids (FFA) present in the sand tube of Phragmatopoma lapidosa californica can induce larval settlement and metamorphosis. Palmitoleic, linoleic, arachidonic, and eicosapentaenoic acids were found in the original inductive fraction. The individual FFA induced metamorphosis over a

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range of concentrations, although abnormal metamorphosis was observed for some FFA at high concentrations only (above 300 µg FFA/g sand). Individual FFA showed relatively low levels of activity at the concentrations at which they were isolated (10 to 50 µg/g sand). In further tests of 37 different commercially available FFA, it was found that cis stereochemistry of double bonds, chain length, and the presence of a free carboxyl group were all important for larval settlement. About a dozen different FFA induced some level of metamorphosis at concentrations of 100 µg FFA/g sand. No single FFA could completely explain the settlement behavior in response to tube sand. Again, high levels of abnormal metamorphosis were observed for many FFA at concentrations above 300 µg/g sand). However, Jensen and Morse (1988) as well as Jensen et al. (1990) argued that fatty acids were contaminants that induce settlement nonspecifically in P. lapidosa californica (settlement and metamorphosis were actually induced by a polymeric protein containing L-DOPA subunits). According to Steinberg et al. (2001) there are at present no studies that unequivocally support the role of free fatty acids (FFA) as settlement inducers for polychaete larvae, nor for any other marine invertebrate. Carbohydrates. Sugars and their derivatives are increasingly implicated as inducers of settlement. Polysaccharides are perhaps the widest studied of the partially characterized larval cues from biofilms. Exopolysaccharides from the bacterium Halomonas (Deleya) marina stimulated settlement in the spirorbid Janua brasiliensis through surface recognition of a galactose-galactose subunit within the polymer (Kirchman et al. 1982a, b). Settlement of J. brasiliensis could be blocked by surface coating the biofilm with specific lectins, suggesting that the cue is surfacebound, as opposed to a water-soluble subunit broken off from the polymer. Microbial biofilms have long been recognized providing settlement cues for polychaete larvae. It could be shown for instance, that larvae of Janua brasiliensis show a strong preference for settlement on the green alga Ulva lobata. However, it could be demonstrated experimentally that the microbial films specific to U. lobata surfaces contain the inductive cue for the settlement of the larvae of J. brasiliensis (Kirchman et al. 1982a). Amino acids, peptides. Probably the most widespread evidence for the importance of soluble primary metabolites as inducers of invertebrate settlement comes from amino acids, peptides, or other proteinacious cues (Zimmer Faust and Tamburri 1994). Protein and small peptides (Jensen et al. 1990) have been claimed to act as potential chemical inducers of larval settlement in the sabellariid Phragmatopoma lapidosa californica (Jensen and Morse 1988). However, their role as a natural inducer remains to be defined, and no such compound have been identified structurally so far. Using bioassay guided isolation and purification procedures, a mixture of free amino acids isolated from the bryzoan Bugula neritina was identified on effective larval settlement inducers of Hydroides elegans (Harder and Qian 1999, see also review by Qian 1999). There was a doubt whether free amino acids were natural cues that directly acted on larvae or simply acted

& Reproductive Biology and Phylogeny of Annelida as nutrients to promote biofilm development in testing vessels. Beckmann et al. (1999) addressed this question when performing still-water laboratory assays with sterile measures in the presence and absence of antibiotic mixtures. These authors showed that the concentration of free amino acids in the testing vessel dropped indeed, while the bacterial density in Petri dishes increased. They assumed that the metamorphic response in these treatments was caused by inductive, antibiotic-resistant bacteria. To pursue this issue further, a series of carefully-designed laboratory experiments were conducted to delineate the effects of biofilm and free amino acids on larval settlement (Jin and Qian 2004). In the latter study the effects of amino acids were decoupled from possible inductive effects of bacteria in the bioassay system. The results showed that the amino acid isoleucine had no inductive effect on larval metamorphosis. Two acidic and 9 aliphatic amino acids directly induced larval metamorphosis, where the effects were concentration-dependent though. It was found that some amino acids induced higher larval settlement while not promoting bacterial growth in testing vessels. These findings indicate that free amino acids had direct inductive effects on larval settlement of H. elegans (Fig. 7.19). Inorganic ions. An elevated concentration of certain ions can induce larval settlement by affecting the electrical potential across cell membranes. Potassium has been effective in inducing larval settlement and metamorphosis of the sabellariid Phragmatopoma lapidosa californica (Yool et al. 1986). In the serpulid Hydroides elegans (Bryan et al. 1997), a slightly elevated concentration of K+ was found to be ineffective in the induction of larval settlement of H. elegans; a highly elevated concentration was found to be toxic, and treated larvae died within 24 hours, and an intermediate concentration triggered a low percentage of larvae to settle but resulted in abnormal metamorphosis (Bryan et al. 1997, Pechenik and Qian 1998, Qian and Pechenik 1998). Pechenik and Qian (1998) also found that it took much longer for the larvae to respond to elevated K+ levels than to other inducers. Also, the juveniles induced to settle by K+ had poorly calcified tubes and grew less within 9 days after settlement. It is unclear as yet why different species respond to K+ in different ways. Carpizo-Ituarte and Hadfield (1998) found that larvae of H. elegans can be stimulated to metamorphose by Cs+ ions or excess K+ ions in seawater but with kinetics quite different from the metamorphic response to biofilms. Application of a K+-channel blocker did not stimulate metamorphosis but instead inhibits both biofilm and Cs+stimulated metamorphosis. Hadfield (1998) performed experiments with Ca2+ ionophores and Ca2+-channels in transducing a metamorphic signal. Exposure to ammonia either had no effect or caused abnormal metamorphosis of H. elegans (Bryan et al. 1997). 7.2.7.8 Inhibitive cues to settlement Much effort in identifying negative cues to settlement has come from the search for antifouling natural products from marine organisms (Pawlik 1992). However, there is little evidence that extracted compounds that are

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Fig. 7.19 A. Mean percentage of metamorphosed larvae of Hydroides elegans in response to 6 amino acids (AA): Ala (alanine), Cys (cysteine), Gly (glycine), Pro (proline), Ser (serine) and Val (valine), in the absence or presence of antibiotics (AB). Data plotted are means ±SD of 5 replicates. B. Bacterial density in test dishes. Data plotted are means ±SD base on counts of 10 fields at 1000 × magnification. Data that are significantly different are indicated by different letters above the bars (one-way ANOVA, p<0.05). FSW: filtered seawater; NBF: marine natural biofilm. From Jin, T. and Qian, P.Y. 2004. Marine Ecology Progress Series 267: 209-218, Fig. 2.

 Reproductive Biology and Phylogeny of Annelida toxic or adversive to invertebrate larvae have any such function in nature. One of the few confirmations for chemical deterrence in situ comes from Woodin et al. (1997). Focusing on halogenated compounds released by many polychaete and enteropneust worms, Fielman et al. (1999), have shown inhibition of settlement of larvae of other species in the presence of these compounds. Walters et al. (1996) tested twelve macroalgae for inhibitive compounds for the larval settlement of the serpulid Hydroides elegans (and bryozoan larvae) and concluded that chemical substances provide at least one cause for the absence of surface fouling of shallow-water algae of Hawaii. However, none of the algae produced substances that inhibited larval settlement by the polychaete and bryozoan representatives. Lau and Qian (1997) found that phlorotannins were crucial in inhibiting the settlement of the polychaete Hydroides elegans on surfaces of brown algae. Investigations of H. elegans settlement has recently been extended to epibiotic communities on algae (Dobretsov and Qian 2002) and sponge (Lee and Qian 2003) surfaces. The aims of these studies were to investigate whether marine macroorganisms have evolved a symbiotic relationship with bacteria that deter invertebrate larvae from settling on their body surfaces. Therefore, recent studies have also investigated the inhibitive effect of bacteria on larval settlement of H. elegans (Lee and Qian 2003). Investigating the inhibitive effect of bacteria on larvae of H. elegans’ settlement is a technical challenge since it cannot be distinguished whether the lack of larval settlement on a bacterial film is a result of inhibition or simply the lack of inductive cue in the bacteria. To overcome this problem, an artifical settlement inducer, 3-isobutyl-1-methylxanthine (IBMX) was introduced to experimental treatments (Jensen and Morse 1988). The exact mechanism of settlement induction by IBMX is not known, but it is suggested to be a result of increasing intracellular cAMP levels in the larvae and thus inducing settlement (Clare et al. 1995). Some breakthroughs have been made in Qian’s laboratory as a group of very effective larval settlement inhibitors for H. elegans have been isolated from sponge tissues, bacteria and fungi associated with soft-bodied organisms, purified, and structurally identified (unpublished data). Young biofilm bacteria were reported by Maki et al. (1988) to inhibit settlement of the barnacle Balanus amphitrite. These results have been contradicted by those of Wieczorek et al. (1995), however, using aged (18day-old) biofilms. If reliable, these observations suggest on interesting future studies on the biological characteristics of biofilms that — depending on age — change from providing negative to positive cues for settlement. Internal processes. Some polychaete larvae settle in response to the organic or inorganic cues released from adult habitats. For instance, Cuomo (1985) reported that hydrogen sulfide (H2S), a naturally occurring product of anaerobic organic matter decomposition, promoted the larval settlement of Capitella sp. I. Larvae of this species settled in response to sulfide regardless of the presence or absence of sediment. The settlement response

Larval Ecology of the Annelida



of Capitella sp. I to H2S was concentration-dependent and the highest settlement rates were always obtained at a concentration range of 0.1-1.0 mM sulfide. Since this species is often found in organically rich sediments with high concentrations of sulfide, Cuomo (1985) suggested that larval response to sulfides may help this pioneering polychaete to locate and colonize newly created and recently disturbed muddy bottoms within a few days. However, Dubilier (1988) argued that sulfide is not a settlement cue promoting habitat selection in this species. Instead, the enhancement of settlement by sulfide in Capitella sp. I is a sublethal toxic effect. She found that larvae of Capitella sp. I ceased swimming and became lethargic at high concentrations of sulfide. The same author also demonstrated that larvae of Capitella sp. I. settled in response to organic-rich sediments in the absence of sulfide and found that an addition of sulfide delayed larval settlement and often led to abnormal metamorphosis. Settlement and metamorphosis were also delayed when larvae were exposed to water without sulfide but only under low or no oxygen conditions. She concluded that larvae of Capitella sp. I might settle in response to rich-organics rather than inorganic sulfide (Dubilier 1988). In a more recent study, Thiyagarajan et al. (2005) clearly showed for the first time a good relationship between sediment biochemical properties and larval habitat selection of Capitella sp. I. Larvae of Capitella sp. I tend to choose mud rich in nutritional values (i.e., sediments with low carbohydrate/protein ratio) not organic rich sediment. Their results suggest that types of sediment organic matter play a more important role in the larval habitat selection process of Capitella sp. I than mere concentration of organic matter. Larval physical condition and age have a profound effect on larval settlement. Qian and Chia (1993) found starved larvae of Capitella sp. and Polydora ligni lost their ability to complete settlement and metamorphosis, while starved larvae of Hydroides elegans lost their ability to respond to certain settlement cues (Qian and Pechenik 1998). Larvae may also lose substratum selectivity with age, as in the terebellid Eupolymnia nebulosa (Bhaud et al. 1990) and in Spirorbis borealis (Wisely 1958). Larvae of this species tended to avoid settling on the seaweed Fucus serratus being epibiotically fouled by adults of their own species when the density exceeded 10 individuals per cm–2. However, larvae that were kept from suitable settling surfaces for increasing periods of time showed successively abbreviations in searching behavior. When they were prevented from settling for 8 h or more, larvae appeared to settle on or very close to the substratum first, regardless of the surface contour. Pawlik and Mense (1994) found that larvae of the polychaete Phragmatopoma lapidosa became unresponsive to natural inducers when starved and regained sensitivity when they were fed again, which suggests that at least partial degradation of the metamorphic pathway occurs during starvation and is reconstructed after feeding is renewed. If larvae indeed have more than one set of receptors or more than one metamorphic

Reproductive Biology and Phylogeny of Annelida

pathway responsible for larval settlement and metamorphosis, it is easy to imagine that different types of inductive compounds may work equally well in inducing larval settlement and metamorphosis of a single species such as Phragmatopoma californica (Pawlik 1990). Qian and Pechenik (1998) found that larvae of Hydroides elegans retained their competence to metamorphose in response to IBMX for up to 10 days after becoming competent, regardless of whether they had been starved or not, which indicated that both, starvation and delayed metamorphosis had little impact on larval responsiveness to IBMX. Delayed metamorphosis, however, had a strong effect on larval responsiveness to larval homogenates as only few larvae metamorphosed in response to larval homogenates after being prevented from settling even for a period as short as 3 days. These results suggest that the two chemicals act at different points in the metamorphic pathway and that only a part of the pathway degrades as metamorphosis is delayed. During the delayed metamorphosis process, the metamorphic pathway in response to IBMX was well retained, while the pathway to conspecific cues was degrading rapidly. In addition, Pechenik and Qian (1998) found that larvae can settle much faster in response to IBMX at a younger age than in response to other artificial or natural inducers.

7.3 ACKNOWLEDGEMENTS We are particularly thankful to the editors for helpful amendments to earlier drafts of the manuscript. The present compilation was supported by the RGC grant (CA04/05.Sc01) of HKSAR to PY Qian.

7.4 LITERATURE CITED Baier, R. E. 1983. Surface chemical factors presaging bioadhesive events. Annual New York Science 416: 34-57. Banse, K. 1986. Vertical distribution and horizontal transport of planktonic larvae of echinoderms and benthic polychaetes in an open coastal sea. Bulletin of Marine Science 39: 162-175. Beckmann, M., Harder, T., and Qian, P. Y. 1999. Induction of larval attachment and metamorphosis in the serpulid polychaete Hydroides elegans by dissolved free amino acids: mode of action in laboratory bioassays. Marine Ecology Progress Series 190: 167-178. Bhaud, M. 1990. The acquisition of benthic life style by Eupolymnia nebulosa (Polychaeta, Terebellidae): An experimental approach with preliminary results. Vie et Milieu 40: 17-28. Bhaud, M. and Cazaux, C. 1987. Description and identification of polychaete larvae: their implications in current biological problems. Oceanis 13: 595-753. Bhaud, M. R. and Cazaux, C. 1990. Buoyancy characteristics of Lanice conchilega (Pallas) larvae (Terebellidae). Implications for settlement. Journal of Experimental Marine Biology and Ecology 141: 31-45. Bhaud, M. R., Cazaux, C., and Mathivat-Lallier, M. H. 1990. Delayed metamorphosis of polychaete larvae and a model for benthic life acquisition. Interaction in

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benthic recruitment between hydrodynamics of water mass and behaviour of larvae. Oceanis (Documentaire Oceanographique) 16: 207-223. Biggers, W. J. and Laufer, H. 1992. Chemical induction of settlement and metamorphosis of Capitella capitata sp. 1 (Polychaeta) larvae by juvenile hormoneactive compounds. Invertebrate Reproduction and Development 22: 39-46. Biggers, W. J. and Laufer, H. 1996. Detection of juvenile hormone-active compounds by larvae of the marine annelid Capitella sp. 1. Archive Insect Biochemistry and Physiology 32: 475-484. Blake, J. A. 1969. Reproduction and larval development of Polydora from northern New England (Polychaeta: Spionidae). Ophelia 7: 1-63. 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. Bryan, J. P., Kreider, J. L., and Qian, P. Y. 1998. Settlement of the polychaete Hydroides elegans on surfaces of the cheilostome bryozoan Bugula neritina: Evidence for a chemically mediated relationship. Journal of Experimental Marine Biology and Ecology 220: 171-190. Bryan, J. P., Qian, P. Y., Kreider, J. L., and Chia, F. S. 1997. Induction of larval settlement and metamorphosis by pharmacological and conspecific associated compounds in the serpulid polychaete Hydroides elegans. Marine Ecology Progress Series 146: 81-90. Burke, R. D. 1983. The induction of metamorphosis of marine invertebrate larvae: stimulus and response. Canadian Journal of Zoology 47: 176-188. Burke, R. D. 1986. Pheromones and the gregarious settlement of marine invertebrate larvae: stimulus and response. Bulletin of Marine Science 39: 323-331. Butman, C. A. 1987. Larval settlement of soft-sediment invertebrates: the spatial scales of pattern explained by active habitat selection and the emerging role of hydrodynamical processes. Oceanographic Marine and Biological Reviews 25: 113-165. Butman, C. A. and Grassle, J. P. 1992. Active habitat selection by Capitella sp. I larvae. 1. Two-choice experiments in still water and flume flows. Journal of Marine Research 50: 669-715. Butman, C. A., Grassle, J. P., and Buskey, E. J. 1988a. Horizontal swimming and gravitational sinking of Capitella sp. I (Annelida: Polychaeta) larvae: Implications for settlement. Ophelia 29: 43-57. Butman, C. A., Grassle, J. P., and Webb, C. M. 1988b. Substrate choices made by marine larvae settling in still water and in a flume flow. Nature (London) 333: 771-773. Carpizo-Ituarte, E. and Hadfield, M. G. 1998. Stimulation of metamorphosis in the polychaete Hydroides elegans Haswell (Serpulidae). Biological Bulletin 194: 14-24. Cazaux, C. 1985. Reproduction et developpement larvaire de l’annelide polychete saumatre Streblospio shrubsolii (Buchanan, 1980). Cahiers de Biologie Marine 26: 207-221. Chia, F. S. 1989. Differential larval settlement of benthic marine invertebrates. Pp. 312. In J. S. Ryland and P. A. Tyler (eds), Reproduction, Genetics and Distributions of Marine Organisms, Olsen and Olsen, Fredensborg, Denmark. Chia, F. S. and Rice, M. E. (eds) 1978. Settlement and Metamorphosis of Marine Invertebrate Larvae. N.Y., Elsevier, pp. 127-164. Clare, A. S. and Matsumura, K. 2000. Nature and perception of barnacle settlement pheromones. Biofouling 15: 57-68.

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Pearce, C. M. and Scheibling, R. E. 1991. Effect of macroalgae, microbial films, and conspecifics on the induction of metamorphosis of the green sea-urchin, Strongylocentrotus droebachiensis (Müller). Journal of Experimental Marine Biology and Ecology 147: 147-162. Pearse, J. S., Bosch I., Pearse, V. B., and Rasch, L. V. 1991. Bacterivory by bipinnarias: in the Antarctic but not in California. American Zoologist 31: 6A. Pechenik, J. A. 1979. Role of encapsulation in invertebrate life histories. American Naturalist 114: 859-870. Pechenik, J. A. 1990. Delayed metamorphosis by larvae of benthic marine invertebrates: does it occur? Is there a price to pay? Ophelia 32: 63-94. Pechenik, J. A. 1999. On the advantages and disadvantages of larval stages in benthic marine invertebrate life cycles. Marine Ecology Progress Series 177: 269-297. 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. Petch, D. A. 1989. Variation in the spionid polychaete Boccardia proboscidea (Hartman 1940), Ph.D. thesis, University of Melbourne. Qian, P. Y. 1999. Larval settlement of polychaetes. Hydrobiologia 402: 239-253. Qian, P. Y. and Chia, F. S. 1989. Sexual reproduction and larval development of Rhapidrilus nemasoma Monticelli, 1910 (Polychaeta: Ctenodrilidae). Canadian Journal of Zoology 67: 2345-2351. Qian, P. Y. and Chia, F. S. 1990. Detritus as a potential food source for polychaete larvae. Journal of Experimental Marine Biology and Ecology 143: 63-71. Qian, P. Y. and Chia, F. S. 1991. Effects of food concentration on larval growth and development of two polychaete worms, Capitella capitata (Fabricius) and Polydora ligni Webster. Bulletin of Marine Science 48 (2): 477-484. Qian, P. Y. and Chia, F. S. 1992. Effects of diet type on the demographics of Capitella sp. (Annelida: Polychaeta): lecithotrophic development vs. planktotrophic development. Journal of Experimental Marine Biology and Ecology 157: 159179. Qian, P. Y. and Chia, F. S. 1993. Larval growth and development as influenced by food limitation in two polychaetes: Capitella sp. and Polydora ligni. Journal of Experimental Marine Biology and Ecology 166: 93-105. Qian, P. Y. and Pechenik, J. A. 1998. Effects of larval starvation and delayed metamorphosis on juvenile survival and growth of the tube-dwelling polychaete Hydroides elegans (Haswell). Journal of Experimental Marine Biology and Ecology 227: 169-185. Qian, P. Y., Thiyagarajan, V., Lau, S. C. K., and Cheung, S. 2003. Relationship between microbial community and the attachment of acorn barnacle Balanus amphitrite Darwin. Aquatic Microbial Ecology 33: 225-237. Qian, P. Y., Rittschoff, D., Sreedhar, B., and Chia, F. S. 1999. Macrofouling in unidirectional flow: miniature pipes as experimental models for studying the effects of hydrodynamics on invertebrate larval settlement. Marine Ecology Progress Series 191: 141-151. Qian, P. Y., Rittschoff D., and Sreedhar, B. 2000. Macrofouling in unidirectional flow: miniature pipes as experimental models for studying the interaction of flow and surface characteristics on the attachment of barnacle, bryozoan and polychaete larvae. Marine Ecology Progress Series 207: 109-121. Qiu, J. W. and Qian, P. Y. 1997. Combined effects of salinity, temperature and food concentration on the early development of the polychaete Hydroides elegans (Haswell, 1883). Marine Ecology Progress Series 152: 79-88.

! Reproductive Biology and Phylogeny of Annelida Railkin, A. I. and Chikadze, S. Z. 1999. Lectin-carbohydrate interactions between hydroid larvae and microfouling films. Zoosyst Russ Suppl 1: 119-128. Rice, S. A. 1991. Reproductive isolation in the Polydora ligni complex and the Streblospio benedicti complex (Polychaeta: Spionidae). Bulletin of Marine Science 48: 432-447. Rodriguez, S. R., Ojeda, F. P., and Inestrosa, N. C. 1993. Settlement of benthic marine invertebrates. Marine Ecology Progress Series 97: 193-207. Roughgarden, J., Gaine, S., and Possingham, H. 1988. Recruitment dynamics in complex life cycles. Science 241: 1460-1466. 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. Polychaetes have evolved feeding larvae several times. Bulletin of Marine Science 67: 391-410. Rouse, G. W. 2000b. Bias? What bias? Gain and loss of down-stream larval feeding in animals. Zoologica Scripta 29: 1-25. 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. Rumrill, S.S. 1990. Natural mortality of marine invertebrate larvae. Ophelia 32: 163198. Scheltema, R. S. 1971. The dispersal of larvae of shallow-water benthic invertebrate species over long distances by ocean currents. Pp. 7-28. In Fourth European Marine Biology Symposium. D. Crisp (ed.). Cambridge University Press, Cambridge. Scheltema, R. S. 1992. Passive dispersal of planktonic larvae and the biogeography of tropical sublittoral invertebrate species. Pp. 195-202. In Marine Eutrophication and Population Dynamics. G. Colomgo et al. (eds.). Olsen and Olsen, Fredensborg, Denmark. Schroeder, P. C. and Hermans C. O. 1975. Annelida: Polychaeta. Pp. x-xx. In Reproduction of Marine Invertebrates, Vol. III A.C. Giese and J.S. Pearse (eds)., Academic Press, London. Pp. 1-213. Simon, J. L. 1967. Reproduction and larval development of Spio setosa (Spionidae, Polychaeta). Bulletin of Marine Science 17: 398-431. Slattery, M. 1992. Larval settlement and juvenile survival in the red abalone (Haliotis rufescens): an examination of inductive cues and substrate selection. Aquaculture 102: 143-153. Snelgrove, P. V. R., Butman, C. A., and Grassle, J. P. 1993. Hydrodynamic enhancement of larval settlement in the bivalve Mulinia lateralis (Say) and the polychaete Capitella sp. 1 in microdepositional environments. Journal of Experimental Marine Biology and Ecology 168: 71-109. Snelgrove, P. V. R., Grant, J., and Pilditch, C. A. 1999. Habitat selection and adultlarvae interactions in settling larvae of the soft-shell clam Mya arenaria. Marine Ecology Progress Series 182: 149-159. Steinberg, P. D., De Nys, R., and Kjelleberg, S. 2001. Chemical mediation of surface colonization. Pp. 355-387. In J. B. McClintock and J. B. Baker (eds). Marine Chemical Ecology. CRC Press, Boca Raton, Florida. Strathmann, R. R. 1990. Why life histories evolve differently in the sea. American Zoologist 30: 197-207. Strathmann, R. R. 1993. Hypotheses on the origins of marine larvae. Annual Reviews in Ecology and Systematics 24: 89-117.

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Strathmann, R. R. 1995. Mechanisms and rates of suspension feeding. Pp. 193-221. In McEdward L. (ed.) Ecology of Marine Invertebrate Larvae, CRC Press, Boca Raton, New York, London, Tokyo. Pp. 1-464. Sutherland, I. W. 1985. Polysaccharide lyases. FEMS Microbial Reviews 16: 323-347. Szewzyk, U., Holmstrøm, C., Wrangstadh, M., Samuelsson, M. O., Maki, J. S., and Kjelleberg, S. 1991. Relevance of the exopolysaccharide of marine Pseudomonas sp. strain S9 for the attachment of Ciona intestinalis. Marine Ecology Progress Series 75: 259-265. Tamaki, A. 1985. Inhibition of larval recruitment of Armandia sp. (Polychaeta: Opheliidae) by established adults of Pseudopolydora paucibranchiata (Okuda) (Polychaeta: Spionidae) on an intertidal sand flat. Journal of Experimental Marine Biology and Ecology 87: 67-82. Tamburri, M. N., Zimmer-Faust, R. K., and Tamplin, M. L. 1992. Natural sources and properties of chemical inducers mediating settlement of oyster larvae: a reexamination. Biological Bulletin 183: 327-341. 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. Thorson, G. 1950. Reproductive and larval ecology of marine bottom invertebrates. Biological Reviews 25: 1-45. Toonen, R. J. and Pawlik, J. R. 1994. Foundations of gregariousness. Nature 370: 511512. Toonen, R. J. and Pawlik, J. R. 1996. Settlement of the tube worm Hydroides dianthus (Polychaeta: Serpulidae): cues for gregarious settlement. Marine Biology 126: 725733. Toonen, R. J. and Pawlik, J. R. 2001. Settlement of the gregarious tube worm Hydroides dianthus (Polychaeta: Serpulidae). I. Gregarious and nongregarious settlement. Marine Ecology Progress Series 224: 103-114. Thiyagarajan, V., Soo, L., and Qian, P. Y. 2005. The role of sediment matter composition in larval habitat selection by the polychaete Capitella sp. I. In press. Unabia, C. R. C. and Hadfield, M. G. 1999. Role of bacteria in larval settlement and metamorphosis of the polychaete Hydroides elegans. Marine Biology 133: 55-64. Walters, L. J., Hadfield, M. G., and Smith, C. M. 1996. Waterborne chemical compounds in tropical macroalgae: positive and negative cues for larval settlement. Marine Biology 126: 383-393. Walters, L. J., Hadfield, M. G., and del Carmen, K. A. 1997. The importance of larval choice and hydrodynamics in creating aggregations of Hydroides elegans (Polychaeta: Serpulidae). Invertebrate Biology 116: 102-114. Watanabe, N., Watanabe, S., Ide, J., Watanabe, Y., Sakata, K., and Okamoto, K. 1998. Chemical signals involved in larval metamorphosis in Hydroides ezoensis (Serpulidae; Polychaeta). Part II: Isolation and identification of a new monoacyl glycerol from adult tube clumps as a metamorphosis-inducing substance. Journal of Marine Biotechnology 6: 11-15. Weinberg, J. R., Starczak, V. R., Mueller, C., Pesch, G., and Lindsay, S. 1990. Divergence between populations of a monogamous polychaete with male parental care: Premating isolation and chromosome variation. Marine Biology 107: 205-213. Wieczorek, S. K. and Todd, C. D. 1998. Inhibition and facilitation of settlement of epifaunal marine invertebrate larvae by microbial biofilm cues. Biofouling 12: 81118.

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Wieczorek, S. K., Clare, A. S., and Todd, C. D. 1995. Inhibitory and facilitatory effects of microbial films on settlement of Balanus amphitrite amphitrite larvae. Marine Ecology Progress Series 119: 221-228. Wilson, D. P. 1932. On the Mitraria larva of Owenia fusiformis. Philosophical Transactions of the Royal Society of London Series B 221: 231-334. Wilson, D. P. 1968. The settlement behaviour of the larvae of Sabellaria alveolata (L.). Journal of the Marine Biological Association of the U.K. 48: 387-435. Wilson, D. P. 1974. Sabellaria colonies at Duckpool, North Cornwall, 1971-1972, with a note for May 1973. Journal of the Marine Biological Association of the U.K. 54: 393-436. 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. Woodin, S. A. 1986. Settlement of infauna: larval choice? Bulletin of Marine Science 39: 401-407. Woodin, S. A., Marinelli, R. L., and Lincoln, D. E. 1993. Allelochemical inhibition of recruitment in a sedimentary assemblage. Journal of Chemical Ecology 19: 517530. Woodin, S. A., Lindsay, S. M., and Lincoln, D. E. 1997. Biogenic bromophenols as negative recruitment cues. Marine Ecology Progress Series 157: 303-317. Wray, G. A. 1995. Evolution of larvae and developmental modes. Pp. 413-447. In McEdward, L. (ed.) Ecology of Marine Invertebrate Larvae. Boca Raton: CRC Press. Yool, A. J., Grau, S. M., Hadfield, M. G., Jensen, D. A., Markell, D. A., and Morse, D. E. 1986. Excess potassium induces larval metamorphosis of four marine invertebrate species. Biological Bulletin 170: 255-266. Young, C. M. 1990. Larval predation by epifauna on temperate reefs: scale, power and the scarcity of measurable effects. Australian Journal of Ecology 15: 413-426. Young, C. M. (ed.) 2002. Atlas of Marine Invertebrate Larvae. Academic Press. Pp. 1-626. Young, C. M. 1995. Behavior and locomotion during the dispersal phase of larval Life. Pp. 249-278. In McEdward, L. (ed.) Ecology of Marine Invertebrate Larvae, CRC Press, Boca Raton, New York, London, Tokyo. Pp. 1-464. Young, D. M. and Braithwaite, L. F. 1980. Orientation and current-induced flow in the stalked ascidian Styela montereyensis. Biological Bulletin 159: 428-439. Zajac, R. N. 1984. Population dynamics and life history variation in Polydora ligni (Annelida: Polychaeta). Ph.D. dissertation, University of Connecticut, Storrs, CN. Zimmer-Faust, R. K. and Tamburri, M. N. 1994. Chemical identity and ecological implications of a waterborne larval settlement cue. Limnology and Oceanography 39: 1075-1087.

II Selected Groups of Annelida

CHAPTER

8

Non-leech Clitellata Barrie G.M. Jamieson1 with contributions by Marco Ferraguti2*

8.1 PHYLOGENY AND SYSTEMATICS 8.1.1 Definition of Clitellates The Clitellata Michaelsen 1919 are annelids which include the oligochaetes (earthworms and their allies), branchiobdellids (ectoparasites of freshwater crayfish) and leeches. They are defined by the possession of a modification of the epidermis, the clitellum, which is located at least partly behind the female pores and which secretes a cocoon in which the eggs are deposited. They were renamed Euclitellata (a term not employed in this volume though widely used) by Jamieson because a clitellum also occurs in questid polychaetes though there anterior to the female pores (Jamieson 1983c). Questa has since been placed, from 18S rRNA, close to the eunicid polychaetes (Erséus and Källersjö 2003) but showed firm relationship to the Orbiniidae (with 95% jackknife support) in an unpublished analysis of 215 annelid taxa, using 18S, 16S, 28S, COI and H3, by Siddall et al. (pers. comm.). In view of the synonymy of the terms Clitellata and Oligochaeta sensu lato, containing oligochaetes sensu stricto, acanthobdellids, branchiobdellidans and true leeches (Siddall et al. 2001; Erséus et al. 2002) it is appropriate to adopt the term Clitellata for the combined assemblage and to dispense with the term Oligochaeta as an official name, a procedure also adopted by Erséus and Källersjö (2003) (but see Chapter 9). Nevertheless, the term ‘oligochaete’ will here be used as a common noun for Oligochaeta sensu stricto, to avoid the inconvenience of referring to them as non-branchiobdellidan, non-acanthobdellid, non-leech Clitellata. Alternatively they may be referred to as ‘oligochaetous clitellates’, though excluding the similarly characterized Acanthobdella. 1

Department of Zoology and Entomology, University of Queensland, Brisbane, Queensland 4072, Australia 2 Dipartimento di Biologia, Università di Milano, Via Celoria 26, I-20133, Milano, Italy *Section 8.4

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8.1.2 Molecular Evidence for Higher Clitellate Relationships Molecular studies have confirmed monophyly of the clitellates, using 18S rRNA (Moon et al. 1996; Struck et al. 2002; Erséus and Källersjö 2003); or 18S rRNA (Winnepenninckx et al. 1998); 18S and mtCOI (Martin 2000; Siddall et al. 2001); cytochrome oxidase I (Siddall and Burreson 1998; Nylander et al. 1999) or elongation factor lalpha (Kojima 1998; see also a review by McHugh 2000). An apparent exception to clitellate monophyly in a parsimony analysis of 18S rRNA was considered questionable (Martin 2001, see below). With regard to the position of the clitellates within the Annelida, molecular analysis has indicated that clitellates form a clade within the Polychaeta and that polychaetes are a paraphyletic or polyphyletic group (Kojima 1998; McHugh 2000; Martin 2001; Rota et al. 2001; Siddall et al. 2001; Erséus and Källersjö 2003). Paraphyly of the Oligochaeta, with leeches and/or branchiobdellidans lying within the oligochaete clade, has long been suspected on morphological grounds (Michaelsen 1928; Brinkhurst and Nemec 1986; Jamieson et al. 1987; Jamieson 1988b; Brinkhurst 1989; Brinkhurst and Gelder 1989; Purschke et al. 1993). Derivation of leeches from a lumbriculid-like ancestor had been suggested on morphological grounds by Michaelsen (1928-1932), Brinkhurst and Nemec (1986) and Jamieson et al. (1987). These relationships are being increasingly confirmed from molecular analyses. Thus, using 18S rDNA and mitochondrial cytochrome c oxidase subunit I, parsimony analysis by Siddall et al. (2001) of the combined data-set gave complete support for monophyly of leeches (Hirudinea), branchiobdellidans (Branchiobdellida), and acanthobdellidans (Acanthobdellida), as already shown in Siddall and Burreson (1998). Monophyly of the oligochaete order Lumbriculida with these three groups was also strongly supported. These results provided support for the hypotheses that leeches and branchiobdellidans are sister groups, that acanthobdellidans are sister to them, as suggested by Livanow (1931), and that together with the family Lumbriculidae they all constitute a clade within Oligochaeta. Synonymy of the Clitellata and the more commonly used Oligochaeta was therefore proposed. It was considered that leeches branchiobdellidans, and acanthobdellidans should be regarded as orders equal to their closest relatives, the order Lumbriculida. However, leaving aside the issue of recognizing ordinal ranks, these clitellate entities are not here considered to be equivalent in rank. Martin (2001) had also investigated phylogenetic relationships of clitellates using 18S rRNA, aligning on the basis of secondary structure and applying maximum parsimony (MP) and maximum likelihood (ML). In contrast to ML, parsimony did not recover the monophyly of clitellates. However, a close scrutiny of the data suggested a spurious attraction between some polychaetes and clitellates. Acanthobdellida and Euhirudinea were reconciled in their traditional Hirudinea clade and were included in the

Non-leech Clitellata

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Oligochaeta with the Branchiobdellida ‘via the Lumbriculidae’ as a possible link between the two assemblages. The exact position of Hirudinea and Branchiobdellida within oligochaetes and of Clitellata within the Polychaeta remained unresolved. Polytomies within annelids were considered to indicate an extremely ancient radiation of polychaetes and emergence of clitellates. It has since emerged that the polychaete sequences taken from another source which led to the non-monophyly of Clitellata by Martin (2001) contained sequencing errors; these have been replaced in genbank with correct sequences (Siddall, pers. comm.). Jamieson et al. (2002) found support, from 28S rDNA, for the following clades, among others: (1) Enchytraeidae; (2) Naididae + Tubificidae; (3) Lumbriculidae; (4) Lumbriculidae + (Hirudinida + Brachiobdellida); (5) Crassiclitellata; and (6), within the Crassiclitellata, Ocnerodrilidae + (Acanthodrilinae + Megascolecinae) (Fig. 8.1). The leech-branchiobdellid relationship has been controversial owing to the concern that molecular artifacts (under the rubric of long branch attraction) undermine confidence in the result. This study revealed significant base composition variation in the 28S (p < 0.001 PAUP* test), in particular the leeches and the branchiobdellids being distinct from other clitellates. Bootstrap support for the leech-branchiobdellid grouping was fairly high in maximum parsimony (MP) (76%) but not sustained in maximum likelihood) (ML) (16%), and the grouping was conspicuously absent using LogDet, a method that is supposed to compensate for nonstationarity (Lockhart et al. 1994). Furthermore, parametric data-sets made to trees that did not contain this clade, when analysed with MP, grouped them at high bootstrap frequency (> 70%); it was therefore suspected that MP support was artifactually high. Conversely, it was considered that low ML support might be due to long branch repulsion effects (see Siddall 1998). Thus, although the branchiobdellid-leech clade was supported it was not without reservation. Erséus and Källersjö (2003) have given a tree based on 18S analysis using Clustal and PAUP (Fig. 8.2) which confirms a sister group relationship of Hirudinida and Branchiobdellida with Acanthobdella as their plesiomorph sister group; the Lumbriculidae formed the plesiomorph sister group of the Acanthobdella through Hirudinida assemblage. Hugall et al. (unpublished), in a study using a different computational method (for details see Enchytraeidae, below) for the three genes 18S, 28S and COI, corroborated this clade but in a sequence ((Hirudinea) ((Branchiobdellida) (Acanthobdella, Lumbriculidae))) (Fig. 8.5).

8.1.3 Phylogeny and Classification of Oligochaetes Sensu Stricto Oligochaetes sensu stricto are marine, freshwater and terrestrial. Unlike leeches, only one species is known to have parasitic forms, in snails. The circummundane naidine Chaetogaster limnaei, found throughout the world in the mantle cavity of snails where it is known to reduce numbers of trematode cercariae (see Fernandez et al. 1991), has a truly parasitic subspecies,

!& Reproductive Biology and Phylogeny of Annelida

Fig. 8.1. 28S rDNA maximum parsimony majority rule consensus of 1000 bootstrap resamples, five random sequence additions each with tree bisection reconnection swapping. Basal polytomy rooted with Haplotaxis gordioides (Hartmann, 1821). Maximum parsimony bootstraps above node, maximum likelihood bootstrap values below (200 resamplings from reduced 36 taxa set using only nine Megascolecidae). Those underlined were not recovered in the majority rule. Family and some higher level groups indicated. From Jamieson et al. 2002. Zoosystema 24(4): 707-734, Fig. 2.  Publications Scientifiques du Muséum national d’Histoire naturelle, Paris.

Non-leech Clitellata

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Fig. 8.2. 18S rDNA phylogeny of basal groups of Clitellata (Annelida). From Erséus, C. and Källersjö, M. 2003. Zoologica Scripta: 33(2): 187-196, Fig. 4.

C. l. vaghini (Gruffydd 1965). An Indian naidid, Haemonais, inhabits the mantle cavity of Vivipara bengalensis. The pheodrilid Astacopsidrilus is commensal in the branchial chamber of Australian crayfish (pers. obs.) and Rota (pers. comm.) reports that Lumbricillus catanensis is abundant in the branchial chambers of freshwater crabs in Sicily; and that Enchytraeus carcinophilus lives in the same habitat in Gecarcinus lagostoma from Trinidade Island, Brazil. The enchytraeid Fredericia parasitica ingests mucus and, apparently, coelomocytes from the surface of Allolobophora robusta, and the

" Reproductive Biology and Phylogeny of Annelida highly modified, sucker-bearing Aspidodrilus kelsalli on an African earthworm probably takes similar food although the gut also contains ‘mud’ and vegetable matter (see Jamieson 2000). Haereodrilus reichelti is a Diporochaeta-like species which lives on the surface of another megascolecine, the giant Digaster grandis in South East Queensland (Dyne, in Jamieson 2000) With the exception of some earthworm-like genera, aquatic oligochaetes are usually small and are loosely termed microdriles. However, some species of the ‘microdrile’ assemblage are as large as earthworms, namely giant enchytraeids and some Moniligastridae. Microdriles are characterized by a plesiomorphic type of clitellum in which, like the epidermis from which it is derived, there is only a single layer of cells. Its simple structure and limited ability to secrete nutrients into the cocoon correlate with the production of small numbers of large, yolky eggs. However, naidines do show some degree of albumenotrophy (see 8.6.6. Embryogenesis. Yolk content and cleavage). Prostate glands usually secreting into the male ducts, or the glandular sperm funnels in enchytraeids, presumably add secretions to the seminal fluid and contribute to the small amount of albumen in the cocoon. A major evolutionary innovation in earthworms (loosely termed megadriles), in contrast, has been the development of a clitellum consisting of several layers of cells with the ability to secrete large proteinaceous reserves into the cocoon (Jamieson 1971c). Correlated with this, the eggs possess little yolk, are therefore small and are produced in large numbers (Jamieson 1992). In morphocladistic analysis, all families with multilayered clitella were found to form a single clade, named the Crassiclitellata (Jamieson 1988b). Thus acquisition of a multilayered clitellum was deduced to be a monophyletic event. However, Omodeo (2000), in an informative review of evolution and biogeography of megadriles, implied that a multilayered clitellum has arisen more than once when he derived the Eudrilidae (with multilayered clitellum) from the Alluroididae (with clitellum consisting of a single layer of cells) independently of other earthworms. Jamieson (1974) had also suggested a relationship between the Alluroididae and Ocnerodrilidae but neither of these familial relationships is supported morphocladistically (Jamieson 1988b). The higher classification of the Oligochaeta sensu stricto adopted here (Table 8.2) is based on the morphocladistic parsimony analysis by Jamieson (1988b), but is modified to take into account molecular (rDNA and RNA) findings (Figs. 8.1, 8.2, 8.5-8.7). The morphocladistic analysis did not include branchiobdellidans and leeches. It produced a single most parsimonious tree resulting in the classification shown in Fig. 8.3 and 8.4A, B. No great significance was attached to Linnean and other rank names above the familial level but they at least served to show the relative levels in the phylogenetic hierarchy, which it is cumbersome to represent by any numerical system. Ranks are omitted from the revised classification in Table 8.2. Detailed definitions of oligochaete families are given by Jamieson (1978b; 1980) and features of their reproductive anatomy are discussed in Section 8.2 below and are partly shown in Fig. 8.4.

Fig. 8.3. Cladogram to show sister-group relationships of superfamilies and high taxa recognized by Jamieson (1988) in a morphocladistic analysis. The Randiellidae (not computed) were tentatively assigned to a new subclass regarded as the most plesiomorphic descendant of a presumed octogonadal ancestor. From Jamieson, B.G.M. 1988. Cladistics 4(4): 367-402, Fig. 4.

Non-leech Clitellata

"

"

Reproductive Biology and Phylogeny of Annelida

Colour Figure

Fig. 8.4. A and B. Cladogram derived from a single most parsimonious tree, represented only to families, showing the taxonomy adopted in a morphocladistic analysis. Naididae were not included in the original

Although the Jamieson (1988b) classification is more complex than that advanced, without recourse to parsimony cladistics by Brinkhurst and Jamieson (1971), it confirmed the order Lumbriculida, family Lumbriculidae and the order Moniligastrida (a single family, Moniligastridae). It restricted the Haplotaxida (previously the Haplotaxidae and putative opisthopore descendants of a common ancestor shared with these) to the Haplotaxidae and Tiguassuidae and recognized the order Opisthopora (male pores more than one segment behind the segment

Non-leech Clitellata

"!

Colour Figure

analysis and are now subsumed in the Tubificidae. Based on Jamieson, B.G.M. 1988. Cladistics 4(4): 367-402, Fig. 3.

containing the posteriormost testes) for the residue of the Haplotaxida (including all the “earthworms”) of the earlier work. The monophyletic clade Crassiclitellata (confirmed in molecular analyses by Jamieson et al. 2002; Erséus and Källersjö 2003) was the apomorph sister-group of the Alluroidoidea, the whole forming the order Opisthopora which with the order Moniligastrida comprised the superorder Metagynophora. The crassiclitellates contained, in the morphocladistic analysis, two sister-clades, with independent, ecological

"" Reproductive Biology and Phylogeny of Annelida support. One of these, named the Aquamegadrili, contained the aquatic megadrile families Sparganophilidae, Biwadrilidae, Almidae (with Criodrilus), and Lutodrilidae, with, questionably, the Kynotidae. The other, named the Terrimegadrili, contained the predominantly terrestrial families Ocnerodrilidae, Eudrilidae, Microchaetidae, Hormogastridae, Glossoscolecidae, Lumbricidae and Megascolecidae (Jamieson 1988b). We will see in the discussion of molecular analyses that recognition of Aquamegadrili and Terrimegadrili is not supported, though at present only a single family (Komarekionidae) transgresses the division. The morphocladistic analysis (Jamieson 1988b) attributed a basal position to the Eudrilidae and especially the Ocnerodrilidae relative to the other terrimegadrile families, whereas these two families are usually considered close to the Megascolecidae. We will see that molecular data do not support this distant position for ocnerodrilids but reunite them with the Megascolecidae. The single eudrilid included in the molecular analysis (Fig. 8.1) associates with the Glossoscolecidae and is not confirmed as a sister taxon of the Ocnerodrilidae + Megascolecidae but this finding requires confirmation from larger samples. In the morphocladistic analysis (Jamieson 1988b), the Haplotaxidae and Tiguassu, confirming suspected paraphyly of the Haplotaxidae, were successively basal to the Metagynophora, the combined clade being the Diplotesticulata which questionably contained the Lumbriculidae (no longer placed there), the latter forming the subclass Lumbriculata. The microdrile families Tubificidae (and Naididae) and the Phreodrilidae, and the Enchytraeidae formed a monophyletic clade, termed the Tubificata, the sistertaxon of the Lumbriculid-Diplotesticulata clade. As discussed below, this basal position of the Enchytraeidae is particularly challenged by some molecular analyses (Siddall et al. 2001; Erséus and Källersjö 2003; Hugall et al., unpublished). Impact of molecular studies. In the analysis of leech relationships by Siddall et al. (2001), which used polychaete outgroups, a sufficient representation of oligochaete families was included to allow assessment of hitherto uncertain interrelationships of major groups of families. Thus, although Siddall et al. (2001) did not refer to it, the suggestion of Stephenson (1930) that Tubificidae and related families were derived groups was contraindicated. Enchytraeidae were divorced from the Tubificidae + Naididae and had an unexpected association with the two oligochaete families represented. Their analysis pointed to the existence of the following main groups of oligochaetes [my enumeration]: 1) an aquatic ‘microdrile’ clade consisting of Tubificidae (including Naididae) and Phreodrilidae and 2) a clade consisting of 2a) the Enchytraeidae (aquatic and terrestrial and sometimes of ‘megadrile’ proportions) with earthworms (e.g. Lumbricidae and Megascolecidae) and 2b) a well defined clade consisting of Lumbriculidae + Acanthobdellidae + Branchiobdellidae + leeches. It should be noted, however, that linkage of the latter clade (2b) with the enchytraeid + earthworm clade was given no numerical support

Non-leech Clitellata

"#

in their consensus tree. The 18S analysis of Erséus and Källersjö (2003) (Fig. 8.2) with its more detailed representation of the Oligochaeta sensu stricto, confirmed the major groupings of Siddall et al. (2001). It should be noted, however, that in the study of Erséus et al. (2002), the Enchytraeidae and Lumbricidae, arbitrarily chosen as the outgroup, failed to form a monophylum in relation to the ingroup consisting of Tubificidae, Naididae, Phreodrilidae and Lumbriculidae. Many families remain to be analyzed for molecular sequences. It is particularly desirable that the Moniligastridae, putatively the most basal group of the Metagynophora and the plesiomorph sister-group of the Opisthopora, and the Alluroididae, which are putatively the most basal of the Opisthopora and form the plesiomorph sister-group of the Crassiclitellata (Jamieson 1988b), should be included.

8.1.4 Phylogeny of the Oligochaete Families The following section will be structured according to the morphocladistic analysis (Jamieson 1988b) but departures from the latter will be noted, some additional families included and some groupings and names recognized in the analysis will be questioned or replaced in light of new evidence chiefly from molecular analyses. Features of the reproductive systems of the various families will be included in Section 8.2, below. After this discussion in the context of the morpocladistic analysis, a revised, simplified classification of the oligochaetous clitellates will be given. Capilloventridae. This family was not included in the morphocladistic analysis (Jamieson 1988b) though it was provisionally placed in the Tubificata. Despite the complex and probably apomorphic condition of the midpiece of the spermatozoon, Ferraguti et al. (1996) considered spermatozoal ultrastructure to indicate that capilloventrids were basal oligochaetes. The 18S analysis of Erséus and Källersjö (2003) (Fig. 8.2) confirms capilloventrids as the most basal known oligochates and the plesiomorph sister group of the Proppapidae + Haplotaxidae + Phreodrilidae + Tubicidae clade. The basal position of Capilloventer is further confirmed in the 215 taxon analysis (Siddall, pers. com.). Randiellata. Whether the little known Randiellidae deserve a position, as the subclass Randiellata, basal to all other oligochaetes requires further investigation. Evidence for recognition of the subclass Randiellata is discussed in the taxonomic review of reproductive systems below (8.2.11.1). Tubificata. Order Tubificida. The Tubificida was the sole taxon of the Tubificata and the latter name can therefore be regarded as redundant. These were defined as having one pair of testes followed by one pair of ovaries; male ducts plesioporous (penetrating one intersegment). They included the Tubificidae, Enchytraeidae, Phreodrilidae, Dorydrilidae, and Opistocystidae. The Capilloventridae, though not included in the character matrix, were tentatively included but are here removed (see above). It was considered possible that the imperfectly known Randiellidae (see above) should be included. Inclusion of the Enchytraeidae is now questionable.

"$ Reproductive Biology and Phylogeny of Annelida Propappidae. These were not included in the morphocladistic analysis but appear to be referable to the Tubificida. Separation (Coates 1986) of the Propappidae from the Enchytraeidae in which they were formerly grouped appears to be supported by the 18S analysis of Erséus and Källersjö (2003) (Fig. 8.2) in which Propappus forms a clade with the Haplotaxidae, Phreodrilidae and Tubificidae, far separated from the Enchytraeidae. Enchytraeidae. The position of the Enchytraeidae particularly needs reinvestigation. From general morphology (Jamieson 1988b) and sperm ultrastructure (Jamieson 1981c, 1983a; Ferraguti and Erséus 1999) the Enchytraeidae have been considered to be a more basal group than the Tubificidae and other oligochaetes and to have no close affinity with crassiclitellates. Siddall et al. (2001) demonstrated a grouping (2a) of Enchytraeidae and earthworms (crassiclitellates) on the basis of 18S rDNA and mitochondrial cytochrome oxidase subunit I. However, the analysis, and the enchytraeid + (lumbricid+Pontodrilus) clade, may not be robust as one of the lumbricids (Eisenia fetida) forms the sister taxon of the megascolecid Pontodrilus, the remaining lumbricid (Lumbricus terrestris) being the sister taxon of that clade. Lumbricids show greater molecular homogeneity in our studies (Jamieson et al. 2002) and Pontodrilus is an undoubted, if somewhat aberrant, member of the Megascolecinae from analysis of the three genes 28S + 12S + 16S. In support of the above proposition of Siddall et al. (2001), Erséus and Källersjö (2003) (Fig. 8.2) show enchytraeids as the plesiomorph sister group of a small sample of crassiclitellate species in the Megascolecidae, Lumbricidae and Almidae (Criodrilus). Insofar as crassiclitellates must have arisen from a ‘microdrile’ group, origin of crassiclitellates from an ancestor shared with the Enchytraeidae deserves consideration. Enchytraeids resemble at least some crassiclitellates in having terrestrial members, simple chaetae (though the crassiclitellate Pontoscolex has some bifid somatic chaetae), in including some species of ‘megadrile’ proportions, and in location of the spermathecal pores well anterior to the testes, a location which seems basic for crassiclitellates (with haplotaxids). Other ‘microdrile’ families have been elimnated in molecular studies from sharing direct ancestry with crassiclitellates. The sister group relationship of enchytraeids and crassiclitellates cannot yet be considered certainly established, however. Thus, Hugall et al. (unpublished) (Fig. 8.5; Table 8.1), used combined nuclear 18S, 28S and mitochondrial COI genes to assess oligochaete, and particularly enchytraeid, relationships. Computation used the MrBayes Markov Chain Monte Carlo (MCMC) likelihood program developed in the Manhattan project. MCMC provides an efficient means of obtaining an estimate of relative likelihood (or relative probability of occurrence) of models across a very large number of competing possibilities while giving the relative support for each possibility. A consensus tree was obtained for all three genes, excluding (Fig. 8.5A), or including (Fig. 8.5B), the third position in mtDNA CoI. Taxa included in the analysis are listed in Table 8.1. Most of the operational taxonomic units were

Non-leech Clitellata

"%

Fig. 8.5. Analyses of phylogenetic relationships of annelids using combined nuclear 18S, 28S and mitochondrial COI genes. Genes used 18S, 28S (nuclear genes) and COI (mtDNA). Not all taxa have all genes (see Table 8.1). Computation uses the MrBayes Markov Chain Monte Carlo likelihood program. Consensus trees for all three genes but excluding the third position in mtDNA CoI. The dashed lines indicate the result of constraining enchytraeids as the sister group of crassiclitellates. A. Excluding third bases in mtDNA COI. B. Including third bases. Taxa are as listed in Table 8.1. Note that most of the sequences are combined from more than one species. For instance, the megascolecine combines different genes for Pontodrilus litoralis, Perionychella cf kershawi, and Pheretima sp. In contrast Lumbriculus variegatus denotes a single species for all three genes. (From Hugall et al., unpublished.)

"& Reproductive Biology and Phylogeny of Annelida Table 8.1 Species sampled, accession numbers and base sites for molecular phylogenetic analysis. The Operational Taxonomic Units listed in the first column are those included in the cladograms in Fig. 8.5 (From Hugall et al., unpublished). 18S

OTU Lumbricidae Lumbricid 1 Eisenia fetida Megascolecidae Megascolecinae Enchytraeidae Enchytraeid 1 Enchytraeid 2 Lumbriculidae Lumbriculus variegatus Lumbriculid 2 Rhynchelmis Tubificidae Tubificid Naidinae Dero Branchiobdellida Cambarincola Branchiobdellid 2 Hirudinea Hirudinea 1 Hirudinea 2 Acanthobdellida Acanthobdella Aphanoneura Aeolosoma Mollusca Bivalve Gastropod Polychaeta Polychaete 1 Polychaete 2

18S

Accession

Sites

Lumbricus rubellus Eisenia fetida

Z83753 X79872

1711 1713

Pontodrilus bermudensis

-

0

Enchytraeus sp., Kim Enchytraeus sp., Aguinaldo

Z83750 U95948

1716 1713

Lumbriculus variegatus Trichodrilus diversisetosus Rhynchelmis shamanensis

AF209457 AJ252318 AJ308512

1709 480 1705

Tubifex sp.

U67145

1712

Dero digitata

AF021879

1708

Cambarincola holti Xironogiton victoriensis

AF115975 AF115977

1712 1711

Erpobdella punctata Hemiclepsis marginata

AF116002 AF115981

1713 1714

Acanthobdella peledina

AF099953

1711

Aeolosoma sp.

Z83748

1706

Glycymeris sp. Helix aspersa

X91978 X91976

1711 1708

Nereis limbata Lanice conchilega

U36270 X79873

1701 1710

Lumbricid sp. Eisenia fetida

AY048498 AY048508

670 670

Perionychella kershawi

AY048484

670

Achaeta bohemica Fridericia bisetosa

AF406595 AF406596

664 659

28S Lumbricidae Lumbricid 1 Eisenia fetida Megascolecidae Megascolecinae Enchytraeidae Enchytraeid 1 Enchytraeid 2

Table 8.1 contd

"'

Non-leech Clitellata Table 8.1 contd

Lumbriculidae Lumbriculus variegatus Lumbriculid 2 Rhynchelmis Tubificidae Tubificid Naidinae Dero Branchiobdellida Cambarincola Branchiobdellid 2 Hirudinea Hirudinea 1 Hirudinea 2 Acanthobdellida Acanthobdella Aphanoneura Aeolosoma Mollusca Bivalve Gastropod Polychaeta Polychaete 1 Polychaete 2

Lumbriculus variegatus Lamprodrilus sp. Rhynchelmis brachycephala

AF406594 AF406592 AF406593

651 668 668

Branchiura sowerbyi

Jamieson et al. 2003

659

Dero aulophorus

AF406598

648

Cambaricola pamelae Xironodrilus formosus

AF406601 AF406600

657 663

MGLeech FWleech

Jamieson et al. 2003 Jamieson et al. 2003

667 649

Acanthobdella peledina

-

0

Aeolosoma sp.

-

0

Mytilus edulis Chloritis anaxbanneri

Z29550 Hugall et al., unpublished

666 658

Phyllodoce Amphitritides harpa

Hugall et al., unpublished AF185180

669 265

Lumbricus terrestris Eisenia fetida Pontodrilus bermudiensis Achaeta bohemica Fridericia tuberosa Lumbriculus variegatus Agriodrilus vermivorus Rhynchelmis alyonae Tubifex tubifex Dero digitata Cambarincola holti Xironogiton victoriensis Hemiclepsis marginata Erpobdella punctata Acanthobdella peledina Aeolosoma litorale Mytilus edulis Albinaria coerulea Neanthes japonica Paralvinella palmiformis

U24570 AF003256 AF064047 other AJ252319 AJ252320 U74076 AF054195 AF116012 AF116014 AF003259 AF003275 AF003264 AF054188 M83762 NC_001761 D38032 U74070

651 0 650 0 573 632 573 573 651 573 651 651 651 651 651 573 622 648 504 651

COI Lumbricid 1 Eisenia fetida Megascolecinae Enchytraeid 1 Enchytraeid 2 Lumbriculus variegatus Lumbriculid 2 Rhynchelmis Tubificid Dero Cambarincola Branchiobdellid 2 Hirudinea 1 Hirudinea 2 Acanthobdella Aeolosoma Bivalve Gastropod Polychaete 1 Polychaete 2

# Reproductive Biology and Phylogeny of Annelida represented by sequences combined from more than one species where these unequivocally belonged to the same, named OTU. For instance, the megascolecine OTU combined different genes for Pontodrilus litoralis, Perionychella cf kershawi, and Pheretima sp. In contrast some OTUs denote a single species (Lumbriculus variegatus, Acanthobdella peledina, Eisenia fetida) or a single genus (Rhynchelmis, Cambarincola, Dero, Aeolosoma) for all three genes. A composite bivalve (Glycymeris, Mytilus), gastropod (Helix, Chloritis) and two composite polychaetes constituted the outgroup. With regard to the position of the Enchytraeidae, represented by two composite OTUs, the enchytraeid clade was the plesiomorph sister-group (at 94%) of a large crassiclitellate + leech (Hirudinea + branchiobdellid + acanthobdellid + lumbriculid) clade when third position COI was excluded. The molecular analysis of Erséus et al. (2002) also failed to group enchytraeids exclusively with earthworms (represented by Lumbricidae), as is also the case in Fig. 3 (not shown) of Erséus (2003). In the Hugall et al. analysis, if enchytraeids were constrained as the sister-group of the crassiclitellates, support was only 48% (Fig. 8.5A). When third position COI was included, however, the enchytraeid clade was the plesiomorph sister-group of the Crassiclitellata represented by lumbricids and megascolecids (Fig. 8.5B) as suggested by Siddall et al. (2001) and Erséus and Källersjö (2003). However, in this case the support was at only 52% whereas if enchytraids were constrained as the sister-group of the larger clade (including leeches and allies), support was a close 48%. In both analyses the tubificid-naid clade was the most basal in the Oligochaeta (Hugall et al., unpublished) but capilloventrids were not included in the analysis. Tubificidae and Naididae. The combined data set of Siddall et al. (2001) did not resolve relationships within the tubificid + naidid clade. Phylogenetic analyses by Christensen and Theisen (1998) based upon the D3 domain of the nuclear 28S rRNA gene and part of the mitochondrial COI gene indicated that the family Naididae is a subordinate group within the family Tubificidae and is most closely related to the subfamily Rhyacodrilinae. Based upon relative genetic distances they hypothesized that the origin of Naididae occurred relatively early in the evolution of Tubificidae, though a fairly late event compared to the basic radiation among Annelida. Erséus and Källersjö (2003) and Erséus et al. (2000, 2002), using cladistic analysis of sequences of the 18S rDNA gene, also showed that the Naididae must be subsumed in a monophyletic Tubificidae, and are related to some “rhyacodriline” groups within the Tubificidae but further stated that “naidids” may not even constitute a monophyletic group. The family name Naididae was formally suppressed by Erséus et al. (2002) as a junior synonym of the Tubificidae (Application to the ICZN by Erséus and colleagues for recognition of this decision is pending as the name Naididiae in fact has chronological precedence). The subfamily Naidinae, within the Tubificidae, was recognized by Erséus and Gustavsson (2002) and Erséus (2003). A tubificid+naid clade was confirmed from 28S in Jamieson et al.

Non-leech Clitellata

#

(2003) (Fig. 8.1) and in the combined 18S, 28S, and COI analysis of Hugall et al. (unpublished) (Fig. 8.5). The analyses of Erséus et al. (2000, 2002) corroborated the idea that the Tubificinae are monophyletic but indicated that the Rhyacodrilinae and the Phallodrilinae are not. Bathydrilus did not appear to be closely related to other “phallodriline” genera. Limnodriloidinae appeared monophyletic in the first study but were unresolved in the second. Of tubificid genera tested for monophyly, several were fully supported by the data, namely Ainudrilus, Bathydrilus, Limnodrilus, Smithsonidrilus, Thalassodrilides. Strong (but not 100%) jackknife support was obtained for Heterodrilus, Tectidrilus and Heronidrilus. The gutless genera Inanidrilus and Olavius were found not to be sister-taxa but collectively formed a reasonably well supported group (85%). Only one genus (Tubifex) was refuted, and two (Tubificoides and Limnodriloides) were unresolved from other taxa (Erséus et al. 2002). Tubifex tubifex siblings. Molecular analyses have also proved of value in determining species limits. Thus, an analysis, using mitochondrial 16S ribosomal DNA, of the relationship within the genus Tubifex identified six genetically distinct lineages of T. tubifex from North America and Europe that were separated by genetic distances comparable to those found for “welldefined” species of Limnodrilus. Therefore, the existence of several morphologically indistinguishable, thus cryptic, species of Tubifex in North America and Europe was suggested (Beauchamp et al. 2002). Narapidae. This South American family (not included in the morphocladistic analysis) has only one known species, in freshwater (Righi and Varela 1983) (see also 8.2.11.2). Narapa resembles the enchytraeid Achaeta in (always?) lacking chaetae. Phreodrilidae. The 18S gene of the phreodrilid Insulodrilus bifidus provided the first molecular evidence that phreodrilids are closely related to tubificids (Erséus et al. 2002), corroborating previous conclusions based on morphology (e.g. Jamieson 1988b). Phreodrilids, albeit represented only by a single species, formed the sister to a clade consisting of all included Tubificidae and Naididiae. The combined Phreodrilidae + Tubificidae (including Naididae) formed the sister to the Lumbriculidae. Leeches and branchiobdellids were not included in the analyses of Erséus et al. (2002). However, in the later analysis, which included these (Erséus and Källersjö 2003), phreodrilids were the sister-taxon to a clade containing Propappus, Haplotaxis, and a monophyletic tubificid assemblage (Fig. 8.2), though support was inconsistent and may have depended on the questionable coding of gaps as fifth states. Parvidrilidae. This family is known only from Parvidrilus strayeri, described by Erséus (1999) from a stream-bed in northern Alabama, U.S.A., and Parvidrilus spelaeus from subterranean waterbodies of southern Europe (Martinez Ansemil et al. 2002). Parvidrilus has the capilloventrid character of hair chaetae in ventral and dorsal bundles (a symplesiomorphy?). Its affinities have not been investigated in morphocladstic or molecular

#

Reproductive Biology and Phylogeny of Annelida

analyses. The Parvidrilidae have been considered to be most closely related to two, largely Southern Hemisphere, aquatic clitellate families, Capilloventridae and Phreodrilidae, and to have possible relationship to the Narapidae (Erséus 1999). Erséus (2003) suggests that they may be the Northern Hemisphere sister group of the Phreodrilidae. Martinez et al. (2002) endorse the phreodrilid relationship. They consider familial ranking of parvidrilids to be sustained on the basis of three autapomorphies: the location of the chaetae in a markedly posterior position within the segments; the lateral development of the clitellum, with large clitellar cells in relation to the body diameter; and the singular glandular pouches present in the mid-dorsal line of the mesosomial segments. Opistocystidae. This little known family contains Dorydrilus with a few species, in the Americas and Africa (see Stacey and Coates 1996), and Lycodrilus, with five species in Lake Baikal. Lycodrilus was considered to merit a new family, Lycodrilidae, by Chekanovskaya 1962 (see Chekanovskaya 1981) who considered the Opistocystidae to be very closely related to the Naididae. Dorydrilidae. This monogeneric family, with three species, occurs in western Europe. They and the Opistocystidae have not been included in morphocladistic or molecular analyses. The reproductive systems are discussed in 8.2. Lumbriculata. In the morphocladistic analysis (Jamieson 1988b), the Lumbriculata, erected (as a subclass) in that work, contained only the order Lumbriculida, family Lumbriculidae (acanthobdellids, branchiobdellids, and leeches were not included in the analysis). The name Lumbriculata is now available for the lumbriculid + branchiobdellid + acanthobdellid + hirudinid (leech) assemblage that has been confirmed from molecular investigations (Siddall et al. 2001; Jamieson et al. 2002; Erséus and Källersjö 2003; Hugall et al., unpublished; see discussion above). Lumbriculidae. The definition of the Lumbriculidae in Jamieson (1988b) included the following characters: Sperm with weak connectives. Acrosome short. Some male ducts recurved from the funnels to end in the same segment as their testes, i.e. quasiprosoporous. Often multigonadal. Nephridia intersegmentally connected. Myofilaments square in cross section. Some of these characters have been investigated in only a small number of species. For further details of the reproductive system see 8.2. Diplotesticulata. The name Diplotesticulata was coined by Yamaguchi (1953) for all non-microdrile oligochaetes. As a grouping it corresponded with the Opisthopora sensu lato of Michaelsen (1928-1932) (see Michaelsen 1928) which included the non-opisthoporous family Haplotaxidae in which typically each pair of male pores is in fact plesioporous (i.e., in the segment immediately following the corresponding pair of testes) in addition to the ‘megadriles’ which are truly opisthoporous in having the male pores more than one segment behind the most posterior testes. The Opisthopora was retained (Jamieson 1988b) as a more restricted group, excluding haplotaxids, as in this account. The Diplotesticulata, if the Lumbriculidae are excluded,

Non-leech Clitellata

#!

also corresponds with the order Haplotaxida (sensu Jamieson 1978b, 1980). The Diplotesticulata formed a clade in the morphocladistic analysis (Jamieson 1988b) (Figs. 8.3, 8.4A,B) with or without inclusion of the Lumbriculidae. The diplotesticulate condition, connoting two pairs of testes, was recognized as a symplesiomorphy but its use was satisfactorily descriptive in view of the cohesion and monophyly of the group. The group had a single unifying apomorphy: presence of well-developed connectives from the secondary tube to the axial rod of the spermatozoon. However, the Diplotesticulata, although not specifically tested, does not appear to be supported in the molecular analysis of Erséus and Källersjö (2003) (Fig. 8.2) in which the Haplotaxidae grouped with the Propappidae. Haplotaxidea. The superorder Haplotaxidea is redundant as it contained only the Order Haplotaxida sensu stricto, the latter being paraphyletic for the Haplotaxidae and the monotypic Tiguassuidae. There is a trend to loss of the posterior ovaries, a loss which is constant in Tiguassu. This contrasts with loss of the anterior ovaries in the Metagynophora (Jamieson 1988b). Haplotaxidae. From a combination of classical and computer-aided methods Brinkhurst (1988) divided the Haplotaxidae into several genera: Haplotaxis, Alphadrilus, Delaya, Hologynus, Heterochaetella, Pelodrilus and Villiersia. Haplotaxis was retained for the gordioides-like species with specialized pharynx and chaetae associated with a predaceous habit. Adenodrilus was placed in Haplotaxis in Brinkhurst and Jamieson (1971). Tiguassu (reginae) was elevated to the family Tiguassidae Metataxis (eliae) sensu Omodeo was placed incertae sedis in the order Lumbricina (Brinkhurst 1988). Haplotaxis has been regarded by many authors as a representative of a putative octogonadal ancestor of all megadriles (references in Erséus 2003), a view supported in the morphocladstic analysis. However, if the single Haplotaxis species included in the18S analysis of Erséus and Källersjö (2003) was indeed the type-species, H. gordioides, Haplotaxis would appear to be a member of the tubificid clade in which it is closely allied to Propappus (Fig. 8.2). However, support for this relationship was weak as two of the four trees were unresolved and the other two differed from each other. Only when gaps were treated as new states was there greater than 50% jackknife support for the tubificid relationship, but this placed Haplotaxis in a large clade. In the 28S rDNA analysis (Jamieson et al. 2002) the position of Haplotaxis gordioides was unresolved apart from the demonstration that it lies outside the Crassiclitellata. Paraphyly of the genus Haplotaxis, suggested by Gates (1972) and several later workers, was endorsed as the two Haplotaxis species did not group together, one forming the plesiomorph sister group of the lumbriculid through leech clade (the Lumbriculata sensu lato) (Fig. 8.1). Metagynophora. The Metagynophora, then a new coinage, formed a superorder within the Diplotesticulata (Jamieson 1988b). Loss of the anterior ovaries with retention or possibly acquisition of ovaries in segment 13, so that a segment lacking gonads intervenes between the last or only testes and

#" Reproductive Biology and Phylogeny of Annelida the ovaries, diagnoses all megadrile oligochaetes (moniligastrids through the Megascolecidae). This condition is also present in some supposed haplotaxids. This unifying condition, the sole universal synapomorphy of its possessors, has warranted the term Metagynophora (Jamieson 1988b) for this assemblage. Only three species in the entire assemblage are known to have two pairs of ovaries, in segments 12 and 13. These are the almid Glyphidrilus kukenthali, the glossoscolecid Enantiodrilus borellii, and the megascolecid Diplocardia sandersi (see Jamieson 1978b, 1980). Where seen in megadriles, this rare, quadriovarian condition presumably represents a reversal to, rather than retention of, the presumed plesiomorphic condition but has an interesting implication of retention of the underlying genome, or of an epigenetic switch, for production of ovaries in both segments. The metagynophoran clade has not been tested by molecular means. Moniligastrida. The plesiomorphic sister-group of the Opisthopora contains only the family Moniligastridae. For nomenclatural purposes these were placed (Jamieson 1988b) in an order Moniligastrida (an arbitrary ranking) coordinate with the order Opisthopora. Moniligastrids are indigenous in southeast and eastern Asia. They are terrestrial and, despite a common belief that they are amphibious, no moniligastrids are known to be aquatic or limicolous (Gates 1972). They have yet to be subjected to molecular analysis. Reproductive characters are discussed in 8.2. Opisthopora. Nested immediately within the Metagynophora is the alluroidid through megascolecid assemblage (Jamieson 1988b). The nominal apomorphy of the Opisthopora is movement of the male pores from the plesiomorphic location one segment behind the last or only testes (the plesioporous condition) to a more posterior location, commencing with segment 13 or its homeotic equivalent. The name Opisthopora was coined by Michaelsen (Nachtrag 1932 in Michaelsen 1928) for this condition though for reasons which are not clear he included the plesioporous family Haplotaxidae (= Phreoryctidae), which are excluded here and in Jamieson (1988b). In one known haplotaxid, Haplotaxis violaceus, the anterior vasa deferentia traverse more than one segment, discharging near the posterior pair of vasa in segment 12 but the latter ducts remain plesioporous. Remarkably, the validity of the Opisthopora has not been tested by molecular phylogenetics, as its most plesiomorphic family, the Alluroididae, has not been sequenced. However, the largest opisthopore group, the Crassiclitellata, is strongly confirmed as monophyletic (Fig. 8.1, 8.2) (Jamieson et al. 2002; Erséus and Källersjö 2003). These, at least, exclude Haplotaxis. Alluroidina. Alluroidoidea. These have opisthoporous male pores in segment 13. The Alluroidoidea of Brinkhurst and Jamieson (1971) computed in the morphocladistic analysis (Jamieson 1988b) as the plesiomorphic sister-taxon of the Crassiclitellata. It contains the Syngenodrilidae Smith and Green 1919, and the Alluroididae Michaelsen 1900, neither of which has been included in molecular analyses. Their reproductive systems are discussed in 8.2.

Non-leech Clitellata

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Alluroididae. The Alluroididae have been reviewed by Brinkhurst (1964), Jamieson (1968b, 1971a) and Omodeo (1996). They occur in inland waters of equatorial Africa and South America but have also been found (Jamieson 1968b) in moist soil near streams. Syngenodrilidae. No specimens of the East African family Syngenodrilidae have been found since definition of the family for Syngenodrilus lamuensis by Smith and Green (1919). The habitat of Syngenodrilus is unknown. Crassiclitellata. The remainder of the Oligochaeta are unified by a striking synapomorphy, a multilayered clitellum. This group was termed (Jamieson 1988b) the Crassiclitellata. The Crassiclitellata is confirmed as a monophylum in the molecular analyses (Jamieson 2000; Jamieson et al. 2002; Hugall et al., unpublished), a result which is not contradicted by 18S analysis (Siddall et al. 2001; Erséus and Källersjö 2003). Thus, at least for the taxa represented, the multilayered clitellum arose only once. It is noteworthy that in the morphocladistic analysis Crassiclitellata remained a monophyletic group even when the character pertaining to the clitellum was omitted from the analysis (Jamieson 1988b). The maximum parsimony majority rule consensus tree for 549bp of 28S only (Fig. 8.1) gives 100% bootstrap support for the Crassiclitellata versus outgroup taxa. Maximum likelihood analysis differs little from a maximum parsimony bootstrap tree. Aquamegadrili and Terrimegadrili. Some molecular analysis was made (Jamieson 2000; Jamieson et al. 2002) of division of the Crassiclitellata on morphocladistic evidence into two groups, the Aquamegadrili and Terrimegadrili (Jamieson 1988b). Aquamegadrili have an aquatic or semiaquatic mode of life, and include the families Sparganophilidae (Holarctic), Biwadrilidae (Japan), Almidae (mostly warm tropics but including Criodrilus, in the Mediterranean region etc.) and Lutodrilidae (Southern Neartic). It is not unlikely that the aquamegadrile families, irrespective of mono- or polyphyly of the group, have always had an aquatic or amphibious existence. The remainder of the Crassiclitellata were predominantly terrestrial, hence the term Terrimegadrili. These included the superfamilies Lumbricoidea, Eudriloidea and Megascolecoidea. The 28S analyses (Jamieson et al. 2002) are inconsistent with partition of crassiclitellates into Aquamegadrili and Terrimegadrili and with monophyly of the Aquamegadrili but this is entirely due to the inclusion of Komarekiona as sister-taxon of Sparganophilus. This pair was previously supported in a phenetic analysis, though in a phylogenetic analysis Komarekiona had the terrimegadrile Microchaetus as its nearest neighbor (Jamieson 1978b); it was also supported in a cladistic phylogeny by Omodeo (1998). Qiu and Bouché (1998) consider the Komarekionidae to be the junior synonym of the terricole family Ailoscolecidae, a relationship mooted by Jamieson (1978b). The representatives (Sparganophilus, Criodrilus and Lutodrilus) of the original aquamegadrile taxa, with Komarekiona, lie within a paraphyletic Terrimegadrili (Figs. 8.1 and 8.5). The superfamilies originally

#$ Reproductive Biology and Phylogeny of Annelida constituting the Aquamegadrili may, nevertheless, be retained, though where monofamilial they are possibly redundant. Michaelsen (1932) divided what are here termed the Crassiclitellata into two groups: the Lumbricina and Megascolecina. The 28S rDNA analysis (Jamieson et al. 2002) (Fig. 8.1) indicates a monophyletic core for each of the groups Megascolecina (Megascolecidae and Ocnerodrilidiae) and Lumbricina (Sparganophilidae, Komarekionidae, Almidae (Criodrilus lacuum), Lutodrilidae, Hormogastridae, Lumbricidae, and Microchaetidae) but both groups are rendered paraphyletic by the successsive basal taxa Glossoscolecid sp. + Eudrilus (Eudrilidae) and Pontoscolex (Glossoscolecidae). Sparganophiloidea. This contained only the Sparganophilidae in the morphocladistic analysis (Jamieson 1978b, 1988b). Sparganophilidae. This is a Nearctic freshwater family, with representation, possibly by introduction, in the United Kingdom and France. The type-species, Sparganophilus tamesis Benham (Fig. 8.40) was shown to be the senior synonym of the American S. eiseni Smith by Jamieson (1971c). The Komarekionidae and Ailoscolecidae were separated from the Sparganophilidae, in the Terrimegardili, in the morphocladistic analyses (Jamieson 1978b, 1988b) but the 28S rDNA analysis suggests a very close relationship of Sparganophilus and Komarekiona. Evidence for inclusion of the Ailoscolecidae with these is given in the discussion of reproductive systems in 8.2.11.4. Komarekionidae. This family is known from a single, terrestrial species, Komarekiona eatoni Gates 1974, from North America (Sims 1980, 1982) included Komarekiona in the Ailoscolecidae. Komarekiona seems little advanced beyond a sparganophilid level of organization. It is therefore noteworthy that Komaerkiona formed a clade with Sparganophilus, with very high bootstrap support, in molecular analyses (Jamieson et al. 2002) (Figs. 8.1, 8.6). Ailoscolex has yet to be included in molecular analyses. Ailoscolecidae. The single genus, Ailoscolex, occurs terrestrially in southwestern France. Ailoscolex appears to have close affinities with the family Komarekionidae (see 8.2), which was subsumed in it by Sims (1980, 1982), and with the Sparganophilidae. Biwadriloidea. Biwadrilidae. Criodrilus bathybates Stepenson (synonym C. miyashitai Nagase and Nomura) was made the type of the monotypic Biwadrilus, in the subfamily Biwadrilinae, by Jamieson (1971c). As the family Biwadrilidae it was placed in a superfamily Biwadriloidea by Jamieson (1988b) which formed the plesiomorph sister-taxon of the Almidae + Lutodrilidae. Biwadrilus has yet to be subjected to molecular analysis that might clarify its relationships. Omodeo (2000) suggests that Biwadrilus (removed from Criodrilus by Jamieson 1971c) may belong to his resurrected Criodrilidae Superfamily Almoidea. As consituted in the morphocladistic analysis these contained the Almidae, Lutodrilidae and tentatively the Kynotidae. Almidae (including Criodrilus). The Almidae Duboscq was resurrected as the Alminae (containing Alma, Callidrilus, Glyphidrilus, Drilocrius,

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Glyphidrilocrius and Criodrilus) within a broad Glossoscolecidae by Jamieson (1971c) who argued against placement of its species in the Microchaetidae by Gates (1972). An innovation was the inclusion of Criodrilus, in a tribe Criodriliini, within the Alminae. The Alminae was subsequently restored to familial rank (Jamieson 1988b). The family is aquatic, falling within the Aquamegadrili and occurs in Tropical Africa and the Nile Valley; Central and South America; India; Oriental Region. Criodrilus occurs in Europe and the Middle East. In the 28S rDNA analysis the sole representative, Criodrilus lacuum (Criodrilinae) was the sister-taxon of Sparganophilus + Komarekiona (Fig. 8.1). There is a need for inclusion of members of the Alminae, particularly Alma, in molecular analyses. Omodeo (2000) has restricted the Almidae to Alma (the other almine genera were placed by him in a family Glyphidrilidae) but it is here considered that the restricted Almidae is insufficiently distinct in comparison with the differences between other families. Furthermore, Drilocrius alfari (Fig. 8.38C) which resembles Alma in having claspers, though with male pores basal to, not apical on these, constitutes a convincing link with the Almidae sensu lato. Pop et al. (2002) gave a maximum parsimony tree for 15 lumbricid species and Criodrilus lacuum based on 16S rDNA and for 15 species based on CO1 analysis, both cladograms being rooted with a mollusc species. Agreement between the two trees was low. Criodrilus necessarily associated with lumbricids as only these accompanied it in the analyses. They nevertheless considered that “separation [of Criodrilus lacuum] from the Lumbricidae to the family Criodrilidae or Glossoscolecidae (Oligochaeta) is supported by these genetic characters”. Inclusion of Alma in an analysis will be necessary to test the present author’s contention that the affinities of C. lacuum lie with the Almidae. Lutodrilidae. The family Lutodrilidae is known form a single species of ‘swamp worm’ in Louisiana (McMahan 1976, 1979). Morphocladistic analysis suggests that the Lutodrilidae are closely related to the Almidae (Jamieson 1988b). In the 28S rDNA analysis they lie in the same clade as the almid Criodrilus (in the absence of Alma from the study) but do not show an unequivocal close relationship with this (Fig. 8.1). Kynotidae. The Kynotidae is a monogeneric, terrestrial family with about 12 species, restricted to Madagascar. The phylogenetic position of the Kynotidae is uncertain. In morphocladistic analysis it was questionably the sister-group of the Almoidea (Jamieson 1988b) (Fig. 8.4A,B). It had been considered the sister-group of the Lumbricoidea in an intuitive phylogeny (Jamieson 1971b) and maintained this position in a previous Hennigian study (Jamieson 1978b). Glossoscolecidae. The Glossoscolecidae forms the dominant group of earthworms in tropical South America north of the Juramento-Salado River (to the south of which acanthodriles predominate) and much of the Caribbean. They occur in a wide range of terrestrial habitats although most species are forest dwellers. Pontoscolex corethrurus is circummundane in warmer regions of the world. One of the largest species (Rhinodrilus fafner)

#& Reproductive Biology and Phylogeny of Annelida has been reported to reach over 2 m in length and 2.5 cm in diameter (Sims 1982). The tribe Glossoscolecini sensu (Jamieson 1971c) is usually, as here, given familial rank (Righi , in many papers; Gates 1972; Sims 1980, 1982; Jamieson 1988b; Jamieson et al. 2002). The family appeared paraphyletic in molecular analyses (Jamieson 2000; Jamieson et al. 2002) (Fig. 8.1) and included Eudrilus but this relationship requires further testing. Samuel James (pers. com.) has independently noted the similarity of Eudrilus to glossoscolecids in morphological studies. Eudrilidae. Eudrilids are restricted to Africa (Ethiopian Region) as natives. One species, Eudrilus eugeniae, has become widely distributed around the warmer parts of the world and is cultured as the ‘African Night Crawler’. The male pores of eudrilids lie in segment 17, as is also typical of Ocnerodrilidae, with which they have been considered to be closely related. Molecular studies do not support this relationship but suggest glossoscolecid affinities (Jamieson 2000; Jamieson et al. 2002). The Eudrilidae has formerly been associated with the Megascolecidae and Ocnerodrilidae in a superfamily Megascolecoidea (Jamieson 1978b) or in a separate subfamily Eudriloidea as the sister-group of the Lumbricoidea + Megascolecoidea sensu stricto, in the morphocladistic analysis (Jamieson 1988b) (Figs. 8.3, 8.4 A,B). In the molecular analyses (Jamieson 2000; Jamieson et al. 2002) (Figs. 8.1, 8.6), Eudrilus always has a glossoscolecid sp. (a lumbricoid sensu Jamieson 1978b) as its sister-taxon, with moderate bootstrap support. The other glossoscolecid exemplar, Pontoscolex, may or may not link with these. There is no molecular support in these analyses for regarding eudrilids as the unique sister-group of the Ocnerodrilidae + Megascolecidae assemblage. The molecular placement of the Eudrilidae with the Glossoscolecidae may be correct but it requires further investigation as there are morphological similarities between this family and the Ocnerodrilidae with which both a cladistic and phenetic analysis linked them in a previous study (Jamieson 1978b); furthermore, ocnerodrilid and eudrilid interchaetal ratios are closely, and uniquely, similar (Jamieson unpublished). Omodeo (2000) recognized the Eudriloidea (as did Jamieson 1988b); he derived eudrilids independently (from alluroidids), thus also noting their distinctness, but that origin goes contrary to the present confirmation of the monophyletic nature of the Crassiclitellata. Lumbricoidea. The Lumbricoidea as defined by Jamieson (1978b, 1988b) included the Lumbricidae (Holarctic), Glossoscolecidae (Neotropical), Microchaetidae (Ethiopian, south of the Kalahari), Hormogastridae (Western Palaearctic, Tyrrhenian), Komarekionidae (Nearctic), and Ailoscolecidae (Palaearctic), of which the latter two families are here placed in the Sparganophiloidea. Relationships of some of these families and the validity of the grouping Lumbricoidea were investigated from molecular sequences by Jamieson (2000) and Jamieson et al. (2002) (Figs. 8.1, 8.6). The Lumbricoidea is incompatible with the 28S data (Fig. 8.1) or combined gene data (Fig. 8.6) but principally because the glossoscolecid sp.

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never groups with the remainder of the lumbricoids in the best trees but groups equivocally with Eudrilus which is here tentatively placed in the resurrected superfamily Glossoscolecoidea. However, testing the unity of the majority of the remaining Lumbricoidea will necessarily require more sampling (Jamieson et al. 2002). In these molecular analyses the Lumbricoidea is a paraphyletic clade, though with considerable monophyly (Sparganophilus, Komarekiona, Criodrilus, Lutodrilus, Hormogaster, lumbricids, and the microchaetid forming a clade). This paraphyletic lumbricoid assemblage also contains the Ocnerodrilidae and Megascolecoidae in addition to the Eudrilidae. In the revised classification (Table 8.2) the Lumbricoidea sensu Jamieson (1988b) is provisionally revived for the non-megascolecoid families, with the exception that the Eudriloidea is tentatively subsumed in the Glossoscolecoidea, as indicated by the molecular analyses. In Table 8.2, lumbricoid families are listed in the order that they occur in the DNA analyses (Figs. 8.1, 8.3). Microchaetidae. Microchaetidae are restricted to South Africa and include the largest known earthworms, Microchaetus microchaetus, with a maximum reported length of 7 m and diameter of 7.5 cm. Molecular analysis (Jamieson 2000; Jamieson et al. 2002) (Fig. 8.1) indicate that microchaetids lie in a different clade from the Glossoscolecidae in which they were formerly placed as a tribe (Jamieson 1971c). With the Hormogastridae they occupy a sparganophiloid + almoid + lumbricoid clade (Fig. 8.1). Lumbricidae. Lumbricids are native in the Holarctic, chiefly in Europe. From the 28S analysis they share a clade with the Sparganophilidae, Komarekionidae, Criodrilus lacuum (an atypical examplar for the Almidae), Lutodrilidae and Hormogastridae (Fig. 8.1). Hormogastridae. Hormogastrids are known from two subfamilies that occur mainly around the western Mediterranean. Molecular analyses (Jamieson 2000; Jamieson et al. 2002) (Fig. 8.1) indicate that hormogastrids lie in a different clade from the Glossoscolecidae in which they were formerly placed as a subfamily (Jamieson 1971c). They show somewhat unresolved relationships with the Sparganophilidae, Komarekionidae, Almidae (Criodrilus), Lutodrilidae and Lumbricidae on molecular grounds (Fig. 8.1). Hormogastrids were considered the sister taxon of the Lumbricidae by Rota and Omodeo (1992) and Omodeo (2000). Tumakidae. In the Colombian family Tumakidae, monotypic for Tumak hammeni Righi 1995, the calciferous glands closely resemble those of some Glossoscolecidae but the gizzard instead of being preclitellar, is postclitellar, in segment 25. As noted by Omodeo (2000), in accordance with the ‘rule’ of Bouché (1972) the gizzard thus lies outside the clitellar zone. The calciferous gland, at 20/21, lies within the posterior limit of the clitellum. Tumak differs from the Glossoscolecidae in the posterior and reverse position of its calciferous glands and gizzard, intraparietal male ducts and presence of dorsal pores. Despite these differences, it seems likely that Tumak originated

$ Reproductive Biology and Phylogeny of Annelida Table 8.2. A revised, simplified classification of the Clitellata

Clitellata Michaelsen 1919 Ì Eoclitellata1- Capilloventridae Harman and Loden 1984 DNA Ì Neoclitellata2 - all other Clitellata Ë Unnamed clade, here unresolved ¥ Randiellata? Jamieson 1988b Randiellidae Erséus and Strehlow 1986 ¥ Tubificata Jamieson 1988b Tubificidae Vejdovsky 1884 DNA (including Naidinae Ehrenberg 1831 DNA) Narapidae? Righi 1983 Opistocystidae? Dorydrilidae? Cook 1971 Parvidrilidae Erséus 1999 Phreodrilidae Beddard 1891DNA Propappidae Coates 1986 DNA Haplotaxidae? Michaelsen 1900 DNA Tiguassuidae? Brinkhurst 1988 Ë Unnamed clade Ø Lumbriculata Jamieson 1988b q Lumbriculidae Vejdovsky 1884 DNA q Unnamed clade “ Branchiobdellida Odier 1823 DNA3 “ Hirudinea Lamarck 1818 DNA Acanthobdellida Livanow 1905 DNA Hirudinida sensu Siddall (Euhirudinea Lukin 1956) Ø Unnamed clade Enchytraeidae Vejdovsky 1879 DNA4 Metagynophora Jamieson 1988b z Moniligastridae Claus 1880 z Opisthopora Michaelsen 1932 Y Alluroidina Jamieson 1978b Alluroididae Michaelsen 1900 Syngenodrilidae Smith and Green 1919 DNA Y Crassiclitellata Jamieson 1988 ­ Paraphyetic, requiring resolution Biwadriloidea Jamieson 1978b Biwadrilidae Jamieson 1971 Glossoscolecoidea Emend. Glossoscolecidae Michaelsen 1900 DNA Eudrilidae Claus 1880DNA? Tumakidae Righi 1995 Sparganophiloidea Jamieson 1978b5 Ailoscolecidae Bouché 1969 Komarekionidae Gates 1974 DNA Sparganophilidae Michaelsen 1918 DNA Lumbricoidea Gates 1976. Emend. Microchaetidae Michaelsen 1900 DNA Lumbricidae Claus 1876 DNA Table 8.2 contd

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Table 8.2 contd

Kynotidae? Jamieson 1971 Hormogastridae Michaelsen 1900 DNA Almoidea Jamieson 1978b6 Lutodrilidae McMahan 1978 DNA Almidae Duboscq 1902 (including Criodrilus DNA) ­ Megascolecoidea Jamieson 1978b Ocnerodrilidae Beddard 1891 DNA Megascolecidae Rosa 1891 DNA Acanthodrilinae Vejdovsky 1884 DNA Megascolecinae Rosa 1891 DNA 1

Eoclitellata, new taxon, is defined as Clitellata with capillary setae in the dorsal and ventral bundles. Atria, prostates and penes are absent 2 Neoclitellata, new taxon, is defined as Clitellata lacking capillary setae in the ventral bundles or if these are present (Parvidrilidae) they are markedly anterior in their segments, a peculiar lateral clitellum is present and glandular pouches are present in the mesosomial segments. 3 Whether the Acanthobdellida or the Branchiobdellida is nearest to Hirudinida is equivocal from molecular evidence. Neither shows the anterior extension of the acrosome tube diagnostic of Hirudinida. The single mitochondrion is here considered a clear synapomorphy of Acanthobdellida and Hirudinida, contrasting with the plesiomorphic multiple mitochondria of the sperm of branchiobdellids and oligochaetes sensu stricto. However, the retention of some setae in in Acanthobdella but not in branchiobdellids is plesiomorphic. 4 The phylogenetic position of the Enchytraeidae is particulary controversial (see text). 5 The close grouping of the Sparganophilus and Komarekiona in the molecular analyses prompts extension of the subfamily to include the Komarekionidae and, because of its morphological resemblance to the latter, the Ailoscolecidae. 6 Only Lutodrilus and Criodrilus were included in the molecular analysis. They lay within the ‘lumbricoid’ clade and were paraphyletic relative to each other. It is nevertheless envisaged that the Lutodrilidae and Almidae are closely related. The position of Criodrilus , which grouped with Alma in the morphocladistic analysis (Jamieson 1988b) remains controversial.

within the Glossocolecidae or, at most, is its sister-taxon and separate familial rank is questionable. Ocnerodriloidea and Megascolecoidea. The families Megascolecidae, Ocnerodrilidae and Eudrilidae were tentatively included in the superfamily Megascolecoidea in Jamieson (1978b, 1980). However, in the morphocladistic analysis (Jamieson 1988b) (Fig. 8.6A,B), the Eudrilidae (superfamily Eudriloidea) and especially the Ocnerodrilidae (superfamily Ocnerodriloidea) occupied a basal position relative to the other terrimegadrile families. Molecular data (Jamieson 2000; Jamieson et al. 2002) do not support this distant position for ocnerodrilids but reunite them with the Megascolecidae (Figs. 8.1, 8.6). As we have seen, the single eudrilid included in the molecular analysis associates with the Glossoscolecidae and is not confirmed as a sister taxon of the Ocnerodrilidae + Megascolecidae. Megascolecidae. The largest, most speciose, earthworm family is the Megascolecidae for which a Pangean origin has been suggested (Jamieson 1981a). They are native in the Nearctic, Ethiopian, Oriental, Australian, eastern Palaearctic (China, Japan, Korea) and southern Neotropical Regions,

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Reproductive Biology and Phylogeny of Annelida

Colour Figure

Fig. 8.6. Strict consensus of six maximum parsimony trees using the combined data. Taxa with one, two or three genes indicated. Heuristic search, 1371 sites unweighted 50 random additions tree bisection reconnection with steepest decent, with bootstrap consensus values from 500 resamplings (heuristic search, unweighted, five random additions, tree bisection reconnection with steepest descent). 1, 28S (12S only for Rhododrilus glandifera Jamieson 1995); 2, 28S + 12S; 3, 28S + 12S + 16S. The narrow colored column indicates the attribution of species which would conform with the classification of Gates (1959, 1972); note the extreme polyphyly of the Acanthodrilidae and Megascolecidae sensu Gates. The Octochaetidae sensu Gates represented (with two or three pairs of prostates) are poly- or paraphyletic and lie within the Acanthodrilinae sensu Jamieson. Other ‘octochaetids’ differing in having megascolecin male pores (not included in this analysis, such as Celeriella) would not be expected to group within the Acanthodrilinae. Redrawn after Jamieson et al. 2002 Zoosystema 24(4): 707-734, Fig. 3.  Publications Scientifiques du Muséum national d’Histoire naturelle, Paris.

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with Central America. In S. America North of the Juramento-Salado River the large family Glossoscolecidae largely replaces the Megascolecidae. In the Ethiopian Region, particularly in Tropical West and East Africa, the family Eudrilidae greatly exceeds megascolecids in numbers of genera. Currently recognized subfamilies of the Megascolecidae are the Acanthodrilinae and Megascolecinae, with or (as supported here) without the Ocnerodrilinae. Megascolecidae sensu stricto differ from the Ocnerodrilidae in that calciferous glands, if present, are not in, or are not restricted to, segments 9 and 10; their reproductive systems are discussed in Section 8.2. Indigenous acanthodriles are predominant in the earthworm faunas of the southern and eastern portions of North America, Mexico, Guatemala, southern S. America, S. Africa, New Zealand, New Caledonia, and parts of Australia (especially the Northern Territory and Queensland). The native range of the Ocnerodrilidae includes the warmer parts of N. and S. America, the Dominican Republic, Africa, India and Burma (Jamieson 1981a). Several, sometimes widely divergent classifications of megascolecoid earthworms have been proposed since publication of Stephenson’s monograph (1930) of the Oligochaeta (see Michaelsen 1928-1932; Pickford 1937; Omodeo 1958; Gates 1959; Lee 1959; Sims 1966, 1967; Jamieson 1971d, 1971e, 1971f, 1978b, 1988b). Particular attention has been paid to the system of Gates (1959), supported by Sims (1966, 1967), and the alternative classification of Jamieson (1971d, e, f) as both of these systems are widely used. Detailed discussion of the alternative classifications may be found in the 1971 papers. Analysis of 28s rDNA (Fig. 8.1), complemented by 12s and 16s mitochondrial rDNA (Fig. 8.6), of megascolecoid oligochaete sequences, has been performed using maximum parsimony, maximum likelihood, bootstrap and neighbor joining procedures, with one (Lumbricidae) to nine nonmegascolecoid families as the outgroup (Jamieson 2000; Jamieson et al. 2002). The widely used classification by Gates (1959) of the non-ocnerodriline Megascolecidae into three groups (Acanthodrilidae, with tubular prostates and holonephridia; Octochaetidae, with tubular prostates and meronephridia; and Megascolecidae, with racemose prostates and holo- or mero-nephry) was not supported by the molecular data. Thus the Octochaetidae and Acanthodrilidae, so defined, were not endorsed as Neodiplotrema (Octochaetidae sensu Gates) paired unequivocally with Diplotrema (a true acanthodriline), and Dichogaster (Octochaetidae sensu Gates) grouped with these. Diagnosis of Megascolecidae by racemose prostates is invalid, as Amynthas, Begemius, Didymogaster, Digaster, Heteroporodrilus, Propheretima, Spenceriella, and Perionyx (all with racemose prostates and Megascolecidae sensu Gates) did not collectively form an exclusive monophylum. The Ocnerodrilidae, represented by Eukerria, was sister-group of the Acanthodrilinae + Megascolecinae, and joint monophyly of the three taxa was supported (Figs. 8.1, 8.6, 8.7). If the Ocnerodrilidae are regarded as a family, the Acanthodrilinae and Megascolecinae are subfamilial in the Megascolecidae.

$" Reproductive Biology and Phylogeny of Annelida Although Gates’ (1959) classification was not supported, the alternative classification of Jamieson (1971d, e, f) also required modification in the light of the molecular study. Thus subdivision of the Megascolecinae into the tribes Perionychini, Dichogastrini, and Megascolecini (long suspected of being grades rather than clades) did not have molecular support (Figs. 8.1, 8.6, 8.7). The Perionychini proved to be a para- and polyphyletic assemblage. The

Fig. 8.7. Maximum Likelihood tree from the sum of the 28S rDNA and mtDNA partitions optimized for GTR-G model separately (SUM model best tree). Selected from 14586 near parsimonious reverse constraints trees. The Maximum Parsimony bootstrap tree is the same. Dashed lines indicate the two topology differences found in the combined data model Maximum Likelihood tree (COMBO model). Bold letters are clade labels for Table 4 in Jamieson et al. 2002. Maximum Parsimony and Maximum Likelihood boostrap values above and below branches respectively. Partition support shown in Table 4. From Jamieson, B. G. M. et al. 2002. Zoosystema 24(4): 707-734, Fig. 4.  Publications Scientifiques du Muséum national d’Histoire naturelle, Paris.

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Dichogastrini were disbanded as Dichogaster associated with the Acanthodrilinae. Maximum likelihood testing confirmed non-monophyly of the Dichogastrini. This confirmed the observation from an earlier cladistic study (Jamieson 1978b) that “the diphyletic origin of the Dichogastrini emerges clearly.” It is uncertain that the residue of the Dichogastrini, represented by Digaster and Didymogaster, with male pores on segment 18, is monophyletic. Jamieson (2000) referred to them as Megascolecinae with the notoscolecin arrangement of nephridia (the Notoscolex-group). The Megascolecini (defined by meronephridia differing from those of notoscolecins) were not found to be monophyletic. Recognition of further higher groupings and confirmation of genera awaits a more detailed survey.

8.1.5

Spermatozoal Phylogeny

Jamieson (1983a) compared spermatozoal ultrastructure of a range of oligochaetes with a holomorphological phylogeny of the group. Plesiomorph characters for the oligochaete spermatozoon were proposed. The chief trends from these plesiomorphies were deduced to have been elongation of the acrosome and its tube; withdrawal of the acrosome vesicle and the axial rod into the acrosome tube and development of a knob-like tip on the rod (capitulum); development of connectives from the secondary tube to the axial rod (though some possibility was noted that the reverse, absence of connectives, is plesiomorph); detorting and shortening of the midpiece (or possibly, again, the reverse) with an increase in numbers of mitochondria from the plesiomorph four to eight or a reduction to two (though capilloventrids suggest that the multiple mitochondria might be plesiomorphic); modification of the base of the tube to form a limen of variable form; and, in one line (lumbricids) flattening of the tip of the nucleus and correspondingly of the limen. Sperm ultrastructure, examined for 9 oligochaete families, corresponded well with taxonomic and phylogenetic groupings. However, convergent similarity of the phreodrilid sperm to that of the Lumbricina suggested a corresponding alteration of fertilization biology in the phreodrilids. The results indicated that the Haplotaxidae were the plesiomorph sister-group of the opisthopores, though they did not unequivocally contraindicate acceptance of a Haplotaxis-like form as a stem form of the Haplotaxida (opisthopores and Haplotaxidae) and Tubificida (Jamieson 1983a). In a parsimony analysis of spermatozoal ultrastructure, Jamieson et al. (1987) drew the following conclusions as to relationships within oligochaetes. The Haplotaxida (Haplotaxis through Megascolecidae) were found to form a discrete monophyletic group with inclusion, however, of the reproductively exceptional tubificidan Phreodrilus. Bythonomus, representing the Lumbriculidae, the phylogenetic position of which had been so controversial, formed the plesiomorphic sister group of the Haplotaxida, a position which was supported from other evidence. The Tubificida, represented by two tubificids (Rhizodrilus and Limnodriloides) and by the enchytraeid Lumbricillus, appeared plesiomorphic relative to the

$$ Reproductive Biology and Phylogeny of Annelida lumbriculid + haplotaxid assemblage and to lie at the base of the tree, but all three appeared mutually paraphyletic. Monophyly of the Tubificida could not, however, be considered conclusively refuted from the small sample used. Lumbricillus (Enchytraeidae) appeared to have the most pleisomorphic sperm in the investigated oligochaetes. The Megascolecidae (Amynthas and Fletcherodrilus) formed the highest affinity and most apomorphic group. The implications of relative apomorphy of the Lumbriculida (Bythonomus) were considered profound. It was stated that ‘Branchiobdellids and leeches are generally regarded as sharing a common ancestry with lumbriculids and would, because of the revised position of the latter, cladistically constitute part of the Oligochaeta sensu lato’ (Jamieson et al. 1987). A major example of the utility of spermatozoal ultrastructure for testing phylogenetic relationsips, in this case at the species level, is that for Enchytraeus by Westheide et al. (1991). Species of Enchytraeus are morphologically rather similar; they are difficult to distinguish or even inseparable on conventional light microscopic methods. In individuals from 19 populations of different origin and in some cases unknown species identity the ultrastructure of mature spermatozoa was investigated. Their morphometric and qualitative data were used as morphologic taxonomic characters. With the exception of a genetically polymorphic field population of E. albidus, the material investigated came from genetically monomorphic laboratory inbred strains. The structure of the thread-like spermatozoa generally conformed with the sperm type known for the family. Numerical characters used were 1) length of the acrosome, 2) length of the corkscrew-like nucleus, 3) number of helical spires of the nuclear flange, 4) periodicity of the flange spires proximally and distally, 5) angle of the flange to the longitudinal axis of the nucleus, 6) length of the midpiece, 7) number of the mitochondrial helical spires, and 8) length of the flagellum. Besides the shape of the nucleus, the complex structure of the acrosome provided further specific qualitative differences such as, length ratio of primary acrosome vesicle to the entire acrosome, extension of a subdistal electron-dense material, and absence or presence of specific structures underneath the primary acrosome vesicle. Eight populations showed highly significant differences from each other and from the remaining populations on the basis of morphometric data alone. In addition, these populations—recognizable as different species on conventional methods—could also be discriminated by qualitative differences of their spermatozoa. The spermatozoa of 10 other populations appeared to be highly similar on the length of their nuclei. Number of nuclear spires, length of acrosome, flange spire periodicity and especially structural details of the acrosome, however, separated these 10 populations into two significantly different groups, one with six species and the other with four. The group of four, which had also been recognized in parallel, independently conducted non-morphological investigations, was considered to belong to one species. Certain morphological data divided the group of six into two groups of three, as was also supported by non-morphological methods and crossbreeding experiments—both groups being distinct species. Statistical

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%$comparisons generally showed no or only slight variability of morphometric data for sperm of one individual. The spermatozoal variability between individuals of genetically monomorphic laboratory cultures was higher but in no case did it prevent species identification. The greatest intraspecific variation was observed in an E. albidus field population: spermatozoa of individuals differing in the pattern of single enzymes deviated significantly in their morphometric data, and on the basis of these data alone they were inseparable from spermatozoa of certain other species. However, qualitative spermatozoal characters were not affected and species could easily be discriminated on the basis of acrosomal structure (Westheide et al. 1991). The influence of patterns of spermatozoal ultrastructure on hypotheses of phylogenetic relationships within the Tubificidae was examined by Erséus and Ferraguti (1995) for species representing 15 different genera. A parsimony analysis of a combination of spermatozoal and conventional morphological characters supported that the Phallodrilinae, Limnodriloidinae and Tubificinae are monophyletic taxa, and that the Rhyacodrilinae as currently defined is a paraphyletic group (Erséus and Ferraguti 1995). The sperm ultrastructure of three Naididae (Paranais frici, P. litoralis and Stylaria lacustris) and two Lumbriculidae (Rhynchelmis brachycephala and R. alyonae) was decribed to supply spermatological evidence towards a better understanding of the phylogeny of the two families. Naidid spermatozoa were found to be similar to one another, particularly with regard to the acrosome (with the acrosome tube bent to one side), and the shape and number of mitochondria. The various morphological characters did not militate against the previously proposed inclusion of naidids within the Tubificidae. The two lumbriculid species examined showed a characteristic conical indentation at the apex of the nucleus, a feature formerly described only in Branchiobdellida, recognized as having possible affinities with Lumbriculidae (Ferraguti et al. 1999). Presence of an acrosome tube apears to be distinctive of clitellates (Jamieson 1981c). A structure resembling an acrosome tube was recognized in vestimentiferan pogonophorans by Jamieson (1987b); however, this has since been found to differ from that of the oligochaete sperm in having continuity with the nucleus although it does not show nuclear (DAPI) staining (Ferraguti and Bright, pers. comm.). Features of oligochaete sperm shared with onychophorans (see Jamieson 1986) are: a cylindrical midpiece interpolated between the basal body and the nucleus; presence of subacrosomal material possibly homologous with an acrosome tube; location of the sole, distal, centriole (basal body) behind the midpiece; and absence of a proximal centriole at maturity. However, an onychophoran-clitellate sistergroup relationship is not widely supported.

8.1.6 Revised Classification of the Clitellata The following revised classification of the Clitellata, with particular reference to oligochaetous clitellates is based chiefly on the morphocladistic

$& Reproductive Biology and Phylogeny of Annelida classifications (Jamieson 1978b,1988b) and the classification of Brinkhurst and Jamieson (1971), modified according to molecular analyses discussed above. Sister-groups are indicated by similar symbols.

8.2 ANATOMY AND EVOLUTION OF THE OLIGOCHAETE REPRODUCTIVE SYSTEM Before considering the anatomy of the reproductive system of individual families, some discussion of shared features, such as the gonoducts, clitellum and male pores will be given.

8.2.1 Gonoducts Oligochaete gonoducts are coelomoducts but otherwise the genital ducts and associated glands in oligochaetes offer no clear homologies with those of polychaetes (see Westheide 1988). Retention of separate coelomoducts and metanephridia, universal for oligochaetes, has been reported only in capitellid and some nereidid polychaetes. Oligochaete features of few gonadal segments (basically apparently two testicular followed by two ovarian segments), and presence of spermathecae and a clitellum are seen in questid polychaetes. From a consideration of spermatozoal ultrastructure resemblances between questids and oligochaetes were considered to be homoplastic convergences (Jamieson 1983b) and this is confirmed from analysis of 18S sequences (Erséus et al. 2002; see also Siddall et al. pers. comm. 8.1.1). Oligochaetes are hermaphroditic but some reports of protandry exist. A tubificid, Mitinokuidrilus excavatus, has been described in which the sexes are separate, though whether this condition represents consecutive hermaphroditism is uncertain; ‘male’ worms have testes, seminal vesicles, male ducts and small spermathecae; ‘female’ individuals possess clitellum, ovaries, ovisac, female ducts and fully developed spermathecae. No mature worms with intermediate sexual condition were observed (Takashima and Mawatari Shunsuke 1998).

8.2.2 The Clitellum The structure of the oligochaete clitellum was investigated by light microscopy by several workers from 1890 to 1980; for reviews, particularly of its ultrastructure and function, see Fernandez and Benito (1987); Jamieson (1981c, 1988a,b, 1992) and Welsch et al. (1984). The structure of the clitellum in several crassiclitellate families is illustrated by light microscopy in Brinkhurst and Jamieson (1971). The clitellum is a specialization of the epidermis for secreting the cocoon in which the eggs are deposited and into which spermatozoa, received into spermathecae from the partner, are extruded to bring about fertilization. Some eudrilids are exceptional in fertilizing the eggs internally before they are shed into the cocoon, entry of sperm from the spermatheca into the ovarian system being demonstrated (Jamieson 1958, 1967; Sims 1967). The clitellum is one

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cell thick in all microdriles and in the Haplotaxidae, Moniligastridae and Allurodidae but in all other oligochaetes, comprising the Crassiclitellata, as the name suggests, the clitellum consists of more than one layer of cells. The single-layered condition of the moniligastrid clitellum is confirmed under that family below (Fig. 8.30). The crassiclitellate condition is interpreted as a modification allowing increased secretion of nutrient materials into the cocoon, a change correlated with a great reduction in both yolk content and size of the eggs and increasing dependence of the embryo for its nutrition on the fluid contained in the cocoon (albumenotrophy) (Jamieson 2000). In megascolecids, where there are usually one or more spermathecal diverticula, the albumen received into the cocoon from the clitellum is presumably augmented from the contents of the spermathecal atrium, the allosperm usually entering from the diverticulum. Our knowledge of the products and function of the clitellar secretions has recently been considerably augmented. In the clitella of Eisenia fetida, the amount of total proteins decreases but the level of soluble proteins increases during puberty. In the cocoons, an increase of total proteins and soluble proteins correlates with the development of the embryos. Among the soluble proteins of the mature clitellum, three major proteins (A, B and C) have been separated by electrophoresis: A, glycolipoproteinic (molecular weight = 450 kDa), B, glycoproteic (MW = 350 kDa) and C, also glycoproteinic (MW = 150 kDa). Only A was present in the cocoons at all stages of development. Although this protein is not provided by the oocyte, it has the same characteristics as a typical vitellogenin. After experimental denaturation of the soluble proteins, four polypeptides were obtained. Two of them, a and c, are involved in the making of this ‘vitellogenin’ (Rouabah Sadaoui and Marcel 1995). With regard to the glucidic and lipidic components of the clitellar epithelium and the cocoon albumen in Eisenia fetida it has been shown that in the clitellum, the sugar concentration increases greatly (about 5-fold) during differentiation. Several monosaccharides increase in quantity during puberty. In addition, glucose is prominent in immature clitella, whereas mannose is the main sugar in mature clitella. About 50% of the dry weight of recently deposited cocoons is carbohydrate. The sugar concentration decreases in the albumen as hatching approached. The amount of total lipids in the clitellum increases during maturation. The levels of neutral lipids, however, varies very little. Phospholipids begin at low levels at the beginning of differentiation, and increased dramatically thereafter. The amount of fatty acids in the clitellar epithelium reached a maximum during the submature stage. Large amounts of fatty acids are stored in the cocoon. Palmitic, stearic, oleic and vaccenic acids preponderate in both the clitellum and cocoon (Rouabah Sadaoui and Marcel 1995b). The reproductive innovation of crassiclitellates allows juveniles to hatch when they have attained a size and energy resources suitable for the terrestrial environment. The size of worms newly hatched from the cocoon is

% Reproductive Biology and Phylogeny of Annelida said to be proportional to the number of clitellar segments involved in secreting the cocoon but also to the amount of yolk (Omodeo and Magaldi 1951, cited in Omodeo 1998). In some species of lumbricids at least, the cocoons are deposited in cocoon chambers composed of faeces from the maternal earthworm; the principal importance of the chambers lies in protection against loss of water but it is suggested that the chambers also supply the cocoons with nutritive fluid and that they are a means of incubation care (Ramisch and Graff 1985). The oligochaete clitellum commonly occupies only a small number of segments in megadriles and as few as two segments in microdriles. The greatest development of the clitellum, in terms of length, occurs in the Almoidea and particularly in the Almidae. In this family, in the genus Alma, the number of clitellar segments varies from 20 to 69. Alma, like other almoids, is also unusual in the posterior location of the clitellum. This is correlated with the post-testicular position of the spermathecae as in oligochaetes the cocoon, after secretion by the clitellum effectively moves forwards to receive sperm from the spermathecae. In Alma the anterior border of the clitellum varies from segment 35 to 247 and the posterior border from segment 49 to 295. In Criodrilus the clitellum begins from segments 14 to 16 and extends for 30 to 34 segments. In the Microchaetidae, the clitellum is also long, beginning on segment 11 to 14 and occupying as many as 44 segments though sometimes a more modest six segments; and in Hormogaster it begins on segment 13 or 14 and extends to segment 24 or as far as 32. The oligochaete clitellum is always behind, or includes, the female pore(s), as the cocoon has also to receive the ova in its forward movement. In the Glossoscolecidae the male pores are intraclitellar, as in Pontoscolex corethrurus, or (Opisthodrilus) postclitellar, they are also intraclitellar, near the anterior border of the clitellum, in the Microchaetidae, Hormogastridae and Almidae, including Criodrilus, though anteclitellar in some almids. In Biwadrilus bathybates, the male pores are immediately anteclitellar, relative to a clitellum of approximately 20 segments. The shorter clitellum is invariably well anterior to the male pores in the Lumbricidae. The clitellum is anterior to the male pores for most or all of its length in the Ocnerodrilidae, Eudrilidae and Megascolecidae in which it occupies a small number of segments (the minimum being three segments, in pheretimoids and some Spenceriella species). The ocnerodrile Nematogenia lacuum is exceptional for this assemblage in the length of the clitellum, occupying approximately 13 segments, commencing on segment 14, with the result that the male pores (on segment 17) are well forward of its posterior end. Some details of clitellar histology are given by Jamieson (1971b), exemplifed for Sparganophilus and Alma in Fig. 8B,D,E (see also those genera in 8.2.11). A detailed review of the ultrastructure and function of components of the clitellum is given by Jamieson (1992). No ultrastructural investigation of the megascolecid clitellum has been published. The histology and histochemistry of the clitellum of the pheretimoid Amynthas hawayanus

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(=Pheretima hawayana) has been investigated by light microscopy (Lufty 1965). The following components of the clitellum were observed: cuticle, supporting cells, sensory cells, mucus secreting cells, large granule cells, fine granule cells and, around the female pore, cells of the “plug.” In taxa with elongate clitella the cocoons are correspondingly long and in almines they are many times longer than wide. The longest recorded in the Alminae are those of Alma multisetosa where the clitellum may occupy 57 segments and in which the cocoons attain a length of 130-155 mm and may contain 32 embryos. In A. nilotica, with a clitellum of 40 to 60 segments, the cocoons (Fig. 8.8F) reach a length of 110 mm and each contains 8-22 young. They average 42 mm (by 12 mm) in Glyphidrilus annandalei in which each contains, on average, seven ova. They reach 70 mm in Criodrilus lacuum, in which they release from 2-8 young, Their elongate spindle-shape in the Alminae contrasts with the more nearly ovoid form of cocoons of other families and appears to be characteristic (diagnostic?) of the subfamily as perhaps is the relatively large number of young released. The cocoons of Sparganophilus tamesis (=S. eisenti), with a clitellum of 14 segments (Fig. 8.8C), are intermediate in length, almost ovoid, though with attenuated extremities, and release from 1-4 young (References in Jamieson 1971b). The cocoons of large earthworms, such as the megascolecid Megscolides australis are several centimeters long and ovoidal.

8.2.3 Chaetae Associated with Reproduction Modified chaetae with a function in reproduction are termed genital chaetae. Those at the male and/or prostatic pores are distinguished as penial chaetae as there is evidence that during copulation they are protruded into the spermathecal orifices of the partner. Other genital chaetae may occur in association with glandular modifications of the epidermis. Those associated with spermathecae are termed copulatory chaetae by some authors but are here termed, less ambiguously, spermathecal chaetae. Penial and spermathecal chaetae are known in some tubificids. Penial chaetae occur in the alluroidid Brinkhurstia and in the biwadrilid Biwadrilus bathybates and are widespread in crassiclitellates (see below). Cuadrado and Martinéz-Ansemil (2001) have provided an investigation and discussion of external structures used in sperm transfer in tubificids, from which the following account is drawn. Judging from the location of genital pores, direct sperm transfer with simple apposition of male and spermathecal pores, or with penetration of differentiated structures such as penes or pseudopenes, appears to be the mechanism of sperm transfer (insemination) used by most tubificids. In some it is aided by penial setae. Observations on the genital region of Potamothrix and Psammoryctides suggest that the embrace of the partners, with formation of a ventral protrusion between spermathecal pores and a retraction of the male pores area, could allow contact between male and sperrnathecal pores, even in species where the latter are clearly lateral. When male and spermathecal pores are aligned, an embrace is not essential to keep them in contact.

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Reproductive Biology and Phylogeny of Annelida

The ciliated (?) and glandular porophore of Protuberodrilus tourenqui could allow the approach of male and spermathecal pores and the adherence of the two worms during sperm transfer. In Peristodrilus montanus, it is likely that the cilia close to the penial chaetal muscles facilitate location of the correct anchorage site. The combined action of the penial chaetal muscles and the strong dorso-ventral muscles could allow the penial chaetae to enter into the anchorage bridge pockets to firmly hold the couple while the pseudopenes enter the spermathecal pores. The small epidermal papilla present in the midventral line of the spermathecal segment of four investigated species of Krenedrilus, combined with their penial chaetae, probably represent a similar anchorage system. The ‘X-shaped mid-ventral slit’ of Bathydrilus rohdei described by Jamieson (1977a) may facilitate the mutual holding of copulating worms (Erséus 1981). A great number of species belonging to several genera from all the tubificid subfamilies have similar penial chaetae that, based on form, location, and orientation, may also have an anchorage function. In Rhyacodrilus falciformis the large and deeply grooved falciform penial chaetae, assisted by several bundles of cilia, may enter into the spermathecal pores, clasping the couple and assisting in sperm transfer. Five of the seven species (Peristodrilus montanus, Protuberodrilus tourenqui, Rhyacodrilus falciformis, Psammoryctides barbatus, Potamothrix bavaricus, P. hammondiensis and P. heuscheri) studied by Cuadrado and Martinéz-Ansemil (2001) had spermathecal chaetae. Protuberodrilus tourenqui also has spermathecal-type chaetae in two non- spermathecal segments (9 and 12). All these chaetae are gutter-shaped, as are most tubificid spermathecal chaetae. Several reasons indicate that they act as piercing chaetae whose principal role is perhaps stimulation, rather than attachment of the concopulants or sperm transfer: (a) they are generally long enough to penetrate deeply into the coelomic cavity; (b) some species from several genera have spermathecal chaetae together with typical anchorage penial chaetae (e.g. Krenedrilus, Rhizodrilus, Limnodriloides); (c) in many species, the spermathecal chaetae are not located near the spermathecal pores, and there are even some species with supplementary sets of spermathecal-type chaetae in non-spermathecal segments (e.g. Protuberodrilus and Krenedrilus). If a mechanical stimulation in fact takes place, the secretions of the large glands associated with spermathecal chaetae, present in many tubificids, could serve to firmly attach the partners while spermathecal chaetae protract and retract alternatively to stimulate the worms. The possibility of chemical stimulation by inoculation of secretions into the blood stream (Feldkamp 1924) deserves further investigation. Either chemical or mechanical stimulation could help to contract the atrium or to contract and expand the coelomic cavity, producing embrace and release, and aspiration, movements aiding sperm transfer. There are other structures and mechanisms, in other tubificids, that probably represent further means of holding partners during sperm transfer: the common median chamber (bursa) into which male pores open in some

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species (e.g. some Rhizodrilus, some Monopylephorus); the development of a cuticular penial sheath of various lengths and forms (e.g. Limnodrilus, Aktedrilus) and the development of special penial chaetae (e.g. Adelodrilus, Inanidrilus). The long and horn-shaped penis sheath of several Aktedrilus species could be interpreted as a means of embracing [holding] the partner and bringing the sperm into the dorsally opening spermatheca (Aktedrilus is devoid of penial chaetae). Erséus (1979) and Erséus and Baker (1982) interpreted the giant penial chaetae of Adelodrilus and Inanidrilus as structures used to transfer sperm into the spermathecae of the mate; the small penial chaetae would be used for attachment (Cuadrado and MartinézAnsemil 2001). Space permits mention of only a few examples of genital chaetae in megadrile families. They do not appear to occur in the Moniligastridae and Ocnerodrilidae. They occur in all ‘aquamegadrile’ families, excepting the Sparganophilidae. In terrestrial earthworms (former Terrimegadrili) they are seen in the Kynotidae, Hormogastridae, Microchaetidae, Lumbricidae, Glossoscolecidae, Eudrilidae and Megascolecidae (Fig. 8.55). In Pontoscolex corethrurus one or both chaetae of the ventral couples in some of segments 14-22 are modified as genital chaetae with longitudinal rows of gouges (Gates 1972; Jamieson, pers. obs.). In the Eudrilidae the penial chaetae assume a great variety of bizarre form (Jamieson 1967) including so-called rolled chaetae (Stephenson 1930). In the acanthodriline Megascolecidae, penial chaetae are often present associated with the prostate glands. In the Australian acanthodriline Diplotrema and its apomorphic relative Neodiplotrema, and the New Caledonian, Acanthodrilus (Fig. 8.55K-N) which appears to be the sister genus of Diplotrema, the penial chaetae may be longer than the thickness of the body and usually have distinctive sculpturing. The spermathecal setae of Acanthodrilus and Diplotrema are sharply pointed and have longitudinal arrays of approximately alternating gouges or notches (Fig. 8.55J,K). Penial chaetae are present in a considerable proportion of the genera of the Megascolecinae, in which they vary interspecifically though they may retain generic or higher characteristics (Fig. 8.55).

8.2.4 Male Pores and Accessory Genital Markings The male pores are the external openings of the vasa deferentia. There is a single pair, even where there are two pairs of testes and vasa deferentia, except in the marine microdrile Randiella, most Haplotaxidae, those Moniligastridae with two pairs of testes, and the Indian megascolecid Hoplochaetella, all of which have two pairs of male pores. Only in the Randiellidae (Erséus and Strehlow 1986) and typical Lumbriculidae are they prosoporous, lying in the same segment as the corresponding testes. By definition, they lie in the segment behind the single pair or last pair of testes in plesioporous oligochaetes: all of the remaining microdriles, which have a single pair of testes; most Haplotaxidae, which typically have two pairs of testes; and the Moniligastridae, which have one or two pairs of testes.

%" Reproductive Biology and Phylogeny of Annelida The segmental positions of the male pores that are typical of the various families are indicated in Fig. 8.4A,B. Location of the pores relative to the clitellum in crassiclitellates is briefly discussed in Section 8.2.2 on the clitellum. In the Megascolecidae, the position of the male pores and their relationship to the openings of the prostates is of great importance taxonomically from the specific to the subfamilial level (Fig. 8.8A). At the specific level, the position of pores relative to chaetal rows is much used. At higher levels the numbers of prostate pores and their locations relative to the male pores are important. The various arrangements in the Megascolecidae are discussed under that family in 8.2.11. In the acanthodrilin condition grooves which is usually regarded as conduits for sperm are present, joining the two pairs of prostate pores to the intermediate male pores (Fig. 8.8A). The genital field illustrated for the African ocnerodrilid Pygmaeodrilus nabugaboensis, suggests utilization of a seminal groove or tract (Jamieson 1957). In the eudrilid genus Stuhlmannia variabilis (Fig. 8.10) it appears that spermatozoa are conveyed via a seminal groove to the summit of the penis, with further incorporation of the two in S. asymmetrica. The most remarkable copulatory mechanism in oligochaetes is perhaps that seen in the genus Alma (Figs. 8.11, 8.56) in which a pair of long claspers, each of which bears a male pore distally, grips the post-testicular, and often far posterior, spermathecal region of the partner. In the Megascolecidae, more than any other oligochaete group, so-called accessory genital markings are well developed, as in Spenceriella penolaensis (Fig. 8.12A) and Heteroporodrilus thomsoni (Fig. 8.12B). These markings usually have the form of paired and/or single pit-shaped or protuberant, segmental and/or intersegmental glandular modifications of the body wall, which may be circular, elliptical or ridge-like or have other forms. They rarely form elongate structures over two or more segments somewhat resembling the tubercula pubertatis of lumbricids. Their function is imperfectly understood but it is likely that they have a role in species recognition and that they may have a “key in the lock” function in copulation, not by mutual insertion, but by precise apposition to structures on the partner. Their prevalence in every other Australian genus makes their absence in the Wet Tropics genus Terrisswalkerius all the more remarkable. Presumably they were lost in an ancestor of the 17 or so currently known species of this genus although a plesiomorphic absence cannot be discounted (they are also absent in the Indian genus Celeriella which appears related). The distribution of accessory genital markings has great taxonomic value for species identification.

8.2.5 Female Pores Much of the variation in the location of the female pores in the Oligochaeta is indicated in Fig. 8.4A,B. They are always located in the segment behind the corresponding ovaries. Predominantly, there is only one pair of female pores as there is only one pair of ovaries in most microdriles and in all but

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Fig. 8.8. A. Arrangement of male and prostatic pores in the Megascolecidae. Original. B. Optical section of clitellum of Sparganophilus tamesis (Sparganophilidae). C. Cocoon of same. D,E. Optical section of clitellum of D. Alma emini and E. A. nilotica (Almidae). F. cocoons of A. nilotica. After Jamieson, B. G. M. 1971b. Anatomy: Glossoscolecidae. Pp. 41-72. In R.O. Brinkhurst and B.G.M. Jamieson (eds), Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto, Fig. 1.6, from various sources.

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Fig. 8.9. Eukerria borellii (Ocnerodrilidae). Genital field, showing seminal grooves connecting prostate and male pores. After Jamieson, B. G. M. 1970. Bulletin of the British Museum (Natural History) Zoology 20: 131-172, Fig. 7F.

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Fig. 8.10. Stuhlmannia variabilis (Eudrilidae). Genital region, showing penis and seminal groove. After Jamieson, B. G. M. 1967. Journal of Zoology (London) 152 79-126, Fig. 12.

three species of the Metagynophora. There are one or two pairs in the Lumbriculidae. Female pores are usually inconspicuous. In the circummundane Eudrilus eugeniae, unusual for the Eudrilidae in having a pair of combined spermathecal and female pores, these are conspicuous and act as vulvae in internal fertilization. In most eudrilids the female pores are separate from the spermathecal pores. The location of the female pores relative to the chaetal lines has some value in taxonomy. Fusion as a single pore is almost invariable in Megascolecinae of the Pheretima group and is seen also in Propheretima and in some species of other genera.

8.2.6 Spermathecal Pores Spermathecal pores are generally inconspicuous, with the notable exception of the Eudrilidae in which they form large, often unpaired orifices (Fig. 8.10). In microdriles they are far pretesticular in enchytraeids but lie in the testis segment in tubificids, including naids, whereas in phreodrilids they occupy

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Fig. 8.11. Variation in claspers of Alma (Almidae). A. Alma basongonis. B. A. emini (Ituri). C. A. emini (=aloysiisabaudiae). D. A. eubranchiata eubranchiata. E. A. eubranchiata catarrhactae. F. A. multisetosa. G. A. nilotica (Egypt). H. A. nilotica (Sudan). I, J. A. pooliana. K. A. stuhlmanni. L. A. kamerunensis. M. A. millsoni millsoni. N, O. A. millsoni millsoni (=schultzei). P. A. millsoni zebangui. Q, R. A. tazelaari .

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S. A. togoensis (Type, Togo). T. A. togoensis (Ghana). U. A. ubangiana. Scale bar 1 mm. With some corrected labelling, after Jamieson, B. G. M. 1971. Glossoscolecidae. Pp. 147-199. In R. O. Brinkhurst and B. G. M. Jamieson (eds). The Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto, Figs. 15.7, 15.8.

& Reproductive Biology and Phylogeny of Annelida

Colour Figure

Fig. 8.12. Example of genital markings in megascolecine earthworms (Megascolecidae). A. Spenceriella penolaensis. After Jamieson, B. G. M. 1974. Earthworms (Oligochaeta: Megascolecidae) from South Australia. Transactions of the Royal Society of South Australia 98: 79-112, Fig. 9B. B. Heteroporodrilus thomsoni Blakemore; specimen from Coolum, Queensland. From Jamieson, unpublished.

the segment of the female pore; in these three groups there is only one pair, with rare exceptions in the Tubificidae. In lumbriculids, spermathecal pores are variable in number, anterior or posterior to the testes (Pinder and Brinkhurst 1994). Haplotaxids have one to four pairs of spermathecal pores, anterior to the testis segments. In alluroidids the pores are pretesticular in one to three of segments 6 to 9, at or near their anterior margins, lateral to dorsal, paired or single. They are also pretesticular in moniligastrids, sparganophilids, Biwadrilus, ocnerodrilids, glossoscolecids (rarely extending into the testis-segments) and megascolecids. In hormogastrids and lutodrilids they are paired or multiple, testicular and posttesticular. In the Almidae, the pores are post-testicular, rarely continued into and in front the

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testis-segments, sometimes translocated into the hindbody, and usually multiple in an intersegment; in Alma they occupy some or all of the eight chaetal lines, or are more numerous, in seven to 37 intersegments (or exceptionally 138 intersegments) from 18/19 to 253/254, thus well behind the testis-segments. In the Kynotidae and Microchaetidae they are posttesticular, though in microchaetids they may also occupy the last testicular segment, and are paired or multiple. In the Lumbricidae spermathecal pores are usually paired, with two to eight pairs in furrows 5/6 to 19/20. They are absent from Criodrilus and Biwadrilus which have external spermatophores (ectospermatophores) and are probably biparental but are also absent from the two circummundane species Ocnerodrilus occidentalis (Ocnerodrilidae) and acanthodrile Microscolex dubius (Megascolecidae, Acanthodrilinae) which lack spermatophores and are deduced to be parthenogenetic (Gates 1972; Jamieson 1974a). The segmental location of spermathecal pores and their position relative to chaetal lines is much used for specific identification in megascolecids. For instance, in Australian megascolecids they are usually intersegmental though in Didymogaster they are segmental in location; there are maximally five pairs, or, in Fletcherodrilus, five midventral pores or are multiple in intersegments 2/3-8/9 in Geoffdyneia rubens (Fig. 8.16) (see also Spermathecae, 8.2.8).

8.2.7 Prostate and other Glands and Bursae Associated with the Male Ducts Various types of so-called prostate glands occur in microdriles. In lower Metagynophora, capsular, smooth-surfaced prostate glands occur in the Moniligastridae and receive the male ducts (Fig. 8.14). These and other accessory sex glands have been reviewed by Adiyodi (1988). The most plesiomorphic of the Opisthopora, the alluroidids, have tubular or bulbous atria which also receive the male ducts or discharge with the latter but separately from them into a terminal chamber; they have an internal epithelium surrounded by a muscular sheath outside which secretory (“prostate”) cells are usually present (see Jamieson 1971a; Omodeo and Coates 2000) (Fig. 8.32). In the crassiclitellate megadriles, dilatations or chambers at the ectal end of the vasa deferentia and protruding into the coelom occur in Kynotus, Criodrilus, commonly in Glossoscolex and some other glossoscolecids and in Biwadrilus. The large gland in Criodrilus is probably responsible for secreting the ectospermatophore of this species and is remarkably similar to the “prostate” of the lumbricid Bimastos palustris. The “copulation glands” of Biwadrilus are glandular prostate-like organs associated with the genital chaetae and are similar in internal structure to the genital chaeta glands of Microchaetus. In the Almidae, only Callidrilus ugandaensis has compact glands which may be considered true prostate glands, though probably not homologous with the prostates of other megadriles. The posterior prostatelike glands of Sparganophilus tamesis and all of these glands in S. smithi,

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Reproductive Biology and Phylogeny of Annelida

resemble those of Callidrilus in having glandular cells grouped around an epithelial layer which lines an intraglandular duct. Sparganophilus differs, however, in the unbranched condition of the duct and, in S. tamesis (=S. eiseni) by differentiation of a terminal, externally recognizable duct (see details in Jamieson 1971a). True prostates, that is glands with a distinct ectal duct and associated with the male terminalia, are typical of the Ocnerodrilidae and Megascolecidae. They have been termed metaprostates (Jamieson 1978b) to distinguish them from the euprostate, which is a dilated ectal loop of the male duct, seen in the Eudrilidae. Three chief types of prostates are recognized in the Megascolecidae (Fig. 8.13) and are here exemplified by Australian taxa. These are tubular, tubuloracemose or racemose. Tubular prostates are slender and elongate and have a single central lumen without side branches. They have their most typical development in the Ocnerodrilidae, and in the Megascolecidae are characteristic of the Acanthodrilinae, though in Diplotrema scheltingai they are tubuloracemose or even racemose and in Exxus they are racemose, and are well exemplified in some Megascolecinae, e.g. Megascolides australis. They are also seen, inter alia, in Cryptodrilus tenuis, Diporochaeta frenchi and Plutellus manifestus (Fig. 8.13A-C) but the latter two species exemplify a tendency in Australian species for some thickening of the gland. In tubuloracemose prostate glands, there is again a central lumen, but thickening of the gland, and often mammilation of the surface, is such as to suggest that groups of cells discharge their secretions by ductules into the central lumen; an example, albeit of relatively simple form, is shown for Graliophilus macedonensis in Fig. 8.13D. The various species of Terrisswalkerius exemplify transitions from tubular to tubuloracemose prostates. In the racemose prostate, the external form is such as to suggest that the duct branches immediately within the gland. This is clearly the case where the gland is bipartite, as in Digaster armifera (Fig. 8.13I). An extreme case of the racemose form is seen in Trinephrus fastigatus (Fig. 8.13E), in which the duct forms several branches external to the gland. A unipartite racemose gland is exemplified by Notoscolex camdenensis (Fig. 8.13H), a deeply incised form by Heteroporodrilus shephardi (Figs. 8.13G), and a unipartite racemose gland, which may be an extreme development of the tubuloracemose type, in Anisochaeta mudgeanus. The structure of the prostate glands of the peregrine megascolecin Lampito mauritii has been investigated by Gupta et al. (1999). Two types of secretory cells are present. Type 1 cells with a broad basal region and a long apical region contain electron-dense oval secretory granules with an increased density at the core region. Numerous electron-lucent granules with fine filamentous and electron-dense amorphous materials also occur at the basal region of these cells. Type 2 cells contain electron-lucent mucouslike secretory granules. This cell type contains exceptionally large Golgi complexes with 20-23 stacked cisternae. Both cell types open into a common

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Fig. 8.13. Prostate glands. Some variation in Australian Megascolecinae (Megascolecidae). A. Tubular (Cryptodrilus tenuis). B. Tubular (Diporochaeta (Vesiculodrilus) frenchi). C. Tubular (Plutellus manifestus). D. Tubuloracemose (Graliophilus macedonensis). E. Racemose, bipartite with external branching of the duct (Trinephrus fastigatus). F. Racemose, unipartite, extending through several segments (Anisochaeta mudgeanus). G. Racemose, deeply incised (Heteroporodrilus shephardi). H. Racemose, unipartite, in a single segment (Notoscolex camdenensis). I. Racemose bipartite, with internal branching of duct (Digaster armifera). From Jamieson, B. G. M. 2000. CD ROM. The Native Earthworms of Australia (Megascolecidae Megascolecinae). Science Publishers, Inc., Enfield, New Hamphshire, Fig. 0.39.

&" Reproductive Biology and Phylogeny of Annelida lumen and numerous microtubules are visible at the apical end. Junctional complexes such as desmosomes and septate junctions are present in this glandular tissue.

8.2.8

Spermathecae

Spermathecae are epidermis-lined sacs which receive spermatozoa from the partner during copulation and store them for later expulsion into the cocoon and fertilization of the eggs. Each spermatheca has a sac-like ampulla and a duct. The duct may be so short that the ampulla is virtually sessile on the body wall, but is usually some significant fraction of the length of the ampulla or longer. The spermathecae of the Moniligastridae are almost unique in the Oligochaeta in the great length of their ducts, a development paralleled by the great length of the vasa deferentia which Bourne (1894) found to measure 9.5 inches in only a single segment. In the advanced genera Drawida and Moniligaster, there is a large atrium at the ectal end of the spermathecal duct and in Moniligaster (Fig. 8.14), this has developed a peculiar “dichotomous gland”. The similarity of the spermathecal atrium to the prostates at the ends of the male ducts (Fig. 8.15) suggests that a common morphogenetic controlling mechanism may be involved in the production of

8.14. Moniligaster troyi. Spermathecal apparatus. After Jamieson, B. G. M. 1977. Evolutionary Theory 2: 95-114, Fig. 5C.

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8.15. Moniligaster troyi. Male genital apparatus. After Jamieson, B. G. M. 1977. Evolutionary Theory 2: 95-114, Fig. 5E.

the spermatheca and prostate (Jamieson 1977b). These appendages of the spermathecal ducts do not, however, store sperm; their tubular constituents are here shown to consist of a tall glandular epithelium (Fig. 8.31C,D,E). In most Megascolecidae and a few species of some other families there are one or more diverticula connected with the spermathecal duct or sometimes with the lower part of the ampulla. In all diverticulate megascolecid spermathecae, except, as far as is known, Megascolides australis (Van Praagh 1995) and Microscolex macquariensis (Jamieson, unpublished), the spermatozoa are stored in the diverticulum. Megascolecid spermathecae are pretesticular. There is usually one pair, occasionally a single midventral spermatheca, per thecal segment but some pheretimoids have several spermathecae per segment. The latter, polythecal condition is known in only one megascolecine Australian species, Geofdyneia rubens (Fig. 8.16). In the Ocnerodrilidae spermathecae rarely (Pygmaeodrilus, some Gordiodrilus) have diverticula; in some ocnerodrile species the diverticula differ from those of megascolecids in not containing spermatozoa, which are stored in the ampulla, as in Pygmaeodrilus montiskenyae (Fig. 8.17), but in at least P. nabugaboensis (see Jamieson 1957), the diverticula and not the spermathecal ampullae are inseminated. In the Glossoscolecidae, usually lacking diverticula, each of the two spermathecae of Glossoscolex schutti has a pair of diverticula of the duct which receive sperm (Michaelsen 1918).

&$ Reproductive Biology and Phylogeny of Annelida

Fig. 8.16. Geofdyneia rubens (Megascolecidae). An example, rare in megascolecids, of multiple spermathecae in a segment. From Jamieson, B. G. M. (2000). CD ROM. The Native Earthworms of Australia (Megascolecidae Megascolecinae). Science Publishers, Inc., Enfield, New Hamphshire, Fig. 14.2.

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8.17. Pygmaeodrilus montikenyae (Ocnerodrilidae). A. Dorsal dissection to show the right spermatheca. B. Vertical section of a spermatheca, showing spermatozoa in ampulla. After Jamieson, B. G. M. 1965. Annals and Magazine of Natural History 8: 95-107, Figs. 4 and 5.

&& Reproductive Biology and Phylogeny of Annelida In the Enchytraeidae, e.g. Lumbricillus rutilus, the ental end of each spermatheca (often in the form of an ampulla) of which there is a pair in segment 5, opens into the lateral aspect of the alimentary canal (Stephenson 1930). Westheide (1999: 199) has stated that, in Marionina preclitellochaeta, the ampullae are intestinal pockets and not part of the epidermal invaginations. In the same paper, Westheide (1999: 210) draws out of the study of the spermathecae-gut connections in three enchytraeid species an interesting speculation on the origin of the whole taxon Oligochaeta: “In principle, all sperm are wasted that do not fertilize an egg, and in general these are the great majority. In a hermaphrodite species with direct transfer of sperm, there is no loss of male gametes. In this case, each individual is capable of reproducing and all the sperm that are not transferred and remain in the body can be resorbed—that is, recycled—and contribute to the energy budget for female reproduction just like all allosperm that are transferred but fail to fertilize eggs. Special structures, such as the genitointestinal connections... may serve to introduce allosperm into the recycling process in a particularly simple and rapid manner. Finally, this idea leads to the assumption that the evolution of hermaphroditism should be favored in small species. ....A conclusive phylogenetic inference that could be drawn from this reasoning for a taxon composed exclusively of hermaphrodites, as the Clitellata are, is that its stem species was small-bodied (about several millimetres) and reproduced by direct sperm transfer”. In the Megascolecidae, the number, shape, and length relative to the ampulla of the spermathecal diverticula are important taxonomically and are used in specific identification or even at the generic level. Thus the genus Simsia in south eastern Australia typically has distinctive rosette-like multiloculate diverticula. Although the form of the spermatheca may allow specific distinction within a genus, the same or a similar form may, nevertheless, occur in other genera. Some major variations in the form of spermathecae exemplified by Australian megascolecids are illustrated in Fig. 8.18. Spermathecae are absent from Criodrilus lacuum, Criodrilus bathybates, Ocnerodrilus occidentalis and Microscolex dubius (see 8.2.6, above). In the megascolecids Amynthas hawayanus, A. morrisi and Metaphire californica, most of the sperm are found within the diverticulum as is usual for Megascolecidae. The ‘bladder’ (ampulla) is the secretory portion and releases a mucous secretion, identified as acid carboxylated and/or neutral glycosaminoglycans linked to neutral proteins. The spermathecal content is thought to play a role in nourishing spermatozoa since the male gamete itself has little intrinsic energy. (Omodeo 2000 suggests that some of the ampullary secretion also enters the cocoon). Cells of the ampulla show a vertical cytoplasmic zonation, and the epithelial surface displays slender, widely spaced, ramified microvillosities. The apical zone is filled with dense vesicles. The central zone with mitochondria, Golgi complexes and rough endoplasmic reticulum is highly developed. The basal zone is made up by vertical compartments consisting of basal plasma membrane folds which

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Fig. 8.18. Contrasted types of spermathecae in the Megascolecinae (Megascolecidae). A. Unidiverticulate and uniloculate (Arthuridrilus burniensis). B. The same, elongate (Terrisswalkerius nashi). C. Unidiverticulate with terminal loculi (Gemascolex similis). D. Bidiverticulate, simple (Anisochaeta enormis). E. Multiloculate (Digaster armifera). F. The same (Anisochaeta (=Oreoscolex) saccarius). G. Bidiverticulate and multifurcate (Heteroporodrilus tryoni). H. With two lumina and apically multiloculate (Terrisswalkerius oculatus). Jamieson, B. G. M. 2000. CD ROM. The Native Earthworms of Australia (Megascolecidae Megascolecinae). In Science Publishers, Inc., Enfield, New Hamphshire, Fig. 0.41.

contain glycogen. Diverticulum cells are of lesser height with the apical surface presenting cytoplasmic projections where sperm cells are stored and vesicles having a dense secretion different from that of ampullary cells (Teisaire and Roldan 1995). In the megascolecid Amynthas rodericensis evidence has been tentatively advanced for holocrine secretion by cells of the epithelium of the

' Reproductive Biology and Phylogeny of Annelida spermathecal diverticula of materials putatively nutritive for spermatozoa contained in the diverticula and it has further been tentatively suggested that the sperm acrosomes contribute to lysis of the epithelium (Jamieson 1992).

8.2.9

Spermatophores

Spermatophores in oligochaetes have been defined by Jamieson (1978c) and discussed in detail by Ferraguti (1999). In oligochaetes, spermatophores have been found attached to the body wall or within the spermathecae; they are termed ectospermatophores and endospermatophores respectively. It may be difficult to determine whether the endospermatophores are produced by the inseminating concopulant or by the spermathecal wall. Spermatophores secreted by the spermatheca, reported in some Eudrilidae and Megascoelcidae are termed thecacysts (Jamieson 1978c). As reviewed by Ferraguti (1999), (ecto)spermatophores have been reported in no less than 27 lumbricid species by Bouché (1975) as structures, presumably deposited by the partner, attached to the exterior of the body; they are less than one mm long, with a flat base and a small mass of sperm wrapped in a chitinous sheath. The sheath is produced by the atrial gland (Perel’ 1978). There are two types of spermatophores: club-shaped ones, and flat ones, inserted into the integument. The presence of spermatophores in Spermophorodrilus albanianus has led Bouché (1975) to suppose that the cocoon, during its anterior migration, would collect the spermatophores and fertilization should occur within the cocoons, as usual in oligochaetes: thus hypodermic impregnation is not suspected. Earlier Jamieson (1971c) had similarly suggested that sperm from the ectospermatophores (received from the partner) of Criodrilus were shed into the cocoon. Ferraguti (1999) considered it questionable that this mechanism could be attributed to all the lumbricid species in which spermatophores have been found (Ljungström 1968) as the segments with spermatophores vary from the clitellum to the preclitellar region (Perel’ 1978). These locations do not here appear to constitute an impediment to deposition of ectospermatophores or their spermatozoa into the cocoon as it effectively moves forward along the body. It is noteworthy in this regard that they are never postclitellar. Where spermatophore-bearing species are known to be parthenogenetic, the spermatophores have been regarded as ‘residual organs’. Perel’ (1978) argues that the exchange of spermatophores adhering to the partner’s body wall could have been the original way of insemination in Lumbricidae, preceding the ‘invention’ of spermathecae. However, it is evident that spermathecae were developed very early in oligochaete evolution (Ferraguti 1999) and Westheide (1999) has argued for direct transfer as the primitive mechanism. Ectospermatophores consisting of an ovoid chamber with an opening at one end and a stalk at the opposite end occur in the tubificids Bothrioneurum and Bacescuella attached to the body wall in the clitellar region (Mann 1984). Both species lack a spermatheca, thus a possible function of the spermatophores is evident.

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In the almoid Criodrilus lacuum ectospermatophores are found in a number variable from two to six attached to the ventral surface, before and behind the genital pores, in some of segments 13-19 (Jamieson 1971c) (Fig. 8.19). (The clitellum lies in segments 14-47.) They have the appearance of small horns, a few millimeters long, with a chitinous wall, containing a large

Fig. 8.19. Criodrilus lacuum (Criodrilinae, Almidae). A. Genital region. B. Genital region, showing attached ectospermatophores. C. An ectospermatophore attached to the body wall. After Jamieson, B. G. M. 1971. Glossoscolecidae. Pp. 41-72. In R. O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World. Oliver and Boyd, Edinburgh, Toronto, Fig. 15.11F,G,H.

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Reproductive Biology and Phylogeny of Annelida

amount of spermatozoa arranged in very regular layers (Ferraguti and Jamieson, unpublished in Ferraguti 1999). (Ecto)spermatophores have also been reported for the lutodrilid Lutodrilus vesiculatus (McMahan 1979) (Cuadrado and Martinéz-Ansemil 2001) and for the bathydrilid Bathydrilus (=Criodrilus) bathybates by Nagase and Nomura (1937). In other clitellates, ectospermatophores have been found in some branchiobdellids and they are widely distributed among hirudineans, being present in all Rhynchobdellida and in Erpobdelliformes among the Arhynchobdellida (Ferraguti 1999; see also chapter 9 of this volume). Jamieson (1967) was not able to confirm the presence of a spermatophore within the spermatheca of the eudrilid Stuhlmannia variabilis which was illustrated by Beddard (1901). That author also reported spermatophores, within the spermtheca, in the same family, in Polytoreutus. Whether they were endospermatophores or thecacysts is uncertain. Internal spermatophores have also been reported for the acanthodriline megascolecid Dichogaster austeni (see Stephenson 1930).

8.2.10

Spermatozeugmata

Spermatozeugmata may be defined as sperm aggregates implanted in the spermatheca by the concopulant, characterized by a repetitive order of the spermatozoa and the presence of some sort of cementing agent, but lacking a proper capsule (Ferraguti et al. 1989; Ferraguti 1999). Spermatozeugmata are said to be formed in the spermathecal ducts during copulation (Dixon 1915). However, the cementing substance has also been said to originate from the atrial epithelium (Jaana 1982). Spermatozeugmata are known from tubificids of the subfamilies Tubificinae, Limnodriloidinae, Phallodrilinae and Rhyacodrilinae. The shape and organization varies within some of the subfamilies, such as Limnodriloidinae but is diagnostic in others, such as Tubificinae (references in Ferraguti 1999). All investigated tubificines, with the exception of Aulodrilus and related forms, have spermatozeugmata with the shape of rods 1-2 mm long, with a diameter of less than 100 µm. The spermatozeugmata are formed by eusperm and parasperm (for terminology see Healy and Jamieson 1981) as they are in some Limnodriloidinae (see sperm dimorphism, 8.4.3, below). Sperm dimorphism is not involved in the Phallodrilinae where there is a single model of spermatozeugma described for Bathydrilus formosus. In Bathydrilus the single spermatozeugma present in the spermathecae is a skein composed of euspermatozoa with the acrosomes in the central portion, and nuclei and tails wound around this (references in Ferraguti 1999). In the Rhyacodrilinae, spermatozeugmata which consist of only one type of spermatozoon have been reported in Rhyacodrilus arthingtonae by Jamieson (1978c) and Rhizodrilus russus by Ferraguti et al. (1994). In both there are also many free spermatozoa. In R. russus, the spermatozeugmata are formed by a sort of cap surrounding the anterior portion of a sperm bundle, the sperm tails being free. The cap is formed by an electron-dense sheath 0.7 µm thick, terminating with a characteristic rim. The spermatozeugmata are apparently

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identical to those described by Martin and Giani (1995) for another rhyacodriline, Epirodrilus michaelseni (see Ferraguti 1999).

8.2.11 Taxonomic Survey of Reproductive Systems Overview. Notable distinctions of the Clitellata from most polychaetes are the restriction of the gonads to a few segments and their hermaphroditism. The plesiomorphic arrangement for oligochaetes is very tentatively considered to be two testicular followed by two ovarian segments (Brinkhurst and Jamieson 1971). In the Lumbriculata the numbers of gonads are very variable (particularly when branchiobdellids and leeches are included). In the Tubificata and Diplotesticulata there are one or two pairs of testes, excepting the lutodrilid crassiclitellate Lutodrilus multivesiculatus (see McMahan 1976) which, by interpolation, has 10 testicular segments. Two pairs of testes (the holandric condition), in segments 10 and 11, is the predominant condition in the megadriles (Metagynophora), but reduction to a single pair in segment 10 (the proandric condition) occurs inter alia in most Alluroididae (Kathrynella guyanae Omodeo 1996 has testes in segment 11, and male pores in 14, a displacement of one segment behind the usual alluroidid positions), some glossoscolecids, some Microchaetus species and, allowing for segmental deletions, in the moniligastrids Hastirogaster and Eupolygaster. The alternative, metandric condition, testes in segment 11 only, is widespread in the Metagynophora, including the moniligastrids Moniligaster and Drawida, allowing for segmental deletions, and characterizes some megascolecid genera (e.g. Begemius and Trichaeta). Development of spermatogonia or spermatocytes through spermatids to spermatozoa usually occurs in pouches of the septa which delimit the testis segments. These pouches, the seminal vesicles, are maximally, in earthworms, four pairs, on the anterior and posterior septa of the testis segments. Sometimes there is a pair of “pseudovesicles” of unknown function, or sometimes containing spermogenetic stages, at septum 13/14. The number and form of the seminal vesicles has some taxonomic value. Leaving aside derived members of the Lumbriculidae, the maximum number of functional ovarian segments in oligochaetes is two, seen only in some haplotaxids and three exceptional species of crassiclitellates among some 28 oligochaete families, the remainder having a single ovarian segment (Jamieson 1992). The arrangement of reproductive organs is summarized in Fig. 8.4A,B. 8.2.11.1 Subclass Randiellata Randiellidae. The Randiellidae is a small marine family the reproductive system of which is imperfectly known (Erséus and Strehlow 1986; Erséus 1997). The number and position of the gonads in this family are unclear, but Randiella caribaea appears to have at least one testis in segment 10 and one ovary in segment 12 (Erséus 1997) whereas two pairs of testes followed by questionably one or two (paired?) ovaries was previously reported (Erséus and Strehlow 1986). The possibly euprosoporous condition of the male ducts

'" Reproductive Biology and Phylogeny of Annelida (male pores in the same segment as their testes), if confirmed, was considered by Jamieson (1988b) to merit placement in a separate basal subclass, the Randiellata. Distribution of the gonads, over more than two segments, certainly appears to merit separation from the Tubificata. Spermathecae are known to be dorsal or lateral. 8.2.11.2 Subclass Tubificata Order Tubificida. Suborder Tubificina. Tubificidae. In the Tubificidae and the Naididae, which were convincingly subsumed as a subfamily in the former by Erséus et al. (2002) there is typically one pair of testes followed in the next segment by one pair of ovaries (the monotesticulate condition). The male pores are in the segment behind the testis segment (the plesioporous condition). The testis segment is usually 10 (Tubificidae sensu stricto) or is more anterior in ‘naids’. The male duct in Tubificidae sensu lato comprises a funnel, a vas deferens, an atrium and, frequently, a copulatory structure (male bursa). There may also be a prostate gland (diffuse or compact) in association with the duct. A tubificid reproductive system is exemplified in Fig. 8.20 by that of Macquaridrilus bennettae, a species which is, however, unusual for the Tubificidae in possessing a spermathecal diverticulum (Jamieson 1968a) and by Rhyacodrilus arthingtonae (Fig. 8.21). The development of the male and oviducal system has recently been clarified by Gustavsson and Erséus (1997, 1999). The funnel and vas deferens in Clitellio arenarius, Tubificoides benedii, Heterochaeta costata, Rhyacodrilus coccineus, Monopylephorus rubroniveus (Rhyacodrilinae) and the ‘naid’ Stylaria lacustris originate from peritoneal (mesodermal) cells in the posterior septum

Fig. 8.20. Macquaridrilus bennetae (Tubificidae). Exemplifying the genital organs of a tubificid. This species is unusual in having a spermathecal diverticulum. After Jamieson, B. G. M. 1968a. University of Queensland papers. Department of Zoology 3: 55-69, Fig. 5.

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Fig. 8.21. Rhyacodrilus arthingtonae (Tubificidae). Photomicrograph of genital organs. After Jamieson, B. G. M. 1978. Proceedings of the Royal Society of Queensland 89: 39-43, Plate 5.

in the testis segment. The atrium develops from an epidermal (ectodermal) invagination but a typical atrium is not formed in M. rubroniveus in which the entire duct is of mesodermal origin. In the latter species, a shallow epidermal invagination occurs, into which both male ducts open; it bears resemblance to a copulatory structure, which usually forms from a secondary invagination, rather than to a proper atrium. The prostate glands have different origins in different taxa; in S. lacustris, R. coccineus and M. rubroniveus they differentiate from the peritoneal (mesodermal) cells surrounding the atrium but in H. costata and T. benedii they develop by a multiplication of the atrial (ectodermal) cells; C. arenarius lacks a prostate gland. The extent of the diffuse prostates in rhyacrodrilines varies: in R. coccineus the cells cover the atrium, whereas in M. rubroniveus they cover only a part of the vas deferens. In all of these species, the spermatheca is confirmed as being of ectodermal origin, developing as an invagination of the epidermis, and the female ducts develop from peritoneal (mesodermal) cells in the posterior septum of the ovarian segment. However, in M. rubroniveus the first rudiment of the oviduct disappears and a proper duct never develops. The position of

'$ Reproductive Biology and Phylogeny of Annelida the oviduct differs; in S. lacustris the female duct is lateral and the pore is in the ovarian segment, while in the tubificids studied the duct is ventral with the pore in the segment behind the one containing the ovary (Gustavsson and Erséus 1997, 1999). Naidinae are notable in having asexual, in addition to sexual, reproduction. They reproduce asexually by paratomic fission, including regeneration of a fixed number of anterior segments in secondary individuals. This provides a great potential for pupulation growth (Erséus 2003). Phreodrilidae. Phreodrilids have a pair of testes in segment 11 and a pair of ovaries and male pores in 12. Spermathecal pores are ventral or dorsal on 13, often with vestibulae; spermathecal ampullae in 13 or usually more posterior. The female funnels are on septum 12/13 or entering into spermathecal vestibulae. Atria are present, usually with a very narrow lumen, without prostate glands. Pendant penes or eversible pseudopenes are usually present. Asexual reproduction is rare (Pinder and Brinkhurst 1994). Internal fertilization is suspected in some species. Opistocystidae. Opistocystids are typical plesiopores but the gonads are unusually posterior in Opistocysta. In O. funiculus (in South America and Africa) testes are in segment 21, ovaries in 22, sperm funnels on 21/22, male pores on 22, female pores apparently in 22/23, and spermathecal pores anterior on 23. Seminal vesicles extend from 21/22 into 22 and ovisacs from 22/23 to 26. O. corderoi has a similar configuration of genital organs but they are further forward, the testes being in segment 14 or 15, with the clitellum at least from 15-16. In Lycodrilus, tentatively placed in the Opistocystidae, the genitalia have the tubificid arrangement: testes in segment 10, ovaries in 11, male pores in 11, female pores in 11/12. Spermathecae are in segment 8 or 10 and spermatophores are present. There are small atria with stalked prostates (Cook 1971b). Dorydrilidae. Dorydrilids are plesioporous, with one pair of testes and funnels in segment 9 and male pores in 10. Spermathecal pores are in segment 10 or 11, or both, spermatheca being atrial or post atrial or both. The ovaries are in segment 10. A pair of vasa deferentia joins a pair of large, musuclar atria in 10 (Cook 1968, 1971a). Cook has espoused derivation of Dorydrilus from a lumbriculid ancestral form but if affinities lie with the Tubificidae this origin is unlikely in view of the wide phylogenetic separation of lumbriculids and tubificids indicated from molecular studies. Enchytraeidae. The Enchytraeidae, Propappidae and Capilloventridae have what may be the most plesiomorphic condition of the male ducts known in oligochaetes, the usual absence of a male atrium. If this is a symplesiomorphy it could not be used to unite these families. However, some enchytraeids have a well developed male atrium (Fig. 8.22) as in most other oligochaete families. The Enchytraeidae is one of the largest families of oligochaetes and is abundant terrestrially as well as in freshwater and marine habitats.

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Fig. 8.22. A. Enchytraeus albidus (Enchytraeidae). Seminal funnel. Note prostate-like preseptal expansion. B. Propappus glandulosus (Propappidae). Seminal funnel. From Michaelsen, W. 1928. Oligochaeta. Pp. 1-118. In W. Kukenthal and T. Krumbach (eds). Handbuch der Zoologie. 2. Walter de Gruyter and Co, Berlin, Fig. 45. C. Mesenchytraeus antaeus. Ectal male apparatus showing atrium, accessory glands, penial glands and a penial chamber or bulb. The apparatus is highly complex, contrary to what is usually envisaged for enchytraeids. After Rota, E. and Brinkhurst, R. O. 2000. Journal of Zoology London 252 (1): 27-40, Fig. 6.

Enchytraeids have a pair of testes in segment 11 followed by a pair of ovaries, in 12, in which segment the male pores are situated. Spermathecae are paired in segment 5, with pores in 4/5 (Fig. 8.4A). They have a long swollen glandular preseptal duct immediately posterior to each seminal funnel (Fig. 8.22A) and this may be regarded as an alternative strategy to having prostate glands for providing seminal secretions. Other male glands, and penial bulbs, are usually absent. However, the absence or weak development of atria and associated glands is possibly size dependent (see Mesenchytraeus antaeus, below). Many details of the reproductive anatomy of Fridericia are given in the valuable monograph of Schmelz (2003).

'& Reproductive Biology and Phylogeny of Annelida Although most enchytraeids are microdriles, some are as large as earthworms. Mesenchytraeus antaeus an unpigmented giant enchytraeid from Vancouver Island, reaches, in fixed material, 61 × 2.9 mm (with the diameter maximal in the midbody). Its spermathecal pores open laterally in the middle of segment 5. The male apparatus has unusually long sperm funnels (extending over 10 segments) and two groups of large accessory glands opening through two circular papillae at the base of each penial bulb, independently of the penial pores (Rota and Brinkhurst 2000). The ectal male apparatus of this species (Fig. 8.22C), which includes an atrium, accessory glands, penial glands and a penial bulb, is highly complex, contrary to what is usually envisaged for enchytraeids. This perhaps suggests that simplicity of the reproductive system in small species is a secondary condition. Most enchytraeids have no sperm sacs (seminal vesicles) or ovisacs, apparently by loss. These sacs are, however, present in Mesenchytraeus, Grania, Randidrilus and a few species in other genera (Omodeo 1998). Propappidae. The Propappidae were removed from the Enchyraeidae because they have characters which are not seen in the latter family: sigmoid, nodulate, bifid chaetae; large epidermal glands posterior to each chaetal bundle; spermathecal pores at septum 3/4; and glandular parts of the vasa deferentia located posterior to 11/12 (Fig. 8.22B) unlike the preseptal glandular enlargement in enchytraeids. Another significant, unique character state is location of the single pair of ovaries in segment 13, except in Propappus arhyncotus, rather than in the enchytraeid location in the male pore segment, 12 (Coates 1986). The presence of a one-segment hiatus (requiring confirmation) between testes and ovaries is here considered homoplastic with the Metagynophora. Capilloventridae. Capilloventrids have one pair of testes in segment 11 and a pair of ovaries and male pores in 12. Atria, prostates and penes are absent. Female pores are in 13 (to be confirmed in some species). Spermathecal pores are lateral at 6/7, with the ampullae in segment 7 (Pinder and Brinkhurst 1994). With their simple organization, including dorsal and ventral bundles of hair chaetae (shared with Parividrilidae), as in some polychaetes, the capilloventrids appear to be the most plesiomorphic oligochaete (and clitellate) family and this position has been confirmed in 18S rDNA parsimony analysis (Erséus and Källersjö 2003). Their plesiomorphy possibly militates against regarding the octogonadal condition as primitive for oligochaetes. Parvidrilidae. In ovigerous specimens of the type-species Parvidrilus strayeri (Fig. 8.23), a clitellum is developed as a pair of lateral rows of a few, large, swollen epidermal cells in segments (9-) 10-12 (-13, -14). It is unclear whether gonads are paired or unpaired; testes (testis?) are in segment 11, ovaries (ovary?) in 12. The other reproductive organs include a U-shaped muscular ‘genital body’, in segment 12 and a V-shaped ‘copulatory organ’, consisting of two conjoined penes (or pseudopenes) in a penis sac or sheath, in segments 12-13; the exact nature and function of these structures are unknown (Erséus 1999). Martinez-Ansemil et al. (2002), for the second known

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Fig. 8.23. Parvidrilus strayeri (Parvidrilidae). A. Schematic horizontal view of segments 12 and 13, showing general outline of ‘genital body’ and ‘copulatory organ’. B. Somewhat horizontal view of segments 11-13 of a paratype. C. Somewhat lateral view of segments 11-13 of a further paratype. Relabeled after Erséus, C. 1999. Proceedings of the Biological Society of Washington 112(2): 327-337, Fig. 2.

species, P. spelaeus, suggest that the genital body and copulatory organ are respectively the atrium and spermathecae. Narapidae. Narapidae, a monotypic family (Righi and Varela 1983), are plesioporous and resemble the Naidinae in having the testes in segment 5 but differ from these in having the spermathecae in the ovarian, not the testicular, segment (Fig. 8.24). Male pores and atria are in 6, and penes are present. The

! Reproductive Biology and Phylogeny of Annelida

Fig. 8.24. Narapa bonettoi (Narapidae). A. Lateral view of segments 6-8. B. Transverse section of atrium. C. Lateral view of male ducts. Relabelled after Righi, G. and Varela, M. E. 1983. Revista De La Asociacion De Ciencias Naturales Del Litoral 14(1): 7-15, Figs. 4-6.

atria are covered by diffuse gland cells. Whereas the testes and efferent ducts are paired, the ovary, in 7, is unpaired. There is a pair of spermathecae in 7. Like the Randiellidae and Propappidae, there is a gonad-less segment between the testicular and ovarian segments. This might represent a proandric condition derived from former holandry. 8.2.11.3 Subclass Lumbriculata We will here deal only with the oligochaetous members, the Lumbriculidae. Other taxa here included are the Branchiobdellida, Acanthobdellida and Euhirudinea which are discussed in Chapter 9. Lumbriculidae. The Lumbriculidae is an Holarctic family with extension into West Asia. Some species have become widely distributed, including the Southern Hemisphere. The reproductive system is very variable. There are one to four pairs of testes, in variable locations. Atria are one to four pairs, located between segments 7 and 15, paired or unpaired, always in a testisbearing segment, each being associated with one or two pairs of testes (Figs. 8.4B, 8.25). There are commonly two pairs of testes in adjacent segments, both with funnels and vasa deferentia feeding a single pair of atria in the same segment as the posterior pair of testes. Sometimes the anterior testes and ducts are absent, leaving a single pair of atria, testes and vasa deferentia within one segment, and then often with this arrangement serially repeated. There are one or two pairs of ovaries beginning one, or rarely two, segments behind the most posterior testis-bearing segments. Spermathecae are variable

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Fig. 8.25. Bythonomus mirus (Lumbriculidae). Atrium and posterior male gonoduct. After Chekanovskaya, O.V. 1981. Aquatic Oligochaeta of the USSR, United States Department of the Interior and the National Science Foundation, Washington, D.C., Amerind Publishing Co. Pvt. Ltd., New Delhi, pp. 513, Fig. 232.

in number and either anterior or posterior to the testicular segments (Pinder and Brinkhurst 1994). Michaelsen (1928-32) (see Michaelsen 1928) brilliantly foreshadowed the findings of molecular phylogenetics when he illustrated (Fig. 8.26) a pathway from lumbriculid organization to that of hirudinid leeches. The progressive stages were exemplified by 1) the lumbriculid Rhynchelmis, with compact testes but long seminal vesicles, extending through several segments; 2) the lumbriculid Agriodrilus vermivorus, in which a chain of testes has developed within the elongate seminal vesicles, though still with a single pair of

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Reproductive Biology and Phylogeny of Annelida

Fig. 8.26. Hypothetical scheme suggesting that the testicular sacs of leeches corrrespond to the seminal vesicles of lumbriculids. A. Rhynchelmis. B. Agriodrilus vermivorus offers an intermediate in which a long series of testicular portions has developed in the original seminal vesicles. In an abnormal condition one of these was seen to be isolated like a testis in the chain of testes occurring in leeches. C. The leech condition: a chain of postovarian testes. Modified from Michaelsen, W. 1928. Oligochaeta. In W. Kukenthal and T. Krumbach (eds). Handbuch der Zoologie 2, Fig. 93.

seminal funnels, and in which the coelom is constricted; 3) Hirudo in which each testicular chamber has acquired its own pair of seminal funnels and the coelom has been reduced to a system of sinuses. The leech distinction from oligochaetes, extension of testes posterior to the ovaries, was thus explained in terms of modification of pre-existing seminal vesicles. 8.2.11.4 Subclass Diplotesticulata The validity of recognizing the Diplotesticulata is discussed in 8.1.4 above. Superorder Haplotaxidea. Order Haplotaxida sensu stricto. The haplotaxid reproductive system usually has two pairs of testes, in segments 10 and 11 (rarely in 9 and 10); the anterior pair is rarely absent. There are one or two pairs of ovaries in the segments following the testicular segments. The male ducts are simple and lead to ventrolateral or lateral male pores. However there is a large glandular mass between the male pores in Hologynus hologynus, in which both pairs of male pores lie in the same segment, the posterior vasa deferentia being reflexed forward. The two pairs of vasa deferentia also open into a single segment in Pelodrilus violaceus but

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in that case the anterior vasa penetrate more than one segment, discharging near the posterior pair in segment 12. In Adenodrilus denticulatus there are four pairs of large copulatory glands which open externally near the ventral setae and are not directly associated with the male ducts (Chekanovskaya 1981) (Fig. 8.27). These glands are reminiscent of those of Sparganophilus, a genus which has in the past been placed in the Haplotaxidae (Tétry 1934), but the molecular study (Jamieson et al. 2002) indicates that at least the type-species, Haplotaxis gordioides, is genetically distant from Sparganophilus. One species, H. brinkhursti, has lost the anterior pair of ovaries and therefore is unique in the known Haplotaxidae in having the metagynophoran condition.

8.27. Adenodrilus denticulatus (Haplotaxidae). Lateral view of genital organs, showing large copulatory glands. After Chekanovskaya, O. V. 1981. Aquatic Oligochaeta of the USSR, United States Department of the Interior and the National Science Foundation, Washington, D. C., Amerind Publishing Co. Pvt. Ltd, New Delhi. pp. 513, Fig. 204.

Tiguassuidae. The Tiguassuidae was recognized as a family by Jamieson (1988b) and by Brinkhurst (1988) in morphocladistic analyses for Tiguassu reginae which Righi et al. (1978) had placed in the Hapolotaxidae. In the analysis of Jamieson (1988b) Tiguassu proved paraphyletic relative to the Haplotaxidae sensu lato and formed the plesiomorphic sister-taxon of the Metagynophora. Its sole autapomorphy was restriction of the hearts to segment 10. The large proboscis-like prostomium (not computed) was a unique apomorphy in the entities included but is known homoplasically in the glossoscolecid Enantiodrilus bilolleyi Cognetti and is approached in some naids and lumbriculids. In its reproductive system (Fig. 8.28) Tiguassu provides evidence of reduction from two pairs of testes, and possibly from an ocotogonadal condition, in having two pairs of seminal funnels (in 10 and 11) of which those in 10 are vestigial in the absence of testes. Well developed testes are present in 11. The female system is progynous, as in most haplotaxids, with a single pair of ovaries in 12 immediately succeeding a testis-segment. There are no atria or other modifications of the male ducts, presumably as plesiomorphic conditions. There are two pairs of small,

!" Reproductive Biology and Phylogeny of Annelida

Fig. 8.28. Tiguassu reginae (Tiguassuidae). Reconstruction of the anterior 15 segments based on serial sections, showing the reproductive system with vestigial anterior seminal funnels. The smaller figure shows the proboscis-like prostomium. After Righi, G. et al. 1978. Acta Amazonica 8 (3 Supplement 1): 1-49, Figs. 3, 1.

adiverticulate spermathecae, in segments 9 and 10. The ova have a diameter of 40-50 µm (Righi et al. 1978). The co-occurrence of spermathecae with male funnels in 10 is a condition also seen in the Tubificidae, as is the great elongation of the seminal vesicle, but ovisacs were not found. Superorder Metagynophora. Loss of the anterior ovaries of a hypothetical octogonadal set, with retention of ovaries in segment 13 so that a segment lacking gonads intervenes between the posterior testes and the ovaries, or two segments in proandric taxa such as alluroidids, diagnoses all oligochaetes above the tubificid-enchytraeid assemblage and the Lumbriculidae, i.e. from the Moniligastridae through the Megascolecidae, loosely termed ‘megadriles’. This synapomorphy characterizes the Metagynophora of Jamieson (1988b) (Fig. 8.4). These are equivalent to the Lumbricida of Brinkhurst (1982). As the most plesiomorphic representatives, the Moniligastridae, Alluroididae and Syngenodrilidae, have not been sequenced for DNA, monophyly of the Metagynophora awaits confirmation from molecular analysis. Order Moniligastrida. Moniligastridae. Reproductive features among unambiguous synapormorphies for the Moniligastrida, as represented by Desmogaster and Moniligaster, are: ovaries in septal chambers; testis-sacs suspended on the posterior septum of the testicular segment; seminal vesicles absent; prostates capsular; spermathecae with non-seminal diverticula. Brinkhurst and Jamieson (1971) had already recognized the Moniligastrida as a separate order. Jamieson (1977b) re-interpreted the long debated nature of the testis-sacs, showing that they were neither reduced segments, as proposed by Stephenson (1922, 1930), nor intraseptal cavities, as argued by Gates (1962), but that they were normal testis-sacs which, with their enclosed testes, had become detached from the original testis-bearing septa (Fig. 8.29). It was recognized that moniligastrids are extraordinarily primitive in retaining a plesiopore condition, the extremely plesiomorphic

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Fig. 8.29. Distribution of genital organs in relation to existing segmentation and hypothetical segmental homologues in Moniligastridae. Origin of the ‘intraseptal’ testis-sacs from premoniligastrid sacs attached, with their testes, to the anterior septa of segments 10 and 11, as in other metagynophorans, is hypothesized. From Jamieson, B. G. M. 1977. Evolutionary Theory 2: 95-114, Fig. 3.

state (in Desmogaster) of 2 pairs of male pores in consecutive segments (both conditions seen elsewhere only in the Haplotaxidae) and a single layered clitellum with large yolked eggs. The moniligastrid clitellum is here shown to consist of a single layer of tall, slender modified epidermal cells with basal nuclei and dense granular secretory contents which discharge at the outer surface of each cell (Fig. 8.30A,B). They contrast with the wider, more robust goblet cells (putative large orthochromatic mucous cells) which predominate in the general epidermis (Fig. 8.30C). The reproductive system of a moniligastrid is here exemplified by that of Moniligaster troyi described by Jamieson (1977b). Details of the system are

!$ Reproductive Biology and Phylogeny of Annelida

Fig. 8.30. Light micrographs of the reproductive anatomy of Moniligaster troyi (Moniligastridae). A. Longitudinal section of the clitellum confirming that is consists of a single layer of cells. B. Same, through an intersegmental furrow. C. Longitudinal section of the general epidermis, showing the goblet cells. D. Longitudinal section of a testis-sac showing that it has anterior and posterior portions suspended in a septum. E. Morulae of spermatids in a testis-sac; a spermatid is labeled in a layer of spermatids encircling and attached to the cytophore. Abbreviations: c.m, circular muscle; clit.gl, secretory contents of a cell of the clitellum; cu, cuticle; cy, cytophore; go.c, goblet cell; l.m, longitudinal muscle; mo, morula of spermatids; sep, septum; spd, spermatid (in a layer of spermatids encircling and attached to the cytophore); te.s, testis-sac wall. From Jamieson, unpublished.

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illustrated from previously unpublished light micrographs (Figs. 8.30, 8.31). Testes and putative funnels are enclosed in a pair of diaphanous iridescent testis-sacs. Each sac is suspended in a septum so that it has pre- and postseptal portions (Figs. 8.29, 8.30D). The vas deferens from each testis-sac joins the sac ventrally at the anterior face of the supporting septum and passes into the anterior segment (segment 9) abutting the septum; it is very long and much coiled in this segment; numerous coils nearest the sac are narrow and iridescent but by far the greater length is wider and non-iridescent, with many hair-pin bends, and forms a large cluster. The vas deferens continues posteriorly to join the glandular portion of the prostates, in segment 11, considerably ectal of the ental end of the gland, and is straight in this segment. Immediately within the testis-sac the vas deferens gives rise to several iridescent ribbons which pass posteriorly for the entire length of the sac and were interpreted (Jamieson 1977b) as a backwardly directed sperm funnel. The testis-sac contains developmental stages of spermatozoa, including morulae of spermatids and free spermatozoa (Fig. 8.30D,E); it thus functions as a testis-sac and seminal vesicle. Each prostate extends from its pore, at 10/11 to intersegment 13/14; it has a clavate, superficially slightly lobulated glandular portion and a shorter, narrow duct which is poorly differentiated from the gland; the duct forms a muscular swelling at the pore which houses the base of the combined male and prostatic porophore (Jamieson 1977b) (Figs. 8.15, 8.31H,J). The wall of the gandular portion of the prostate consists of an outer thick longitudinal muscle layers, a thinner, though still thick, intermediate circular muscle layer, and an inner epithelium which contains gland cells (Fig. 8.31G,I). The ovary consists of folded (fan-like) laminae (Fig. 8.31F,H) on the anterior septum of its segment (11?). Oviducal funnels have yet to be recognized but large elongate ovisacs extend into segment 13 though arising from septum 11/12 against which septum 13/14 is adpressed; some lobules each contain a large-yolked egg (putative primary oocyte) with conspicuous nucleus. Moniligaster troyi has one pair of spermathecae, each with a large, elongate-ovoid ampulla in segment 8, its duct is long and much coiled in this segment but almost straight (Fig. 8.31C) on passing into segment 7 where it joins the apex of the wide, muscular ectal spermathecal duct (Figs. 8.14, 8.31A,B). The latter duct has two branches or horns, one on each side of the apex, each of which bears a large lobulated gland, the dichotomous gland; with the ectal spermathecal duct this constitutes the spermathecal atrium, discharging at intersegment 7/8 on each side. The spermathecal ampulla, in its ectal half, and its duct are exceptional for oligochaetes in being internally ciliated (Fig. 8.31A,B). The dichotmous gland consists of many blind tubules, opening into a common lumen; each tubule consists of a tall, glandular epithelium (Fig. 8.31C-E). Order Opisthopora. All remaining oligochaetes, above the Moniligastridae, from the Alluroididae to the Megascolecidae, form a convincing clade, the Opisthopora (see 8.1).

!& Reproductive Biology and Phylogeny of Annelida

Fig. 8.31 contd

Non-leech Clitellata

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Suborder Alluroidina. Superfamily Alluroidoidea. Syngenodrilidae. Apomorphies of the syngenodrilid reproductive system include presence of longitudinal tubercula pubertatis, intrasegmental testis-sacs, tubular prostate-like glands, all of which occur in other taxa, and a unique location of prostate pores in segments 11, 12 and 13. The male pores are lateral, and separate from the prostate pores, in segment 13. Genital and penial chaetae are present or absent. The clitellum begins in segment 11 and intraclitellar tubercula are present. Female pores lie in segment 14. Spermathecal pores are two pairs, posteriorly in segments 7 and 8. Testes are two pairs in 10 and 11 (rarely on segment more posterior), enclosed in the testis-sacs. The seminal vesicles are of a microdrile type, extending posteriad within the ovisacs through several segments. Alluroididae. Like the Syngenodrilidae, alluroidids represent an evolutionary transition in that they have the microdrile characteristic of a single layered clitellum but have attained the most plesiomorphic opisthoporan condition of male pores in segment 13, as in Righiella jamiesoni (Fig. 8.32A). In alluroidids the unilayered clitellum (Fig. 8.32D) commences on segment 12 or 13. Male pores are ventral to lateral in the chaetal arc of segment 13 or 14. The pair of female pores lies at or near the anterior border of segment 14. Spermathecal pores are paired, lateral, or are single, middorsal, in segments 6-9, maximally in three of these segments, never in line with the male pores. The male gonads are proandric, with testes in segment 10. In Kathrynella guyanae, all gonads are homeotically displaced one segment further posteriorly so that the testes are in 11. The sperm funnels have their mouths directed anterodorsally. Seminal vesicles project into the segment behind that of the testes or are absent; in the latter case spermatogenesis occurs in the testis-segment. Prostates (atria) are tubular or bulbous, receiving the male ducts, or discharging with the latter but separately from them, into a terminal chamber; they consist of an internal epithelium surrounded by a Fig. 8.31 contd

Fig. 8.31. Light micrographs of the reproductive anatomy of Moniligaster troyi (Moniligastridae), continued. A. Longitudinal section (LS) of a spermathecal ampulla. B. Same, showing internal ciliation. C. Passage of the straight region of the spermathecal duct through septum 7/8 into segment 7, where it joins (not shown) the dichotomous gland. Note lobes of the gland. D. Cross section through the dichotomous gland. E. Cross section through a single tubule of the dichotomous gland. F. Longitudinal section of the ovary, showing stages of oogenesis with terminal putative primary oocytes. G. Section through the prostate gland, showing the three layers of its wall, with inner gland cells. H. Section showing all regions of the prostates in segment 11: glandular region, duct and muscular swelling containing the common prostatic and male porophore. The ovary is visible (top left) in the following segment. I. Section of the glandular part of the prostate. J. Approximately horizontal section of the muscular swelling containing the common prostatic and male porophore. Abbreviations: ci, ciliation; c.m, circular muscle; di.g, dichotomous gland; gl.c, gland cell; l.m, longitudinal muscle; ov, ovary; p.o, putative primary oocyte; pr.d, prostate duct; pr. ep, internal epithelium of prostate gland; pr.g, prostate gland; pr. lu, lumen of prostate gland; pr.po, common prostatic and male porophore; sep, septum; sp.amp, spermathecal ampulla; sp.d, straight part of spermathecal duct. From Jamieson, unpublished.

! Reproductive Biology and Phylogeny of Annelida

Fig. 8.32 contd

Non-leech Clitellata

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muscular sheath outside which prostatic (atrial gland) cells are usually present. Ductules from the atrial gland cells penetrate the muscular sheath of the atrium, as in Alluroides brinkhursti brinkhursti (Fig. 8.32E,G), to reach the atrial lumen; an elongate penis, terminally containing a spermatozoal mass, may be present (Fig. 8.32F). Genital or penial chaetae are present or absent. Ovisacs extend posteriorly from the ovarian segment through several segments. Suborder Crassiclitellata. Crassiclitellate relationships are discussed under molecular phylogeny in 8.1.4 above (see also Fig. 8.4A,B). Biwadrilidae.The reproductive apparatus of Biwadrilus bathybates, illustrated by Nagase and Nomura (1937) (Fig. 8.33) lacks spermathecae, an absence shared with Criodrilus, in the Almidae (sensu Jamieson 1988b) and with Ocnerodrilus. In the case of Ocnerodrilus, at least, this appears to be a homoplasy. The male system is holandric, with testes in 10 and 11; testis-sacs are absent; seminal vesicles are two pairs, in segments 11 and 12. Vasa deferentia are intraparietal for much of their lengths, uniting only at the base of the conical male porophore, on each side on segment 13. The ventral chaetae of 13 are replaced by bifid genital chaetae. Prostate glands consisting of numerous lobules with branched ducts, bundles of ducts, and common ducts open into a male slit just ventral to each male pore. A large single ‘copulation gland’, resembling the chaetal gland of Microchaetus, is present on each side in 13, opening into the male slit ventrally to the prostate pores and just external to the genital chaetae; each gland has a terminal duct and a glandular portion consisting of outer peritoneum, a middle muscularvascular layer, an inner glandular layer with three types of cells, and a simple lumen. The lobed ovaries occupy the metagynophoran location of segment 13, with pores in 14. Ovisacs are restricted to segment 14 but an extensive subenteric (non-genital?) septal pouch (also seen in Microchaetus) may be present, arising further anteriorly. The location of the male pores, in segment 13, in Biwadrilus, only one segment behind the plesioporous location, is the most plesiomorphic condition for the Opisthopora and for the Crassiclitellata. It is shared with Fig. 8.32 contd

Fig. 8.32. Righiella jamiesoni (Alluroididae). A. Diagram showing arrangement of genital organs and vascular commissures. B. Transverse section (TS) of spermthecal duct. C. TS of prostate. After Omodeo, P. and Coates, K.A. 2000. Hydrobiologia 463(39): 39-47, Fig. 6. D-F. Alluroides brinkhursti brinkhursti. D. Transverse section (TS) of clitellum, showing single cell layer; the cells with conspicuous secretory granules and each with a basal nucleus. E. TS through the wall of the atrium, showinga group of atrial gland cells with ductule penetrating the muscular sheath of the atrium. F. Longitudinal section through the male pore, showing the ectal end of the atrium, which forms a penis with muscular sheath, ciliated epithelium and rope of spermatozoa in the lumen, forming in the ectal chamber a sperm mass. G. Alluroides pordagei. Oblique section through the atrial bulb, containing a large sperm mass, and the associated atrium. D-G. From Jamieson, unpublished figures from the study of Jamieson, B. G. M. 1971a. Alluroididae. Pp. 708-722. In R. O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh.

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Reproductive Biology and Phylogeny of Annelida

the non-crassiclitellate Alluroididae and the male genital systems of the two families show considerable similarities. Testing of phylogenetic proximity from molecular sequences would be desirable. Glossoscolecidae. The glossoscolecid clitellum is usually saddle-shaped and occupies as many as 15 segments, beginning near or shortly behind the female pores. Male pores are inconspicuous, one pair, rarely two pairs, intraclitellar or (Opisthodrilus) postclitellar. The female pores have the normal crassiclitellate location in segment 14 or exceptionally (Enantiodrilus) there are two pairs, in segments 13 and 14. The spermathecal pores are pretesticular, rarely extending into or behind the testis-segments; and in each intersegment occupied are usually a pair, though sometimes multiple, sometimes absent. Testes are one or two pairs, in segment 10 or segments 10 and 11; testis-sacs are present or absent. Copulatory sacs are present or absent. Spermathecae are absent (Glossoscolex, Fimoscolex, Goiascolex) or, usually, are present, when they extend freely into the coelom and are well differentiated into duct and ampulla or are intraparietal and poorly differentiated; they usually lack diverticula (Gates 1972; Jamieson 1971c; Righi 1995; Sims 1982). Tumakidae. Tumak hammeni (Fig. 8.35C,D) has a saddle-shaped clitellum commencing in segment 14 and occupies 9 segments. Male and female pores are microscopic, the female in the usual crassiclitellate location of segment 14, the male in 18 on the tubercula. Genital papillae surround the ventral setae (a and b separately) in segment 12 and throughout the clitellar region except in 17-20 where there is one pair of rectangular, tumid glandular pads in each segment; the pads on each side collectively considered to probably be homologous with the puberal bands (here termed tubercula pubertatis) of glossocolecids. We may also note the striking resemblance of the genital field to that of the microchaetid Michalakus (Fig. 8.35A,B). Testes and male funnels occupy segments 10 and 11, lacking testis-sacs; seminal vesicles are paired in 11 and 12. Tumak differs from the Glossoscolecidae in having intraparietal male ducts. The ovaries are large, folded and fan-shaped. Prostates and copulatory chambers are absent. The spermathecae are post-testicular, simple, two pairs in each of segments 12-14, lacking diverticula or seminal chambers and opening by microscopic pores in the corresponding anterior intersegments (Righi 1995). Eudrilidae. In eudrilids the male pores lie segment in 17, as is also typical of Ocnerodrilidae. Eudrilids differ from the Megascolecidae in having euprostates (Fig. 8.34), i.e. tubular prostates through which the male ducts discharge and which appear to be reflexed modifications of these ducts, thus more resembling the atria of lumbriculids and monilgastrids than the separate prostates (metaprostates) of megascolecids. However, the ectal ends of the vasa deferentia in some ocnerodriles are enlarged and somewhat resemble euprostates, though accompanied by tubular metaprostates. Eudrilids further differ from megascolecids, and ocnerodrilids, in migration of the spermathecae from the basic earthworm anterior location (in and/or anterior to segment 9) to the vicinity of the ovaries (in 13; sometimes posterior

Non-leech Clitellata

!!

Fig. 8.33. Biwadrilus bathybates (Biwadrilidae). Diagrammatic dorsal view of genitalia. Relabelled from Jamieson 1971c. Glossoscolecidae. Pp. 147-199. In R. O. Brinkhurst and B. G. M. Jamieson (eds), The Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto, Fig. 15.12A, After Nagase and Nomura.

to the male pore) and in the development in many of internal fertilization, foreign sperm passing internally from the spermathecae to ovisacs, on the oviducts, internally (see, for instance, Jamieson 1967, 1969; Sims 1967; Zicsi 1997). Elsewhere in the Oligochaeta, only the Phreodrilidae are suspected of having internal fertilization. The reproductive system in the Eudrilinae is more complex than that of the Pareudrilinae. In the latter transitions are seen in Stuhlmannia from

!" Reproductive Biology and Phylogeny of Annelida

Fig. 8.34 contd

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ovaries free in the ovarian segment, presumably with fertilization in the cocoon, to ovaries enclosed within the spermathecal system and with presumed internal fertilization. Penetration of the wall of the spermatheca by sperm from the partner, thus gaining access to the ovisacs has been demonstrated in Stuhlmannia variabilis by Jamieson (1958, 1967). Transition from free to enclosed ovaries is also seen in the pareudrilines Chuniodrilus and Scolecillus (see Jamieson 1969) (Fig. 8.34). Microchaetidae. In microchaetids the single pair of male pores is intraclitellar, behind segment 16, and female pores are on segment 14. The clitellum is saddle-shaped, beginning on segment 11 to 14 and occupying as many as 44 segments though sometimes a more modest six segments. Spermathecal pores are immediately postesticular or also occupy the last testis segment and are paired or multiple in each intersegment. Testes are in segments 10 and 11 or 10 only, in testis-sacs. Copulatory sacs and prostates are absent. The spermathecae do not project far into the coelom but are sometimes sinuous tubes. Tubercula pubertatis and/or genital papillae are present and have been illustrated by Plisko in several papers (e.g. Plisko 1996a,b) (see, for instance, Michalakus, Fig. 8.35). Lumbricidae. Lumbricidae, native in the Holarctic, are readily distinguished by location of the male pores, on 15, as in Lumbricus terrestris (Figs. 8.36, 8.53) or exceptionally 11, 12 or 13, well anterior to the clitellum. The clitellum is usually saddle-shaped, commencing between segments 17 and 52, and occupying 4-32 segments (Fig. 8.53). The spermathecal pores are preclitellar and usually paired, in two to eight of furrows 5/6-19/20, commonly in 9/10 and 10/11. There are two pairs of testes (Fig. 8.36, 8.37A) rarely one pair, in segments 10 and 11, usually free but occasionally in suboesophageal or perioesophageal testis-sacs. The vasa deferentia are Fig. 8.34 contd

Fig. 8.34. Genital anatomy of some Eudrilidae (Pareudrilinae), showing transition from free to enclosed ovaries with internal fertilization. A-E. Spermathecal and female genital systems in Chuniodrilus and Scolecillus arranged in order of increasing modification. A. C. ghabbouri. B. C. zielae. C. C. vuattouxi. D. C. compositus. E. S. tantillus. Note, in B-E, asymmetry of the oviducal system, with the ovisac vestigial on the left side (contrast Stuhlmannia). F, G. Stuhlmannia variabilis. Dorsal and lateral view of female reproductive system, respectively. Ovaries in the ‘coelomic tube’ discharge eggs into the ovisac on the right side, that of the left side being vestigial. Allosperm received into the spermatheca pass through the wall of the spermathecal atrium into the oviducal system where they are presumed to effect internal fertilization. H. Stuhlmannia asymmetrica. Here the oviducal system is developed on the left side only, having been totally suppressed on the right side. Sperm do not have to penetrate the wall of the spermatheca to reach the oviducal system as there is a wide, ciliated portal between the two. I. Stuhlmannia variabilis. Spermatophore redrawn after Beddard. A-E. After Jamieson, B. G. M. 1969. Journal of Natural History 3: 41-51, Fig. 1, After Omodeo 1958. Mémoires de l’Institut Français d’Afrique Noire 53: 1-109, and Wasawo, D. and Omodeo, P. 1963. Memorie del Museo Civico di Storia Naturale di Verona 11: 211-223. F-I. After Jamieson, B. G. M. 1967. Journal of Zoology, London 152: 79-126, Figs. 2, 3, 7 and 4 respectively. Abbreviations: cd, coelomic diverticulum. cs, coelomic sac; ct, coelomic tube; fp, female pore; o, ovary; oca, ovarian capsule; od, oviduct; of, oviducal funnel; ol, oviducal loop; os, ovisac; sa, spermathecal atrium; sam, spermathecal ampulla; sch, seminal chamber; sdiv, spermathecal diverticulum.

!$ Reproductive Biology and Phylogeny of Annelida

Fig. 8.35. A,B. Genital field of Michalakus initus (Microchaetidae). After Plisko, J. D. 1996. Michalakus, a remarkable new genus of microchaetid earthworm from South Africa (Oligochaeta: Microchaetidae). Annals of the Natal Museum 37: 287-293, Figs. 1,2. C,D. Tumak hammeni. C. Ventral view of segments 10 to 24, showing genital field. D. Spermathecae of segments 13 and 14. After Righi, G. 1995. Studies on Tropical Andean Ecosystems 4: 485-607, Fig. 201A,E.

extraparietal and sometimes coiled behind the seminal funnels to form epididymides. There are two to four pairs of seminal vesicles. Spermathecae are adiverticulate; they lack a distinct duct and are intraparietal, sessile or pedunculate. The ovaries, in segment 13, have a single egg string; each oviduct, discharging at a paired female pore in segment 14, bears a small ovisac (Bouché 1972; Gates 1976; Sims 1980). Kynotidae. The clitellum is annular or saddle-shaped in the region of segments 18-47. Tubercula pubertatis are absent. Male pores (clasper pores) are preclitellar, very conspicuous, on segment 16 or, rarely, 15, on a flat area or, on erection, on everted copulatory sacs (Fig. 8.37B). The spermathecal pores are post-testicular in the region of intersegments 13/14-16/17 and multiple in each row. Distinctive tubular prostate-like glands are associated with the copulatory sacs and with the follicles of preclitellar genital chaetae (Fig. 8.37C). The adiverticulate spermathecae are spherical to tubular (see review in Jamieson 1971c). Hormogastridae. In hormogastrids the male pores are intraclitellar, in the posterior half of segment 15, as in the type-species Hormogster redii (Fig. 8.39B), or, rarely, discharge on the tubercula pubertatis on 22 (as also in Ailoscolex). The clitellum is annular or saddle shaped, commencing on or near segments 12 or 14 and extends posteriorly for about 17 segments. The spermathecae are paired or multiple, in two to four intersegments, at the level

Non-leech Clitellata

Fig. 8.36. Lumbricus terrestris (Lumbricidae). Anatomy revealed by sagittal bisection. Original.

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!& Reproductive Biology and Phylogeny of Annelida

Fig. 8.37. A. Lumbricus terrestris (Lumbricidae). Diagram of the reproductive organs in dorsal view. Relabelled after Jepson, M. 1951. Biological Drawings. Part II. John Murray, London, p. 32. B, C. Kynotus cingulatus (Kynotidae). B. Ventral surface of segment 13-16, showing the pores of three pairs of prostates; a fourth pair discharges at the male pores. The clasper is shown evaginated through the left male pore (clasper pore). C. Internal view of the four pairs of prostates and the bursa propulsoria which contains the clasper. Each prostate is a convoluted tube enveloped in a sac. After Stephenson 1930. The Oligochaeta. Oxford. Figs. 145,146, from Benham.

of the genital segments. Testes are two pairs, in 10 and 11, or a pair in 11 only. Testis-sacs are absent but there are two pairs of seminal vesicles, in segments 11 and 12. Copulatory sacs and prostates are absent. Female pores lie in segment 14 (Bouché 1972; Sims 1980).

Non-leech Clitellata

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Lutodrilidae. Lutodrilus multivesiculatus is unique in the earthworms in having ten pairs of testes, in segments 12-21. It is considered to have interpolated ten segments, the last eight of them testicular, anterior to the normal megadrile location of testes in segments 10 and 11 (Jamieson 1978b). Lutodrilus stands apart from other almoids in having single-stringed ovaries, a feature clearly over-valued by Gates (1976) in aligning Lutodrilus with Lumbricus in his Lumbricoidea. The male pores are in segment 32; the pores discharge on a tumescence that encloses both ventral chaetal couples on 32 and 33. There is one pair of female pores, on segment 24. The clitellum is annular, only slightly swollen, and covers 37-51 segments, between segments 20 to 71. Alae about 1.5-3 mm high, extend through 16-32 segments, through segments 22 to 53 (Fig. 8.38A). Similar alae are also seen in the almoids Glyphidrilus (Fig. 8.38B), and, as segmentally less extensive claspers, in Drilocrius alfari (Fig. 8.38C) and Alma (Fig. 8.11A,B). Genital tumescences surround the ventral chaetal pairs in some of segments 13-51. The male tumescence is an elevated flattened area on the ventrum of 32-33, sometimes also 31 and/or 34. The ten pairs of testes are not enclosed in testis-sacs; each has several strings; vasa deferentia are intraparietal and prostates are absent. Seminal vesicles are largest in 14-22, attached to the posterior facesof their respective septa with the exception of the vesicles of 11 and 12 which attach to the anterior faces of septa 11/12 and 12/13 respectively. The ovaries are paired in segment 23, each with a single egg-string. The spermathecae are ovoidal and intraparietal in 2-5 of intersegments 15/16-25/26, that is, commencing in the gonadal region; they are multiple in each row; the external pores are not recognizable (McMahan 1976,1979). If the ten interpolated sections are deducted, comparison with other megadriles is facilitated and its closest relationship of seen to be with the Oriental Glyphidrilus and Ethiopian Callidrilus. Almidae. The reproductive anatomy of the Alminae will here be considered separately from that of the Criodrilinae. Alminae. In the Alminae genital chaetae, if present, are little if at all modified, except when on claspers. The male pores are one pair, on segments 15-30, always inconspicuous, intraclitellar or preclitellar. Female pores are on segment 14 but Glyphidrilus kukenthali is one of only three megadrile species known to have two pairs of female pores, in 13 and 14. Spermathecal pores are post-testicular (as in microchaetids), but are rarely continued into and anterior to the testis segments; they are sometimes (some Alma species) translocated into the hindbody; and are usually (with the spermathecae) multiple in an intersgement. Testes are paired in segments 10 and 11 or (Areco) 11 only. Prostate-like glands are rarely present. The paired intraceolomic parietal glands described by Righi et al. (1978) in some segments in Areco, although seen in some other almids, are reminiscent of the prostate-like glands of Sparganophilus. This endorses the view (Jamieson 1971b) that sparganophilids have a morphology close to that which might be attributed to proto-almids. A close relationship between Sparganophilus

!  Reproductive Biology and Phylogeny of Annelida

Fig. 8.38. Genital fields in Lutodrilidae and Almidae. A. Lutodrilus multivesiculatus (Lutodrilidae). Anterior end, with genital region, in ventral view, showing alae. After McMahan, M. L. 1979. Proceedings of the Biological Society of Washington 92(1): 84-97, Fig. 1. B. Glyphidrilus kukenthali (Almidae). Anterior end, with genital region, in ventral view, showing alae. C. Drilocrius alfari (Almidae). Anterior end, with genital region, in ventral view, showing claspers. C and D after Jamieson, B. G. M. 1971. Glossoscolecidae. Pp. 147-199. In R. O. Brinkhurst and B. G. M. Jamieson (eds), The Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto, Figs. 15.4B, 15.10A.

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and almoids (represented by Criodrilus and Lutodrilus) is not refuted by molecular data (Figs. 8.1, 8.6). Almines are notable for extensions of the body wall in the vicinity of or including the male pores. These extensions may be mere protuberances, as in some Drilocrius species; or involve a greater extent of the body wall, as in Glyphidrilocrius, or take the form of wing or keel-like structures (alae) in Glyphidrilus (Fig. 8.38B) or paddle-shaped claspers in Drilocrius alfari (Fig. 8.38C) and all species of Alma (Fig. 8.38A,B). In D. alfari, the male pores lie near the bases of the claspers but in Alma they are near the tips of the claspers which are furnished with genital chaetae and sucker-like structures (Figs. 8.11, 8.56). The structure of the clitellum of Alma emini has been described by Grove (1931) (Fig. 8.8D) and corresponds closely with that observed by the same author in the glossoscolecid Diachaeta exul, in the almids, Callidrilus ugandaensis by Jamieson (1971b), Glyphidrilus annandalei by Nair (1938) and Alma nilotica (Fig. 8.8E) by Khalaf El Duweini (1951) and in the sparganophilid Sparganophilus tamesis by Jamieson (1971b) (Fig. 8.8B). The clitellum of the biwadrilid Biwadrilus is similar but has, in addition to the fine- and coarse-grained cells, club-shaped peripheral cells with fine or coarse granules (Nagase and Nomura, 1937). In Criodrilus lacuum, Benham (1887) observed only glandular cells with small spherical globules. In Alma nilotica (Fig. 8.8E) the shortest (outermost) cells are normal epidermal supporting cells with a few sensory cells and mucin-secreting cells irregularly distributed amongst them. The cells that appear to make the middle layer are glandular cells which are fairly numerous and are irregularly distributed. They contain large granules and there is evidence that they secrete the cuticle and membrane of the cocoon. The apparent third layer is composed of cells appearing to form several tiers and arranged in groups that are separated from one another by thin lamellae of connective tissue. These contain fine granules of an albuminous secretion (Khalaf El Duweini 1951). The latter author, as did Grove (1931) for Alma emini and Grove and Cowley (1927) for Eisenia, presents evidence that the fine-granule cells secrete the albuminous contents of the cocoon. Grove (1931) considered this relative abundance of mucin-secreting cells in the clitellum of A. emini to indicate secretion of a copulatory slime tube. Their paucity in the clitellum of A. nilotica corresponds with the absence of a slime tube in this species. Criodrilinae. The Criodrilinae contain a single genus, Criodrilus (Fig. 8.19), including two species, the type-species Criodrilus lacuum and little know species inquirendae, C. ochridensis. In C. lacuum the ventralmost chaetae of at least segments 12, 13, 16-18 are modified as genital chaetae: terminally bearing four deep longitudinal grooves the proximal ends of which grade into irregular transverse jagged tooth-like ridges. The clitellum is indistinctly delimited anteriorly and posteriorly, annular, embracing 14, 15, 16 to 45, 47 (=30, 32, 34 segments). It consists histologically of an outer columnar epidermis continuous with that of the general body surface and three or four

!

Reproductive Biology and Phylogeny of Annelida

layers of club-shaped glandular cells with basal nucleus and filled with highly refractive small spherical globules. Male porophores are very strongly protuberant, transversely placed, ellipsoidal mounds filling, and widening, segments 15 and 16 longitudinally. Each male pore is a transverse cleft deeply bisecting the summit of the male porophore. There may be one to several spermatophores: curved, horn-shaped, hard but flexible structures approximately 1 mm long and maximally about 0.4 mm wide, at the expanded base; attached in the vicinity of the genital field. The female pores are each a small transverse slit in intersegmental furrow 14/15. Spermathecal pores are absent. The testes are free, digitate, or delicate, transversely slightly plicate lobes in segments 10 and 11; posterior to each is a much convoluted sperm funnel. Seminal vesicles are four pairs, in segments 9-12. The vasa deferentia are concealed deeply in the unusually thick body wall musculature, emerging in the coelom of segment 15 where that of each side of the body joins the anterodorsal aspect of a large hemispherical male bursa or prostate gland which is restricted to that segment. The gland consisting of cells similar to and continuous with those forming the epidermis of the clitellum; the muscular layers of the body wall covering the inner surface of the gland are thin; the vas deferens is continuous through the substance of the gland to the male pore. Each ovary is a solid, tongue-like or paddle-shaped lobe showing few external indications of oocytes, almost filling the length of segment 13. Oviducal funnels form small rosettes. Ovisacs, in segment 14, at maturity are at least as large as the ovaries and contain large oocytes; they project into 14 from septum 13/14 and are closely associated with but apparently not directly connected with the funnels (Jamieson 1971c). Ailoscolecidae. The male pores of Ailoscolex are intraclitellar, discharging on the tubercula pubertatis anteriorly in segment 22. The clitellum is annular, on segments 14-23 though incomplete ventrally in the first three segments. The tubercula pubertatis each consist of a gutter bordered dorsally by a pad and ventrally by the chaetal papillae, in segments 22-24. In Ailoscolex lacteospumosus the chaetal papillae form a row of contiguous tubercles from 14-24, of which the last three pairs are fused with the ventral aspect of the large tuberculum pubertatis (Bouché 1972) (Fig. 8.39A). Testes are paired, in 10 and 11; testis-sacs are absent; seminal vesicles lie in 11 and 12. Spermathecae are simple, very large, intracoelomic, pedunculate and globose, in segment 9 and 10. Prostate-like glands occur on the body wall, associated with the tubercula pubertatis, and radiate about a point of maximum density situated on intersegments 21/22-23/24. Ovaries are in segment 13, and large ovisacs in 14 (Bouché 1972). Ailoscolex appears to have close affinities with the family Komarekionidae (see below), which was subsumed in it by Sims (1980, 1982) and with the Sparganophilidae. Komarekionidae. This family is known from a single, terrestrial species, Komarekiona eatoni (Gates 1974), from North America. (Sims 1980, 1982) included Komarekiona in the Ailoscolecidae. There are striking similarities between the two entities, including the unusual location of male pores on

Non-leech Clitellata

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Fig. 8.39. Anterior ends, showing genital fields of A. Ailoscolex lactospumosus (Ailoscolecidae). B. Hormogaster redii insularis (Hormogastridae). After Bouché, M. B. 1972. Lombriciens de France: Écologie et Systématique, Institut National de la Recherche Agronomique, Vol. 72, Fig. 19.

segment 22; the long, saddle-shaped clitellum; tubercula pubertatis on the clitellum; the dorsolateral location of the spermathecal pores; the large number of tubular prostate-like glands associated with ventral chaetae; and the adiverticulate spermathecae. However, Komarekiona shows important differences from Ailoscolex which collectively are here considered to caution against synoymy in the Ailoscolecidae, although it must be admitted that variation of a similar magnitude occurs within other families, for instance the Megascolecidae. These differences are numbers of gizzards (single in segment 6, two, in 6-7 and 8-9, in Ailoscolex); absence of nephridial caeca and an intestinal typhlosole; a pretesticular (not testicular) location of the spermathecae; and presence of two pairs of latero-oesophgeal vessels which are not seen in Ailoscolex. In Komarekiona the clitellum is saddle-shaped, in segments 19-25 or 26, and bears ridge-like tubecula pubertatis. The male pores are inconspicuous, near the equator of segment 22. Spermathecae are adiverticulate, with pores

! " Reproductive Biology and Phylogeny of Annelida in 6/7-8/9. There are two pairs of testes, in segments 10 and 11. The vasa deferentia are supraparietal. Prostates are absent but prostate-like tubular glands, resembling those of Sparganophilus, are associated with the ventral chaetae in any of segments 7 to 26, those in 9-11 are larger. Additional, intraclitellar paired glands occur between the ventral chaetal pairs in some or all of segments 20-26. The ovaries have a single, terminal egg-string. Sparganophilidae. The male pores in Sparganophilus are one pair, inconspicuous in intersegmental furrow 18/19 or anteriorly in segment 19 (Fig. 8.40). The saddle-shaped clitellum is extensive, occupying eight to twelve segments in the region of segments 15-19. Tubercula pubertatis, in the

Fig. 8.40. Sparganophilus tamesis (=eiseni) (Sparganophilidae). A. Dorsal dissection. B. Anterior end, with genital region, in ventral view. After Jamieson, B. G. M. 1971. Glossoscolecidae. Pp. 147-199. In R. O. Brinkhurst and B. G. M. Jamieson (eds), The Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto, Figs.15.13C and B.

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clitellar region, are ridge like or a series of paillae, lateral to the ventral chaetal couples. Female pores are inconspicuous, in front of the ventral chaetal couples of segment 14. Spermathecal pores are inconspicuous, and dorsolateral, in 6/7-8/9, or 5/6 also, a single pair or four pairs per intersegment. Pores of postate-like glands, if these are present, are minute in the vicinity of the ventral chaetae in several segments in the clitellar region and sometimes in a variable number of more anterior segments. Testes and funnels are free in segments 10 and 11; seminal vesicles two pairs, in 11 and 12. Vasa deferentia are intraparietal. Ovaries are of the lumbricid type, i.e. with a single egg string, in 14; small ovisacs are present. Spermathecae are adiverticulate, paired or multiple, and extend far into the coelom (Jamieson 1971c). Resemblances to the Ailoscolecidae are noted under that family, above. Megascolecoidea. Families Ocnerodrilidae and Megascolecidae. This grouping is strongly supported by molecular data (Jamieson et al. 2002) (Fig. 8.1). Ocnerodrilidae. Relationship of ocnerodriles to the Megascolecidae has been widely accepted but they have been given subfamilial or familial status or even dispersed within the Megascolecidae (see Jamieson 1971d). Molecular analyses (Jamieson 2000; Jamieson et al. 2002) indicate that they are the plesiomorph sister-group of the Megascolecidae (Acanthodriliinae + Megascolecinae) (Fig. 8.1). They are divisible into two groups, ranking as subfamilies if ocnerodriles are given familial rank: the Ocnerodrilinae and a small group, the Malabarinae. The Ocnerodrilinae have extramural calciferous glands (esophageal diverticula) in segment 9; they occur from near the Tropic of Cancer in western North America through Central America and some Caribbean Islands into South America near the Tropic of Capricorn and throughout Africa from the Nile Valley and south of the Sahara, into Madagascar and the Seychelles. The Malabarinae lack extramural calciferous glands; they occur in the Indian subcontinent and Burma (Jamieson 1971d; Sims 1980, 1982). Ocnerodriles closely resemble Megascolecidae but differ from these in that calciferous glands, which are frequently absent from megascolecid species, are restricted to segment 9, or, in Malabarinae, 9 and 10. They are plesiomorphic relative to megascolecids in origin of the intestine in segment 12 (sometimes 13 or 14) and in not having added hearts behind segment 11 (Jamieson 1971d). With regard to reproductive anatomy, there are one to three pairs of tubular prostates with pores in the region of segments 16-21, of which one or two pairs are sometimes united with the male pores. Penial chaetae if, rarely, present are little modified. In some genera, including Eukerria (Fig. 8.9) (Jamieson 1970) the male pores and prostates are in the acanthodrilin arrangement which is typical of the megascolecid subfamily Acanthodrilinae. This genus alone was represented in the molecular analysis which confirmed sister-group relationship with the Megascolecidae (Jamieson 2000; Jamieson et al. 2002) (Fig. 8.6) and it would be desirable in

! $ Reproductive Biology and Phylogeny of Annelida further analyses to include species with the typical ocnerodrile arrangement of a pair of united prostatic and male pores on segment 17. The clitellum in Ocnerodrilidae usually occupies up to seven segments, between 12 to 18, but in Nematogenia it is 13 segments long and extends to segment 26. Spermathecal pores are, as in megascolecids, pretesticular but, unlike the latter, rarely bear diverticula. Whereas in Pygmaeodrilus nabugaboensis the spermathecal diverticula are inseminated (Jamieson 1957), in P. montiskenyae the ampulla receives the sperm (Jamieson 1965) (Fig. 8.17). Megascolecidae. Megascolecids usually have male pores on segment 18, fused with or near a pair of prostate glands, or prostates in 17 and 19 with male pores intermediate or fused with one pair of prostate pores. Different arrangements are shown and named in Fig. 8.8 Hoplochaetella is exceptional in having two pairs of male pores. Prostate glands are tubular to racemose (the latter with branched internal ducts, as in Pheretima). The vasa deferentia do not usually enter the glandular part of the prostate and they are therefore metaprostates and not euprostates. Spermathecae are usually diverticulate, rarely (Fig. 8.18) multiple. The evidence of the Ocnerodrilidae, which may have from one to three or more pairs of prostates, suggests that more than one pair of prostates were present in ancestral megascolecids. In the Megascolecidae, two pairs are still seen in the acanthodrilin condition, in which two pairs of prostate pores lie on segments 17 and 19, and the male pores are on segment 18, or the homeotic equivalent of these segments. Correspondingly, there are usually two pairs of spermathecal pores, at intersegments 7/8 and 8/9. This condition of the male terminalia is typical, though not constant, for the Acanthodrilinae (Fig. 8.8) and is seen, and probably of common derivation, in the Ocnerodrilidae such as Eukerria (Figs. 8.9, 8.54A). It is well exemplified by the genus Diplotrema, in which, as is usual for the acanthodrilin condition, the two prostate pores of each side communicate with the male pores by a seminal groove. Michaelsen (e.g. 1928) may well have been correct in proposing the acanthodrilin arrangement as basic to the Acanthodrilinae and that the microscolecin condition (Figs. 8.8, 8.54B) resulted by loss of the posterior prostates (in segment 19) and migration of the male pores into the vicinity of the anterior prostate pores (on segment 17), but a sexprostatic (with prostates in segment 18 also, as in Dichogaster damonis) or multiprostatic precursor (as in some ocnerodriles, see Jamieson 1958) cannot be ruled out. The microscolecin condition is seen, for instance, in Rhododrilus and, with complete fusion of male and prostate pores on segment 17, in Kayarmacia and in the circummundane parthenogenetic Microscolex dubius. The less common balantin condition (Figs. 8.8, 8.54C) was putatively derived by migration of the male pores onto segment 19 where they approached the single remaining, posterior, pair of prostate pores. This condition with male pores at intersegment 18/19 (and correspondingly a single pair of spermathecal pores, in 7/8) is seen in the Yucatan acanthodrile Balanteodrilus and was so derived by Pickford (1937) and is also seen in the New Zealand genus Sylvodrilus, in which the male pore remain on segment

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18 (Lee 1959). The most extreme balantin reduction is seen in the acanthodrile Torresiella from Horn Island, Torres Strait, Australia, in which the male pores and those of the single pair of prostates are united on segment 19 (with spermathecal pores in 7/8) and, as a further profound apomorphy, the nephridia are wholly meronephric (Dyne 1997). The strongly protuberant nature of the male pores in some acanthodriles suggests that in the acanthodrilin arrangement, the seminal grooves serve to pass prostatic secretion to the male pores rather than sperm to the prostate pores. However, the usual correlation of the number of pairs of spermathecal pores with the number of prostate pores might suggest the latter, commonly accepted, alternative. The megascolecin condition of the male pores (Figs. 8.8, 8.54D) is characteristic of the subfamily Megascolecinae. In the megascolecin condition, the male pores are united with the pores of a single pair of prostates on segment 18 or, in a presumably more plesiomorphic condition seen only in the New Caledonian genus Eudiplotrema, are near but not fused with the prostate pores (Jamieson and Bennett 1979). Michaelsen (1913) debated, and in Michaelsen (1928) remained equivocal, as to whether the megascolecin condition was acquired by migration of the remaining anterior or posterior pair of prostates of a former acanthodrilin condition onto segment 18. However, the possibility exists that the prostates of the megascolecin arrangement are plesiomorphically and intrinsically of that segment and, though not necessitated by this proposition, that they are persistent from a longer segmental series of prostates, possibly from a sexprostatic condition, with a pair of prostates in each of 17, 18 and 19. The latter, sexprostatic condition, though exceedingly rare, is known in the ocnerodrile, Diaphorodrilus doriae Cognetti (1910), the acanthodrile Pickfordia magnisetosa Omodeo (1958) and supposedly in the inadequately described Dichogaster damonis Beddard (1889b), from Fiji, the type species of Dichogaster, and has been reported in other Fijian and also in Caribbean Dichogasters (James, pers. com.; Jamieson et al., 2002) (Figs. 8.1, 8.6). The prostates are predominantly tubular in the Acanthodrilinae but they are tubuloracemose or even racemose in Dipotrema scheltingai and are racemose in Exxus Gates 1959.

8.3

OOGENESIS

Oogenesis in oligochaetes is intraovarian (Jamieson 1988a) (though considered extraovarian by Eckelbarger 1988) in that the germ cells are not released from the ovary into the coelom or a diverticulum of this, the ovisac, until they are ripe eggs (metaphase primary oocytes) and have completed vitellogenesis. Here they remain in metaphase of the first meiotic division and are released in this state from the female pores into the cocoon in which they are fertilized (see review by Jamieson 1981c). The size of the egg differs greatly between microdriles and crassiclitellates. Thus in microdriles the primary oocyte is very large, ranging from 300 µm to 1 mm, well within the

! & Reproductive Biology and Phylogeny of Annelida size range for so-called lecithotrophic eggs of polychaetes and other invertebrates, whereas in lumbricid earthworms primary oocytes reach 120 µm (see references in Jamieson 1988a), roughly in the range of planktotrophic eggs of polychaetes. The smaller size in megadriles is correlated with what is considered to be secondary acquisition of a multilayered clitellum, secretions of which, in the cocoon, reduce the necessity for reserves in the egg (Jamieson 1971b, 1988b). Intercellular bridges. Whereas connections between oocytes are rare in polychaetes (Eckelbarger 1988), in earthworms (Eisenia fetida) the oogonia and premeiotic primary oocytes are interconnected, each group developing from a single oogonium. The bridges each have the form of a fuzzy-coated zonula collaris as seen in spermatogenesis and at their confluence they constitute the cytophore. Homology of such structures between developing eggs and sperm is attributed to the hermaphroditic condition. It is inferred that the bridges permit synchronization of development of the gametes but the mechanism of information exchange is unknown (Jamieson 1988a). As shown for Enchytraeus and Tubifex, when the primary oocyte of an oligochaete is released into the coelom it is detached from the cytophore (see review by Jamieson 1981c). Vitellogenic Phase. Oligochaete vitellogenesis is already underway in the primary oocytes and in Enchytraeus albidus is said to begin in the third stage oogonium (Dumont 1969). It appears to be both autosynthetic and heterosynthetic; the distinction between the two being somewhat arbitrarily defined by the size of imported molecules. Heterosynthesis involves endocytosis (pinocytosis) as in polychaete eggs but some transference of yolk to the egg by chloragocytes occurs in enchytraeids (references in Jamieson 1981c). Such mechanisms for rapid incorporation of yolk precursors are characteristic of species having semi-continuous reproductive periods with short periods of oogenesis and frequent egg-laying (Eckelbarger 1988), as is true of oligochaetes, as opposed to monotelic species. For evidence for autoand hetero-synthesis of vitelline materials see (Jamieson 1981c, 1988a; Siekierska 2003). Cortical granules. Conspicuous cortical granules seen in some polychaete eggs, are not characteristic of oligochaete eggs. The cortical zone of the Tubifex primary oocyte in the ovisac is 2-3 µm thick, containing mitochondria and RER, ribosomes, minute vesicles, multivesicular bodies (often open to the surface) and other components in a finely particulate matrix devoid of granules and lipid droplets. Yolk granules and lipid droplets are confined to the endoplasm (Shimizu 1976). Egg envelopes. Oolemmal microvilli first appear in Eisenia in the primary oocytes where they project into a newly acquired acellular sheath, the zona pellucida (ZP). The ZP is regarded as a thickened oocyte membrane, the chorion, by Lechenault (1968) and, with the microvilli, would therefore constitute a primary envelope sensu Eckelbarger (1988). However, the origin of the ZP requires further investigation. The ZP and microvilli may jointly be termed the vitelline envelope. The microvilli in the mature ovarian oocytes are

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much shorter in microdriles (Enchytraeus albidus, Tubifex tubifex) than in crassiclitellates (Fig. 8.41). Their length has been shown in lumbricids to be strongly correlated with the length of the acrosome. They are illustrated for Allolobophora chlorotica in Fig. 8.41F,G and for Lumbricus rubellus in Figs. 8.41H. In tangential section of the ZP of A. chlorotica (Fig. 8.41F) it is seen that the microvilli are interrupted by circular fenestrae (Jamieson et al. 1983); these are perhaps equivalent to multiple micropyles. There is no ZP in tubificid oocytes but an indistinct fibrillar ‘vitelline membrane’ is traversed by the short microvilli. After fertilization the vitelline membrane of the Tubifex egg, becomes a trilaminar fertilization membrane overlying a perivitelline space. The oolemma loses its microvilli. The detached microvilli reach outwards only to the middle layer, suggesting (unless they have retracted) that the outer layer of the fertilization membrane is added from extrinsic sources, possibly from the cocoon fluid (Shimizu 1976, see Jamieson 1981c). Nurse cells (as sterilized oocytes), seen in polychaetes (Eckelbarger 1988), have not been reported in oligochaetes, excepting in a recent paper by Siekierska (2003). As only one or two embryos commonly develop in earthworm cocoons with sixteen or more eggs, the infertile eggs are effectively vitelline cells equivalent to those of neoophoran plathyhelminthes (Jamieson 1981c). Siekierska (2003), for the lumbricid Dendrobaena veneta reported the presence of nurse cells (trophocytes): the ovarian stroma is composed of somatic cells, the processes of which are connected to each other via numerous desmosomes; the somatic cells (identified as follicle cells sensu Jamieson) and their processes envelop the germ cells tightly and play a supportive role; oogonia, oocytes and trophocytes are arranged in distinct zones in the ovary; trophocytes form chains of cells, which are interconnected by intercellular bridges and contain numerous microtubules; the oocytes are distally arranged in the ovary. The function of the nurse cells in D. veneta has not been elucidated. There is evidence that trophocytes are mainly responsible for RNA (mainly rRNA) synthesis (references in Siekierska 2003), RNA being partly or fully synthesized in these cells and then transported to oocytes. They do not seem to be involved in vitellogenesis in D. veneta; the connections between oocytes and trophocytes no longer exist in ovarian zone III (mature oocytes) as the trophocytes degenerate (Siekierska 2003). That these trophocytes are distinct from follicle cells requires further investigation. Follicle cells. In Enchytraeus several layers of squamous epithelial cells (termed follicle cells by Jamieson 1981c but presumed to be modified peritoneal cells) cover the distal surfaces of the stage I and stage II oogonia. The stage III oogonium becomes covered distally and laterally by a thin, electron-dense layer derived from this epithelium by attenuation. Pillars of epithelial cytoplasm project towards the oogonial surface. Follicle cells also occur in lumbricids. In Eisenia fetida, they surround the primary oocyte at least while it retains its connection to the ovary. They are very much branched, with long slender processes forming several beds around the ZP, the more internal projecting into the latter. The projections in Enchytraeus and Eisenia do not establish connections with the egg. As

!! Reproductive Biology and Phylogeny of Annelida

Fig. 8.41 contd

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observed by Eckelbarger (1988) such follicle cells may have a supportive rather than nutritive role. In Enchytraeus, when the oocyte has entered the ovisac it has lost its follicle cells and this is presumably the case for the lumbricid egg (see reviews by Jamieson 1981c, 1988a, 1992). Oogenesis and phylogeny. The modes of oogenesis and vitellogenesis are of little value in determining the phylogenetic position of the Oligochaeta within the Annelida but they do not appear to show specializations which exclude regarding oligochaetes as being near the stem of the Annelida (Jamieson 1988a).

8.4 SPERMATOGENESIS AND SPERMATOZOAL ULTRASTRUCTURE (MARCO FERRAGUTI AND BARRIE G.M. JAMIESON) 8.4.1

Spermatogenesis

Spermatogenesis refers to the process of cell division and differentiation that commences with the primordial germ cells (protogonia) and ends with production of the mature spermatozoa. The latter stage of the process, whereby the spermatids differentiate without division to produce spermatozoa, is distinguished as spermiogenesis. The first spermatogonial divisions occur in the testis. Division is synchronous and results in the development of groups of cells interconnected by cytoplasmic bridges (collars in Ferraguti and Lanzavecchia 1971; bridges in Martinucci et al. 1977; zonulae collaris in Jamieson 1978a) to form morulae (e.g. Jamieson 1981c, 1992), also termed cysts (Ferraguti 1999). Collars surrounding the bridges are illustrated for mature spermatids of Haplotaxis ornamentus (Fig. 8.42H,I) and Eudrilus eugeniae (Fig. 8.43D). Further spermatogonial divisions, and spermiogenesis, may be limited to the coelom of the testis segment, as in Phreodrilidae, but usually occur in diverticula of the septa which project into adjacent segments and constitute the seminal vesicles. Development may also occur in specialized compartments of the testis segments, termed testis-sacs, present in combination with seminal vesicles, as in lumbricids and many megascolecids. The many variations on these themes are beyond the scope of this volume but details are given by Jamieson (1981c, 1992). Light microscopical observations on spermatogenesis in the megascolecids Amynthas hawayanus, A. morrisi and Metaphire californica are given by de Majo (2002a,b). The spermatogonia differ from oogonia in lacking smooth ER (present, however, in spermatids, see Boi et al. 2001), which is mostly situated Fig. 8.41 contd

Fig. 8.41. A-E. Acrosome and proximal portion of the nucleus of spematozoa of Lumbricidae in longitudinal section. A. Eisenia fetida. B. Lumbricus castaneus. C. Allolobophora longa. D. L. rubellus. E. A. chlorotica. F-H. Zona pellucida of unfertilized primary oocytes of Lumbricidae. F. Allolobophora chlorotica. In near-tangential section, showing fenestrae between the microvilli of the ZP. G, H. In vertical section, showing microvilli and portion of adjacent oocyte. G. Allolobophora chlorotica. H. Lumbricus rubellus. Based on the study of Jamieson B. G. M. et al. 1983. Gamete Research 8: 149-169.

!!

Reproductive Biology and Phylogeny of Annelida

Fig. 8.42 contd

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peripherally in the oogonia. The number of nuclei will increase stage by stage following powers of two. Thus morulae with 4, 8, 16, 32, 64, 128 (sometimes more) cells will be found. When the spermatogonial morula or follicle has reached the 8-32 cell stage, the region of confluence becomes a common cytoplasmic mass, the early cytophore. The spermatogonial morulae released from the testes into the coelom, or into modifications of this such as testissacs or seminal vesicles, possess only small cytophores (see Eudrilus eugeniae, Fig. 8.43A,B). In microdriles (Branchiura sowerbyi) (Hirao 1973) and earthworms (Lumbricus terrestris) (Walsh 1954) division of spermatogonia of the 16 cell morula gives the 32 primary spermatocytes. These are distinguished from spermatogonia, whose divisions are mitotic, in subsequently undergoing a meiotic reductional division, the first maturation division. In oligochaetes spermatocytes are smaller than spermatogonia, as there is a precocious enlargement of late spermatogonia. The products of reductional division of the diploid primary spermatocytes (usually 32) are haploid secondary spermatocytes (usually 64) (Hirao 1973). The cytophore further enlarges. The second non-reductional meiotic division of the haploid secondary spermatocytes produces similarly haploid spermatids, 128 in most of the oligochaete species studied (Jamieson 1981c), but 256 in two different lumbriculids (Ferraguti 1999) and more than 400 in some cysts, considered atypical, of the phreodrilid Astacopsidrilus (=Phreodrilus) (Jamieson 1981b). The spermatids transform into spermatozoa without further divisions (details in Jamieson 1981c, 1992; Ferraguti 1999). After detachment of the spermatozoa (Martinucci et al. 1977) or shortly before this in the tubificid Limnodriloides (Jamieson and Daddow 1979) and in Astacopsidrilus (Jamieson 1981b), the cytophore becomes globular (see Eudrilus eugeniae, Fig. 8.43C) and disintegrates. Functions of the cytophore. These are deduced to include support and synchronization of germinal cells through the interconnecting bridges. Boi et al. (2001) have shown that the bridges are kept open by actin rings. When the actin is de-polymerized by the end of spermiogenesis, some nuclei slip Fig. 8.42 contd

Fig. 8.42. Haplotaxis ornamentus. Spermiogenesis by transmission electron microscopy. A. Transverse section (TS) through condensing nucleus of spermatid, surrounded by the micrtoubular manchette. B. Golgi apparatus of spermatid, showing a Golgi lamella contributing to the developing acrosome vesicle.C. Acrosome developing an acrosome tube. D. TS distal centriole with satellite rays of anchoring apparatus. E. TS axoneme showing tetragon arrangement at central singlets. F. Developing acrosome and midpiece accompanied by Golgi apparatus. G. Spermatid nucleus surrounded by microtubular manchette. H. Longitudinal section of a younjg spermatid, showing collar attaching it to cytophore, Golgi apparatus, undondensed nucleus, proximal and distal centrioles, and flagellum. I. Part of a cytophore, containing rough endoplasmic reticulum (RER) stacks, and attached spermatids. J. Detail of RER. Abbreviations: at, acrosome tube; av, acrosome vesicle; co, collar attaching spermatid to cytophore; cy, cytophore; dc, distal centriole; f, flagellum; g, Golgi apparatus; gl, Golgi lamella; m, mitochondria of midpiece; mt, microtubules; n, nucleus; nm, nuclear membrane; pc, proximal centriole; rer, rough endoplasmic reticulum. From Jamieson, unpublished.

!!" Reproductive Biology and Phylogeny of Annelida

Fig. 8.43 contd

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inside the cytophore. The bridges possibly also play an active role in migration of cell elements into the cytophore as evidenced by the abundant cytoskeletal elements and mitochondria, as energy transducers, on the cytophore side. This includes selective intake of cell organelles not required for further spermiohistogenesis such as ribosomes, chromatoid bodies, fibrillar clumps, smooth endoplasmic reticulum, the Golgi body after it has completed its secretory phase, multivesicular and lysosomal bodies, and mitochondria other than the midpiece set. Nuclear projections into the cytophore may provide a means of discarding nucleoplasm or, it is speculated (Jamieson 1981c), transmitting genetic instructions into the cytophore. The large number of mitochondria in the cytophore suggests that it has an important function in providing energy to the developing spermatids as it seems unlikely that the mitochondria of the midpiece precursor fulfill the total energy requirements of these cells. The cytophore at the second meiotic division has a volume which is at least twenty-two times that of the spermatogonial stage (Eudrilus eugeniae, Fig. 8.43C). This is taken as evidence of much endogenous synthesis of materials. Organelles are, clearly, synthesized. Some of them, including RER, remain in the cytophore but it has been suggested that materials involved in cell morphogenesis (precursors of manchette microtubules and of sperm tails) and in energy production (glycogen of sperm tail) probably countermigrate into the spermatids. It is also suggested that the great increase in size, and synthesis of materials, of the cytophore are mediated by transcription products emitted from the nucleus in the cell cytoplasm at earlier, protogonial and spermatogonial, stages (Martinucci and Felluga 1975; Martinucci et al. 1977). The transcription products are possibly represented by the chromatoid bodies and fibrillar mitochondria-associated material Fig. 8.43 contd

Fig. 8.43. Eudrilus eugeniae (Eudrilidae). Stages in spermatogenesis by TEM. A. Morula of primary spermatocytes in metaphase. B. Morula of young spermatids, with some enlargement of the cytophore. C. Morula of elongated spermatids, with rounded cytophore (penetrated by a spermatozoon); spermatids with condensed nuclei in each of which an endonuclear canal has developed. D. Detail of late morula showing spermatid attached to the cytophore by a collar, microtubules surrounding the nucleus with its endonuclear canal. E. Transverse sections of nuclei of advanced spermatids and of a midpiece, both structures surrounded by microtubules. F. Longitudinal section (LS) of an elongating spermatid. The developing acrosome, being secreted by the Golgi apparatus, is still basal, near the distal centriole which has satellite rays. The mitochondria are assembling at the base of the nucleus. G. LS of an earlier spermatid. The acrosome rudiment shows a short acrosome tube and, within this, a shorter secondary tube. Mitochondria are assembling as the midpiece. The satellite apparatus of the distal centriole is visible as a ‘muff’ at the base of the axoneme. H. Later acrosome adjacent to the Golgi apparatus which has dense secretions at the ends of its lamellae. Beneath the dome-shaped acrosome vesicle there are two dense granules and a slender secondary acrosome tube. Outside the latter is the elongating acrosome tube. I. More advanced acrosome with more elongated acrosome tube. Abbreviations: at, acrosome tube; av, acrosome vesicle; ch, chromosomes; co, collar; cy, cytophore; dc, distal centriole; ec, endonuclear canal; f, flagellum; g, Golgi apparatus; m, mitochondria of midpiece; mt, manchette; n, nucleus; nm, nuclear membrane; sc, primary spermatocyte; sec, secondary tube; st, spermatid. From Jamieson, unpublished.

!!$ Reproductive Biology and Phylogeny of Annelida (Martinucci et al. 1977). Finally, the cytophore may play a part in separation of the spermatozoa and its own lysis (see reviews by Jamieson 1981c; 1992; Ferraguti 1999). Origin of early acrosome rudiments. The history of the complex and sometimes confused terminology of the components of the acrosome and of their development is treated in detail by Jamieson (1981c). Only a brief outline can be given here. The earliest anlage of the acrosome vesicle in oligochaetes originates from the Golgi apparatus of the spermatid, as the proacrosome. It becomes a small cap-shaped (concavoconvex) vesicle containing electron-dense material, the acrosome vesicle, as in Haplotaxis ornamentus (Fig. 8.42B,C,F). In all species studied it moves to the outer side of the Golgi under the plasmalemma, the latter becoming convex as a ‘’bleb”, over it. Additional Golgi vesicles may fuse with it during this migration (see review by Jamieson 1981c). Figure 8.42B is unique among published micrographs in showing an entire Golgi lamella fusing with the developing acrosome vesicle in H. ornamentus; this probably involves active movement of the lamellae. The second early anlage of the acrosome consists of one to several electron densities or granules, which develop immediately below the acrosome vesicle. A third structure which appears very early in development of the acrosome is the acrosome tube, as in Haplotaxis ornamentus (Fig. 8.42C,F) and Eudrilus eugeniae (Fig. 8.43G,H,I). This is a diagnostic autapomorphy of clitellates. It develops below the acrosome vesicle and, while growing in length, migrates with the vesicle to the tip of the nucleus which has undergone elongation and condensation. Its origin has been attributed, from micrographs, to the subacrosome granule(s), and/or directly to lamellae or vesicles originating from the Golgi apparatus (as in H. ornamentus, Fig. 8.42) and Limnodriloides or, questionably, to the proximal centriole (for a detailed discussion see Jamieson 1981c). A fourth major component of the acrosome complex termed the secondary tube (periaxial sheath), appears shortly before the axial rod makes its appearance. This is a short sleeve-like tube which appears in Lumbricus to develop from the base of the dense subacrosomal granule. It is attached as its apical end to the encircling rim formed by the bounding membrane of the primary acrosome vesicle and lies within the acrosome tube. Jamieson (1981c) proposed that in oligochaetes the acrosome vesicle, the dense granules, the acrosome tube and (indirectly) the secondary tube are all products of the Golgi apparatus but that the sequence of development of the granules and tube is variable. In the final stages of acrosome morphogenesis the acrosome vesicle is withdrawn except for its terminal domed tip, into the acrosome tube, or in enchytraeids, capilloventrids, and some tubificid species remains external. Differentiation of a fifth major component, the acrosome rod (axial rod), the putative perforatorium also occurs. This rod is already present inside the basal invagination of the vesicle in the earthworms but in the lumbriculid

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oligochaete Bythonomus lemani (see description below) differentiation of the rod occurs when the vesicle is already withdrawn. The rod probably develops from the dense granule(s) in all oligochaetes (see Jamieson 1981c, 1992; Ferraguti 1983, 1999). Migration of the acrosome from its origin at the Golgi apparatus to its emplacement on the tip of the nucleus occurs outside the microtubular manchette and without any association with microtubules (MTs). However, after its emplacement on the nucleus the manchette, which surrounds the nucleus, extends anteriorly to ensheath the acrosome tube and elongation of the acrosome occurs within the manchette. It seems possible that the MTs play a part in the later morphogenesis of the acrosome (see Jamieson 1981c). Some details of acrosome structure in the investigated oligochaete families are given below in 8.4.2. Nuclear morphogenesis. The nucleus of clitellate male germ cells is a long, filiform structure in the mature spermatozoa, but is rounded, as is the cell, in early, so-called isodiametric spermatids. Nuclear morphogenesis consists in the transformation of the nucleus from the spheroidal to the elongate, cylindroid form, and in the condensation of its chromatin, accompanied by major modifications of the microtubules present in the spermatid cytoplasm. The microtubules first appear, during early spermiogenesis in the vicinity of the distal centriole and later surround the nucleus as the manchette which is at first circumferentially discontinuous but becomes continuous (see Haplotaxis ornamentus, Fig. 8.42A,G; Eudrilus eugeniae, Fig. 8.43D,E). That the manchette controls nuclear elongation and chromatin condensation has been the subject of much debate (see, for instance, Fawcett et al. 1971; Ferraguti and Lanzavecchia 1971; Lora Lamia Donin and Lanzavecchia 1974; Webster and Richards 1977; Jamieson and Daddow 1979; Martinucci and Felluga 1979; Troyer 1980; Jamieson 1981c; Ferraguti and Ruprecht 1992; Ferraguti 1999). This subject cannot be revisited here. Suffice it to say that with the progress of spermiogenesis, the diameter of the nucleus decreases, as does the number of microtubules forming the manchette (Jamieson and Daddow 1979; Hodgson and Jamieson 1992). The microtubules discarded from the manchette are found in the cytoplasm of the spermatids, and are later discarded with the residual cytoplasm. Origin of the midpiece. Posterior to the nucleus, and approximately as wide as its base, are the mitochondria of the midpiece. The mitochondria are cristate but are radially adpressed; where several occur, they form a cartwheel configuration in cross section (Haplotaxis ornamentus, Fig. 8.42F; Eudrilus eugeniae, Fig. 8.43E). The fact that in ontogeny of the spermatozoon the midpiece originates as two to 11 separate, rounded mitochondria and that little elongation occurs in most tubificids suggests that a short untorted midpiece is plesiomorphic. There are seven mitochondria in the spermatozoon of Branchiobdella, and there is one in that of leeches. As four is the most common number in “primitive” sperm this has been somewhat arbitrarily assumed to be the basic number for oligochaete sperm, with divergence to two in Tubifex and a maximum of 11 in Capilloventer (see

!!& Reproductive Biology and Phylogeny of Annelida Jamieson 1981c; Ferraguti 1999). However, as Capilloventer appears to be the most basal oligochaete it is possible that many parallel mitochondria is the plesiomophric condition. Transformation of the centrioles. As reveiwed by Ferraguti (1999), two centrioles with the conventional ultrastructure of nine microtubular triplets are present only in the very early euclitellate spermatids (Jamieson 1981c), as here shown for Haplotaxis ornamentus (Fig. 8.42H). Soon after, the proximal centriole disappears. A remnant of the proximal centriole is said to make contact with the base of the nucleus just before the clustering of the midpiece mitochondria (Gatenby and Dalton 1959). The fate of the proximal centriole has never been followed in detail, but its topographical, if not causational, importance as a center for aggregation of the midpiece mitochondria has been repeatedly demonstrated (Ferraguti and Jamieson 1984). Very early in spermiogenesis the distal centriole produces the flagellum when it is connected to the plasma membrane of the spermatid through a complex nine-rayed structure, as in Haplotaxis ornamentus (Fig. 8.42D) and Eudrilus eugeniae (Fig. 8.43F), identical with the anchoring apparatus of ‘primitive’ spermatozoa, i.e. those fertilizing in sea water. The anchoring apparatus disappears progressively as spermiogenesis continues. It has been supposed that the anchoring apparatus in mature spermatozoa loses its function, since the basal body is constrained by the midpiece (Ferraguti 1984a,b). In mature sperm the remnants of the anchoring apparatus assume the shape of a ring or a cylinder of dense material placed under the plasma membrane (annuloid in Jamieson 1982) which may involve the basal body microtubules, as in tubificid oligochaetes and in branchiobdellids or be discontinuous and irregular as in leeches or in megascolecid oligochaetes (Jamieson 1978a). The basal body never shows a conventional triplet appearance in the mature oligochaete spermatozoon. Tannic acid treatment reveals microtubular doublets surrounded by dense material (Ferraguti and Gelder 1991). In the oligochaetes the basal cylinder is a structure with a diameter ranging from 60 to 100 nm and a length from 0.1 to 0.3 µm from which the central apparatus of the flagellum emerges (Ferraguti 1999). It makes its appearance in the early spermiogenetic stages and elongates to fill, to a variable extent, the basal body (Ferraguti 1984a). In branchiobdellids, hirudineans, and acanthobdellids, all lacking a basal cylinder, the basal body is progressively penetrated by the central apparatus of the axoneme from the early spermiogenetic stages. The central apparatus reaches the mitochondrion in acanthobdellids and leeches, or even penetrates into the midpiece axis, as in branchiobdellids. Modifications of the flagellum. The flagellum of the early euclitellate spermatids has a conventional 9+2 appearance. During spermiogenesis, the basic structure is modified by the appearance of glycogen granules surrounding the axonemal doublets, and by the transformations of the central apparatus, as in Haplotaxis ornamentatus (Fig. 8.42E). The glycogen

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granules appear in the mid-spermatids to surround the axoneme for almost its complete length, so that only the terminal portion of the euclitellate flagella has the conventional 9+2 appearance. The central apparatus of the flagellum may be modified by ‘tetragon fibers’ (two fibers at right angles to the two central singlets) (Fig. 8.42E) or by the prominent central sheath. In some oligochaete species, the tetragon fibers grow into the central apparatus of mid-spermatids (Jamieson 1981a); in others, the tetragon fibers are replaced (or embedded?) in later stages by the prominent central sheath. This process has been followed in detail in the tubificid Monopylephorus limosus (Ferraguti 1999) and proceeds from the basal towards the distal portion of the flagellum. In some oligochaete species (for instance in the tubificid Coralliodrilus rugosus, see Erséus and Ferraguti 1995) a mixture of the two modifications of the central apparatus is present in the mature spermatozoa: the basal portion of the flagellum shows a prominent central sheath appearance, whereas a more distal portion shows the tetragon fibers, and a short final tract has a 9+2 appearance without any modification of the central apparatus. The branchiobdellids, acanthobdellids, and leeches, show the progressive involvement of the central tubules by the prominent central sheath (Ferraguti and Lanzavecchia 1977).

8.4.2 Mature Spermatozoa The Clitellata (Oligochaeta sensu lato) possess only introsperm (terminology of Rouse and Jamieson 1987), i.e., sperm that are not introduced into the ambient water if the species is aquatic and are usually (obligatorily in leeches) internally fertilizing, though in most oligochaetes fertilization occurs external to the body, in the cocoon. The spermatozoa of the Clitellata are filiform cells characterized by (1) the presence of an acrosome tube containing and/or supporting the acrosome vesicle which is often withdrawn into it; (2) interpolation of the mitochondria between the nucleus and the basal body of the flagellum; (3) peculiar modifications of the central apparatus of the axoneme and (4) two glycogen granules per doublet cross section (Jamieson 1981c). The oligochaetes sensu stricto are diagnosed by the presence of a basal cylinder situated within the basal body which is lost in brachiobdellids and leeches (Ferraguti 1984a,b; Ferraguti and Erséus 1999); the branchiobdellidans by an apical, conical indentation of the nucleus and by a helical marginal fiber coiled around the tail; the acanthobdellids by a dense sheath and accessory fibers surrounding the axoneme; the euhirudineans by an anterior prolongation of the acrosome tube (Jamieson 1978c, 1981b,c, 1982, 1983a,b,c, 1984, 1986, 1987, 1988a ,b, 1992; Jamieson et al. 1978, 1982, 1987; Ferraguti 1983, 1999; Ferraguti and Erséus 1999). Some caveats apply to these characteristic features. The acrosome tube though highly distinctive, being unknown in polychaetes, occurs convergently, though only superficially similar, in nematomorphs (Valvassori et al. 1999). It is reduced or possibly absent in Bathydrilus formosus, a phallodriline tubificid (Ferraguti et al. 1989). Interpolated mitochondria also occur in Onychophora, with possible

!" Reproductive Biology and Phylogeny of Annelida phylogenetic implications (Jamieson 1986) and, with no phylogenetic significance, in Chondrichthyes (Jamieson 1991) and in some gastrotrichs (Ferraguti et al. 1995). Similarly distributed glycogen is seen, convergently, in the polychaete Micromaldane (Rouse and Jamieson 1987). The clitellate spermatozoal synapomorphies of possession of an acrosome tube and interpolation of the midpiece between nucleus and distal centriole do not occur in Aeolosoma and Potamodrilus, thus confirming that aphanoneurans are not clitellates (Bunke 1985, 1986). A combined immunohistochemical and ultrastructural investigation of the central nervous system and the sense organs in Aeolosoma hemprichi (Hessling and Purschke 2000) also indicated that aphanoneurans could not be included in the Clitellata. Some molecular cladograms based on three combined genes, two nuclear (18S, 28S) and one mitochondrial (COI), gees placed Aeolosoma as the sister-taxon of the Clitellata (Hugall et al. unpublished) (Fig. 8.5). However, Struck et al. (2002) found that 18S rDNA sequences did not unequivocally support a sister-group relationship of Aeolosoma sp. and the Clitellata. Instead, depending on the algorithms applied, Aeolosoma clustered in various clades within the polychaetes, for instance, together with eunicidan species, the Dinophilidae, Harmothoe impar or Nereis limbata. Although spermaozoal ultrastructure asserts monophyly of the Clitellata, it does not prove monophyly of the oligochaetes sensu stricto and, though not conflicting with monophyly of the latter, it could equally support the view that oligochaetes sensu stricto are a paraphyletic congeries, as indicated by molecular data (see 8.1.2 above). The constantly present basal cylinder, though distinctive of oligochaetes, is presumably plesiomorphic. A generalized, plesiomorphic oligochaete spermatozoon is illustrated in Fig. 8.44 to aid in understanding of the components of a mature spermatozoon. Regarding major departures from general oligochaete sperm characteristics, only the eudrilid Eudrilus eugeniae (Jamieson and Daddow 1992) and the tubiificid Rhizodrilus russus (Ferraguti et al. 1994) are known to have an endonuclear canal. In members of the subfamily Tubificinae, and some Limnodriloidinae, a double sperm line produces euspermatozoa and paraspermatozoa (Braidotti et al. 1980; Braidotti and Ferraguti 1982, 1983; Ferraguti et al. 1983, 1988, 1989, 1994, 2002b; Ferraguti and Ruprecht 1992; Boi et al. 2001; Marotta et al. 2003). An account follows on the ultrastructure of spermatozoa in those oligochaete families for which it has been investigated. Enchytraeidae. Lumbricillus rivalis has a simple, short acrosome, with the vesicle external to the tube, a well developed secondary tube, a stout rod, a basal chamber, and a limen (Webster and Richards 1977) (Fig. 8.45A). These characters have been considered to make the enchytraeid acrosome the most primitive oligochaete spermatozoon examined (Jamieson 1983a), though this status is now questionable in view of molecular phylogeny discussed above in 8.1.4. The nucleus is apically flanged and basally straight; the four mitochondria (six in one Mesenchytraeus species, Ferraguti and Fender unpublished) are twisted; the flagellum has a long basal cylinder and an

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Fig. 8.44. Diagrammatic longitudinal section of generalized, plesiomorphic oligochaete sperm to illustrate chief components. From Jamieson, B. G. M. et al. 1987. Cladistics 3(2): 145-155.Fig. 3.

axoneme with a tract with a prominent central sheath followed by one with tetragon fibers (Fig. 9 in Webster and Richards 1977). A profound comparative study on 19 different populations and species of Enchytraeus (Westheide et al. 1991) has confirmed the above description, except for the absence or extreme reduction of the basal chamber in the acrosome and for the nucleus being flanged (or corkscrew-shaped) for the whole length. Significant metric differences allowed the identification of the various populations and species.

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Reproductive Biology and Phylogeny of Annelida

Fig. 8.45. Longitudinal section of a spermatozoon by TEM of A. An enchytraeid (Lumbricillus rivalis). B. A tubificid (Rhyacodrilus arthingtonae). 1. acrosome vesicle, 2. subvesicular space, 3. acrosome tube, 4. secondary tube, 5. axial rod (perforatorium), 6. nuclear pad, 7. nucleus, 8. capitulum, 9. connective, 10. midpiece, 11. proximal core of axoneme. From Jamieson, B.G. M. 1983. Zoologica Scripta 12: 107-14, Fig. 1.

Non-leech Clitellata

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The morphometric and qualitative spermatozoal data were used (Westheide et al. 1991) as morphologic taxonomic characters, as discussed in 8.1.5. Their analysis confirmed the value of sperm ultrastructure for solving taxonomical problems at the species level (Westheide et al. 1991). Capilloventridae. The acrosome of the spermatozoon of the Australian freshwater Capilloventer australis (Fig. 8.46) is similar to that of enchytraeids, and putatively plesiomorphic, in having the vesicle completely external to the tube, but differs in lacking a secondary tube and a limen. The nucleus is again similar to that of enchytraeids, but C. australis has the condition, known in no other clitellate, of eleven mitochondria arranged longitudinally in parallel (not radially) in the midpiece. It is uncertain whether this is autapomorphic or plesiomorphic. The basal cylinder is very long, as in Enchytraeidae, but the tail shows only the prominent central sheath pattern (Ferraguti et al. 1996). Phreodrilidae. The spermatozoon (Fig. 8.47A) of Astacopsidrilus (=Phreodrilus) jamiesoni, epizoic on an Australian freshwater crayfish (Jamieson 1981b), has a long undulating acrosome tube, with the vesicle only partly withdrawn, a small acrosome rod (perforatorium), a short secondary tube, a large basal chamber, a putative nuclear pad, an undulating nucleus, six helical mitochondria forming the unusually long (9 µm) midpiece, and a tail with prominent central sheath basally and tetragon fibers distally. The acrosome resembles that of some crassiclitellates, thus falling among them in the phylogenetic analysis. The sperm of Insulodrilus bifidus is similar in most respects: even longer midpiece; nuclear shape; six mitochondria; prominent central sheath and and tetragon fibres but the acrosome tube is less that 1 µm long, slightly bent to one side, has a limen, and a short basal chamber and the vesicle is completely withdrawn (Marotta, pers. comm.). Phreodrilids are suspected of having internal fertilization, which is rare among oligochaetes, occurring elsewhere in Eudrilidae (Jamieson et al. 1987), and this possibly accounts for the unusual acrosomal morphology in both taxa. Tubificidae. This account is drawn from the review of Ferraguti (1999). The Tubificidae is the most speciose microdrile family (about 600 species) and shows the greatest diversity in sperm morphologies. Variations concerning all the characters were reviewed in detail for 17 species by Erséus and Ferraguti (1995) and Ferraguti et al. (1994). A list of all examined species is given in Ferraguti (1999). A generalized tubificid sperm based on Limnodriloides australis is illustrated in Fig. 8.45B. The acrosome tube may be straight as in Tubificinae and Limnodriloidinae, or bent, as in the rhyacodriline Monopylephorus limosus and in the gutless phallodrilines, or twisted, as in the phallodriline Thalassodrilus prostatus. The acrosome tube has, in M. limosus, a crystalline appearance in negative staining (period of 4.7 nm). The vesicle can be completely external, as in Thalassodrilus prostatus or in Limnodriloides sp., partly withdrawn, as in Rhyacodrilus arthingtonae (Jamieson et al. 1978) or in the Tubificinae, or completely withdrawn, as in M. limosus. A large basal chamber is present, as in Thalassodrilus prostatus or absent, as in M. limosus.

!"" Reproductive Biology and Phylogeny of Annelida

Fig. 8.46. Capilloventer australis (Capilloventridae). Ultrastructure of the spermatozoon. From Ferraguti et al. 1996. The spermatozoon of Capilloventer australis and the systematic position of the Capilloventridae (Annelida: Oligochaeta). Australian Journal of Zoology 44(5): 469-478, Fig. 1.

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Fig. 8.47. Longitudinal section of a spermatozoon by TEM of A. A phreodrilid (Astacopsidrilus (=Phreodrilus) jamiesoni). B. A haplotaxid (Haplotaxis ornamentus). C. A sparganophilid (Sparganophilus tamesis). D. A hormogastrid (Hormogaster redii). E. A lumbricid (Lumbricus rubellus). F. A megascoelcid (Fletcherodrilus unicus). G. A megascolecid (Amynthas gracilis). 1. acrosome vesicle, 2. subvesicular space, 3. acrosome tube, 4. secondary tube, 5. axial rod (perforatorium), 6. nuclear pad, 7. nucleus, 8. capitulum, 9. connective, 10. midpiece, 11. proximal core of axoneme. From Jamieson, B. G. M. 1983. Zoologica Scripta 12: 107-14, Fig. 2.

Some acrosome tubes have a limen, others are simple. In Tubifex tubifex eusperm, and in M. limosus, the plasma membrane at the level of the tube shows a characteristic array of particles. In tubificid sperm, the nucleus is never straight for its whole length, but can be in part straight, or twisted, or corkscrew-shaped, or flanged , or with regions with different shapes. The midpiece mitochondria vary in number from two (Tubifex tubifex) to five (all the Phallodrilinae), and in shape from hemispherical (in T. tubifex) to radial sectors of a cylinder, as in Clitellio arenarius. When elongated, the mitochondria can also be helical, as in Pectinodrilus molestus or in Coralliodrilus rugosus. The axoneme can have the prominent central sheath, or the tetragon fibers, or both in sequence (Erséus

!"$ Reproductive Biology and Phylogeny of Annelida and Ferraguti 1995). The plasma membrane of tubificid sperm tails is characterized by the possession of zipper lines, i.e. nine parallel double rows of particles at the level of the axonemal doublets (Ferraguti et al. 1991). Besides the zipper lines all rhyachodriline sperm tails, except in Rhizodrilus russus, also have “muffs” (Ferraguti et al. 1991, and Marotta, unpublished), double rows of particles running perpendicular to the zipper lines. The interplay between zipper lines, muffs, and other membrane structures gives rise to a complex model of the periaxonemal area. The considerable variation of sperm models within tubificids may be the consequence of the great species differentiation within the family. However, no other microdrile family has been analyzed in such detail. A high amount of structural homoplasy was detected in a cladistic analysis (Erséus and Ferraguti 1995). Naidinae. The spermatozoa of the naidines Paranais litoralis, Nais communis and Slavina appendiculata (Gluzman 1998, 1999), Stylaria lacustris and two species of Paranais (Ferraguti et al. 1999) have the usual oligochaete sperm components Paranais litoralis has a straight nucleus, five midpiece mitochondria and a complex flagellum with prominent central sheath. Nais communis, on the other hand, has a twisted nucleus and, questionably, only one mitochondrion. The spermatozoon of Slavina appendiculata has a ‘corkscrew’ nucleus, 5 or 6 mitochondria in the usual radial arrangement. The axonemes have the tetragon fiber configuration (Gluzman 1998, 1999). In Stylaria lacustris (and two species of Paranais, see Ferraguti et al. 1999) the acrosome is short and straight, with the vesicle withdrawn, no secondary tube, and the acrosome rod barely visible. The nucleus is twisted for most of its length, but basally straight. There are five parallel mitochondria, and a tail with prominent central sheath basally and tetragon fibers distally. This description is confirmed for Unicnais uncinata (Marotta, pers. comm.). The ultrastructure of the naid spermatozoon is consistent with inclusion of Naididae within the Tubificidae (Erséus 1990), as the Naidinae (Erséus and Gustavsson 2002). Lumbriculidae. Ultrastructural descriptions exist for the spermatozoa of Bythonomus lemani (Ferraguti and Jamieson 1987) (Fig. 8.48), Kinkaidiana sp., Rhynchelmis limosella, R. alyonae (Martin et al. 1998), R. brachicephala (Ferraguti et al. 1999) and an undetermined species from Lake Baikal (Ferraguti 1999). The acrosome is short, bent to one side, with the vesicle deeply withdrawn, well developed rod, secondary tube with possible connections to the rod, no basal chamber, no limen, and possibly the tube closed at its base (or a nuclear pad fused with the tube?). The nucleus is twisted for the whole length (Bythonomus and Rhynchelmis alyonae) or apically corkscrew-shaped and basally straight (Kinkaidiana), or completely straight as in R. brachicephala. In all Rhynchelmis species examined so far there is a deep apical concavity at the apex of the nucleus. In Kinkaidiana sp. there is also an apical concavity reminiscent of that present in most Branchiobdellidae and, possibly, in some leeches, a feature consistent with molecular phylogeny. There are six, highly twisted mitochondria in the lumbriculid midpiece, and a prominent central

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Fig. 8.48. Bythonomus lemani (Lumbriculidae). Longitudinal section of a spermatozoon by TEM. From Ferraguti, M. and Jamieson, B. G. M. 1987. Hydrobiologia 155: 123-134, Fig. 23.

!"& Reproductive Biology and Phylogeny of Annelida sheath in Rhynchelmis and Kinkaidiana, but a sequence of prominent central sheath and tetragon fibers in the axoneme in Bythonomus lemani. Haplotaxidae. The spermatozoa of the Tasmanian Haplotaxis ornamentus (Jamieson 1982) (Fig. 8.47B) and Pelodrilus leruthi, a subterranean species living in the Pyrenées, are substantially similar to each other (Ferraguti 1999). There is a long acrosome tube, and a comparatively small vesicle, only partly withdrawn into the tube. A thin acrosome rod with an apical enlargement (capitulum sensu Jamieson 1978a) is housed in the subacrosomal space, leaving a large basal chamber. A secondary tube is connected to the rod in Haplotaxis, but not so evidently in Pelodrilus. The tube ends with a limen in Pelodrilus, but is possibly closed in Haplotaxis. A flat nuclear pad separates the acrosome tube from the straight nucleus. Six mutually parallel mitochondria follow, and a flagellum with tetragon fibers. The two species examined are distant in space and phylogeny, thus the haplotaxid sperm models appear remarkably uniform. Sparganophilidae. The acrosome tube of the spermatozoon of Sparganophilus tamensis, investigated by Jamieson et al. (1982) (Fig. 8.47C), is characterized by a long basal chamber ending proximally against a large limen and distally with the basal extremity of the vesicle. This vesicle is only partly withdrawn into the tube, projecting anteriorly as a bleb. The acrosome rod is contained within the subacrosomal space and partly projects outside the tube anteriorly. A short secondary tube surrounds the base of the rod. In Sparganophilus, as in the ‘higher’ families described below, the secondary tube has two clearly distinct parts: a cylinder surrounding the proximal extremity of the rod, and an oblique connection between the cylinder and the proximal extremity of the acrosome vesicle. A nuclear pad with a central boss separates the acrosome from the straight nucleus. The midpiece is formed by eight parallel mitochondria, two of which are in line, one over the other. The flagellum has the typical tetragon fibers of the megadriles, as well as the usual glycogen granules. Ocnerodrilidae. The spermatozoon of the peregrine ocnerodrilid species Nematogenia panamensis is 35 µm long and shows the conventional clitellate sequence of acrosome, nucleus, middle piece and tail. The acrosome is asymmetric, showing an acrosome rod crossing the vesicle to nearly touch the tube, and re-crossing the vesicle anteriorly. This condition is unique among investigated euclitellates, as is the structure of the acrosome tube, which seems to be decorated by longitudinal spiral furrows. The secondary tube and the limen are similar to those of other earthworms. The nucleus is straight as are the six midpiece mitochondria. The flagellum has one of the two usual oligochaete arrangements: a 9+2 axoneme with two central tetragon fibers, surrounded for most of its length by glycogen granules. The secondary tube and the limen are similar to those of earthworms (Bondi et al. 1993; Ferraguti et al. 1999). While the general features of Nematogenia spermatozoon are undoubtedly of crassiclitellate type, characters such as the shortness of the acrosome and the basal chamber were considered to indicate a plesiomorphic condition within the group.

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Eudrilidae. Only a single species of the Eudrilidae has been examined for spematozoal ultrastructure, the circummundane Eudrilus eugeniae (Jamieson and Daddow 1992) (Fig. 8.49). Spermatogenesis in this species is illustrated for the first time in Fig. 8.43). This is an internally fertilizing species and investigation of species which extrude spermatozoa from spermathecae into the cocoon in the usual oligochaete mode is desirable. The

Fig. 8.49. Eudrilus eugeniae (Eudrilidae). Highly diagrammatic representation of the ultrastructure of the spermatozoon by TEM. From Jamieson, B. G. M. and Daddow, L. 1992. Journal of Submicroscopic Cytology and Pathology 24(3): 323-333, Fig. 2.

!# Reproductive Biology and Phylogeny of Annelida Eudrilus spermatozoon shows the following unique or unusual features relative to other oligochaetes: 1) The acrosome tube bears externally a spiral ridge or flange (also seen in some other oligochaetes, e.g. Coralliodrilus, Ferraguti and Erséus, 1999). The tube and its extension also bear helical ridges in leeches and some branchiobdellidans; it has a spiral tendency in lumbriculid and phreodrilid sperm (Jamieson 1981c; Ferraguti and Gelder 1991). The undulating nucleus and helical midpiece are unusual among earthworms. 2) The acrosome tube is greatly elongated, at 7.7 µm, though shorter than the maximum in other oligochaetes of 12.7 µm, reported for Allolobophora chlorotica by Jamieson et al. (1983). The tube is shorter in most leeches but in the rhynchobdellid leech Theromyzon tessulatum reaches the remarkable length of 55 µm, and 66 µm in the branchiobdellid Cambarincola pamelae (Gelder and Ferraguti 2001). 3) the acrosome tube is thickened around the subacrosomal space by oblique ribs. 4) Presence of an endonuclear canal containing the axial rod (an endonuclear canal is present in the tubificid Rhizodrilus russus, see Ferraguti et al. 1994, but does not contain a rod). 5) Presence of a subplasmalemmal sheath of dense material in the basal portion of the tail, reminiscent of that of the fish leech Acanthobdella peledina. 6) Presence of a wide band of cytoplasm beneath the plasma membrane of the anterior region of the axoneme. 7) Replacement of the two glycogen granules usually associated with each axonemal doublet in clitellates with radial rows of glycogen granules which occupy a wide band of cytoplasm peripheral to the axonemal doublets. The large amount of glycogen in a broad cytoplasmic zone resembles the condition in tubificine parasperm though it is there γ-glycogen (see 8.4.3). As noted by Jamieson and Daddow (1992), it is not possible categorically to state which unique features of the sperm of Eudrilus are adaptations to the requirements of internal fertilization. Examination of sperm of externally (cocoon) fertilizing eudrilids is needed. However, all are apomorphies not seen in other oligochaete eusperm and coexist with highly apomorphic modifications of the reproductive system for internal fertilization in Eudrilus. The reproductive system rivals that of the similarly internally fertilizing leeches in its complexity. The apomorphies may therefore reasonably be considered to be adaptations for aspects of internal fertilization, presumably including specific requirements of altered sperm metabolism (increased glycogen storage), migration and storage of of sperm within the female system and peculiarities of sperm-egg interaction. The significance of elongation of the acrosome together with the extraordinary elongation of the axial rod, which varies between approximately 14 and 20 µm in length, are uncertain beyond its presumed relationship to internal fertilization. The length of the acrosome tube was shown in other oligochaetes to be highly correlated with that of the microvilli which constitute the zona pellucida of the egg (Jamieson et al. 1983). The egg of Eudrilus has yet to be studied. Microchaetidae. Spermatogenesis in Microchaetus pentheri follows the familiar pattern known for other oligochaetes (Hodgson and Jamieson 1992).

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As usual, spermatogenic stages develop around an anucleate cytophore from which they separate as mature spermatozoa. During spermiogenesis the nucleus elongates and becomes surmounted by a complex, elongate acrosome: the flagellar axoneme develops from the distal centriole. The centriole is positioned posterior to the midpiece. Microchaetus shows many plesiomorphic features in the structure of its acrosome, which are also seen in two other taxa of the Diplotesticulata, Haplotaxis (Haplotaxidae) and Sparganophilus (Spartganophilidae). The spermatozoon has a long (3.8 µm) acrosome, with a tube only partly containing the acrosome vesicle, which projects anteriorly in a spheroidal bleb (terminal bulb in Hodgson and Jamieson 1992). The vesicle is deeply introflected at its basis, delimiting a large subacrosomal space extending up to the bulb. In the subacrosomal space there is a 1 µm long rod entirely contained in the tube. The rod is surrounded basally by a nodelike sheath probably homologous to the secondary acrosome tube. A large basal chamber is delimited distally by the end of the acrosome vesicle and proximally by the limen terminating the nuclear end of the tube. A thin nuclear pad separates the acrosome from the rectilinear, 24 µm long, nucleus. Six parallel, radial mitochondria form the midpiece. A conventional megadrile flagellum follows, with tetragon fibers and glycogen granules (Hodgson and Jamieson 1992). Hormogastridae. The Sardinian species Hormogaster redii has a typical megadrile (metagynophoran) spermatozoon (Fig. 8.47D). The acrosome is 2.7 µm long, with the vesicle completely withdrawn into the tube. An axial rod is housed in the basal invagination of the vesicle. A short secondary tube surrounds the base of the rod. There is a limen at the base of the tube, and a pad with central boss separates the acrosome from the nucleus. The straight nucleus is followed by a midpiece with six (or seven) straight mitochondria (Ferraguti and Jamieson 1984). Lumbricidae. Comprehensive descriptions exist for spermatozoa of Lumbricus terrestris (Cameron and Fogal 1963; Anderson and Ellis 1968; Anderson et al. 1967, 1968; Anderson and Curgy 1969; Lanzavecchia and Lora Lamia Donin 1972; Shay 1972; Bergstrom and Henley 1973; Henley 1973) and of Allolobophora sp. (Troyer 1980; Troyer and Cameron 1980); see also an interpretation of their structure by Jamieson (1978a, 1981c) (Fig. 8.47E). The acrosome has been described and measured in several lumbricid species (Jamieson et al. 1983): Eisenia fetida; Lumbricus castaneus; Allolobophora longa; L. rubellus; Allolobophora chlorotica (Fig. 8.41A-E); Dendrobaena octaedra, Eiseniella tetraedra and Aporrectodea caliginosa. The observations point to a uniform sperm model for the family, with variations affecting the length of different portions. The acrosome has a length variable from 2.33 µm in L. rubellus to 12.7 µm in A. chlorotica (Jamieson et al. 1983), with the tube completely enclosing the vesicle, a 1 µm long rod in the basal invagination of the acrosome vesicle, and a short secondary tube around the base of the rod. The acrosome tube ends basally with a limen. There is a thin nuclear pad between the acrosome and the straight nucleus. The nucleus is 14.3 µm

!#

Reproductive Biology and Phylogeny of Annelida

long in Allolobophora sp. (Troyer 1980), and 9 µm in Lumbricus terrestris (Henley 1973). Six straight, 2 µm long, radial mitochondria follow, then a flagellum with tetragon fibers and glycogen granules. The tail plasma membrane shows two modifications: a flagellar necklace, i.e. a triple row of parallel particles at the base of the flagellum, and “long rows and aggregates of individual particles” longitudinally arranged (Bergstrom and Henley 1973) reminscent of the zipper lines of tubificids already mentioned. Megascolecidae. The ultrastructure of the spermatozoa of six species of megascolecid was described by Jamieson (1978a); data discussed also in Jamieson (1981c) and reviewed by Ferraguti (1999): Fletcherodrilus unicus (Fig. 8.47F), Cryptodrilus sp., Digaster longmani, Spenceriella sp., Amynthas (=Pheretima) sp., Amynthas corticis (=diffringens) and that of Amynthas rodericensis was illustrated in a discussion of spermathecal function by Jamieson (1992) (Fig. 8.47G). There are variations in the size of the different organelles in the various species examined, but the general scheme is uniform. The acrosome is 1.7 (Amynthas) to 2.6 µm (Digaster) long, thus being much shorter than that of lumbricids. The acrosome vesicle is completely withdrawn and deeply invaginated at its base. The acrosome rod lies within the invagination, only its basal portion being external to it and surrounded by a distinctive secondary tube very close to it and obliquely connected to the posterior rim of the acrosome vesicle. The node-like form of this secondary tube is diagnostic of the investigated Megasolecidae. Posterior to this there is a short basal chamber. The acrosome tube terminates with an obvious medianly directed shelf-like exension, the limen, surmounting a thin nuclear pad and the domed extremity of the nucleus. The nucleus is straight and about 10 µm long. A midpiece follows, with six parallel mitochondria, of variable length (0.5-1.4 µm). The tail has tetragon fibers and glycogen granules. The low level of ATP in oligochaete (Amynthas hawayanus) sperm is discussed by Teisaire and Del (1989).

8.4.3 Double Spermatogenesis in Oligochaetes The literature on oligochaete spermatogenesis contains many reports on the existence of “atypical” spermatozoa, reviewed by Fain-Maurel (1966) and Christensen (1980), uniformly interpreted as degenerating cells, without any genetic role in fertilization. In was not until 1980 that two functional types of spermatozoa were found in Limndodrilus hoffmeisterii, a species belonging to the tubificid subfamily Tubificinae (Block and Goodnight 1980) and their function interpreted in another member of the same subfamily, Tubifex tubifex, as joining to form the spermatozeugmata (defined above) (Braidotti et al. 1980). They were the large rods (up to 2 mm long) already described in tubificids in the 19th century (Claparède 1861; Lankester 1871). The spermatozeugmata. The first modern ultrastructural studies on spermatozeugmata were made on Tubifex tubifex, and showed that they contain two regions: an inner axial cylinder, and an outer cortex (Fig. 8.50I) (Braidotti and Ferraguti 1982; Ferraguti et al. 1988). In the axial cylinder the

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fertilizing eusperm, resembling the conventional oligochaete spermatozoa are orientated in parallel. In the cortex a large number of paraspermatozoa is arranged with the nuclei facing the interior and the tails spirally coiled around and tightly connected by cell junctions (Fig. 8.50H). Only the extremities of the sperm tails are free (Fig. 8.50I) (Ferraguti et al. 1988). This organization proved to be valid, with minor variations, for other tubificine species examined (see below, and Ferraguti 1999). The structure of Tubifex tubifex spermatozeugmata provides hints as to its functions: the spermatozeugmata hold together, by means of the junctional complex which connect parasperm tails, a large amount of euspermatozoa “ready for use”; the parasperm form the cortex of the spermatozeugmata and envelope the eusperm; the free ends of the parasperm tails are able to move, forming a metachronal wave; this probably transports fertilizing sperm towards the opening of the spermathecae at the moment of fertilization (Ferraguti et al. 1988). Other functions for parasperm cannot, however, be excluded: the parasperm may filter or process substances passing from the spermathecal lumen to the axial cylinder; the parasperm could also be a lowcost material to fill up the spermatheca of the partner, thus preventing further copulations (the ‘eunuch effect’ proposed for lepidopteran parasperm by Silberglied et al. 1984). When the study of spermatozeugmata was extended to other tubificine species it was found that, while the presence of a cortex produced by parasperm and of an axial cylinder produced by eusperm was a common feature in all tubificines, the shape of parasperm nuclei and the presence of cell junctions varied in the different species. In Tubifex tubifex and in Clitellio arenarius, the parasperm nuclei change their shapes in the spermatozeugmata: in the first species they degenerate visibly even showing myelin figures, whereas in the latter the nuclei are coiled on themselves, leaving only the acrosomes outside the skein. In Isochaetides arenarius (Ferraguti et al. 2002b), in members of Tubificoides (Ferraguti et al. 1989, and unpublished), and in Heterochaeta costata (unpublished) the parasperm nuclei in the spermatozeugmata maintain their rectilinear shapes. Cell junctions of two different types connect parasperm tails of Tubifex tubifex in the spermatozeugmata: septate junctions in the main tract, and scalariform junctions in a more distal portion (Ferraguti et al. 1988). In the second model of spermatozeugma studied, that of Clitellio arenarius (Ferraguti and Ruprecht 1992) the junctions were completely absent, whereas in species of Tubificoides only a limited number of septate junctions were present (Ferraguti et al. 1989, and unpublished). Heterochaeta costata, Psammoryctides barbatus (unpublished), and Isochaetides arenarius (Ferraguti et al. 2002b) showed a large number of septate junctions. As noted by Ferraguti et al. (2002b), it is possible that there is a connection between a freshwater habitat of a species and the production of large numbers of septate junctions. Species of the sister subfamily to Tubificinae, the fully marine Limnodriloidinae, also produce spermatozeugmata (Marotta et al. 2003). In the genus Limnodriloides the five species examined have ‘tubificine-type’

!#" Reproductive Biology and Phylogeny of Annelida

Fig. 8.50. A, B. Tubifex tubifex eusperm. A. Apical portion of the head. B. Basal portion of the nucleus, mitochondrial midpiece and proximal portion of the flagellum. Note the reduced size of the mitochondria and the ‘conventional’ aspect of the flagellar plasma membrane (× 20 000). C. Eusperm acrosome of T. tubifex fixed in the presence of tannic acid. The acrosome vesicle (paler) is almost completely withdrawn into the acrosome tube (× 55 000). D. Transition area between axial cylinder and cortex of a spermatozeugma of Fig. 8.50 contd

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spermatozeugmata, formed by eusperm surrounded by parasperm (those last are not connected by septate junctions, however), whereas members of Smithsonidrilus have spermatozeugmata of two types, each one formed by euor para-sperm only. Spermatozeugmata were also found in members of the other limnodriloidine genera examined, Doliodrilus and Thalassodrilides. Species of both genera produce eusperm only, but their spermatozeugmata differ significantly: a parsimony analysis indicates that these spermatozeugmata may even have arisen independently (Marotta et al. 2003). The same analysis suggests that, despite some morphological differences, the spermatozeugmata composed of both eusperm and parasperm may be homologous in the Tubificinae and Limnodriloides and that the simpler spermatozeugmata observed in Smithsonidrilus may be the result of an apomorphic secondary transformation of tubificine-like spermatozeugmata (Marotta et al. 2003). The paraspermatozoa. The two types of sperm (eusperm and parsperm) differ in all their parts (Fig. 8.50). Table 8.3 lists the differences discovered to date in Tubifex tubifex. The other models studied in some detail in both tubificines and limnodriloidines showed the same type of differences, with species-specific features. In general it may be said that in paraspermatozoa: 1. Acrosomes are reduced in size and contents, or even absent (as in all Limnodriloidinae) (Fig. 8.50E,C) 2. Nuclei are much shorter (up to one tenth those of the eusperm in tubificines) and slender (Fig. 8.50A-B,G). Chromatin shows uncondensed areas in all tubificines 3. Mitochondria are always fewer in number then in euspermatozoa (only T. tubifex has two mitochondria in both sperm types). In the tubificines, however, the volume of the parasperm midpiece is about double that of eusperm (Fig. 8.50B,G). 4. Plasma membrane surrounding the flagellum is largely separated from the axoneme in all tubificines and most limnodriloidines ( Fig. 8.50D,G,H). Fig. 8.50 contd

Isochaetides arenarius. In this area, cross-sections of different regions of both eusperm and parasperm are visible (× 20 000). E. Parasperm acrosome of T. tubifex. The acrosome tube appears ‘empty’ and the vesicle is completely external to the tube (× 55 000). F. Ciliated funnel of T. tubifex: both parasperm and eusperm are visible among the cilia (× 3 000). G. An entire parasperm head of T. tubifex. Note the plasma membrane largely separated from the nucleus and from the axoneme, and the large mitochondria (× 20 000). H. Cross section of the cortex of an I. arenarius spermatozeugma. Prominent septate junctions connect parasperm tails (× 45 000). I. Spermatozeugma of T. tubifex broken and seen under scanning electron microscope. Part of the axial cylinder with eusperm is visible, as well as part of the cortex formed by parasperm. In the main portion of the cortex, parasperm tails are tightly packed, only their extremities are free, and form a metachronal wave (× 2 500). Abbreviations: a, acrosome complex; c, cilia; ce, ciliated epithelium; f, flagellum; m, mitochondria; n, nucleus. Arrowheads point to some eusperm sections; arrows point to some parasperm sections.

!#$ Reproductive Biology and Phylogeny of Annelida Table 8.3 Differences between the characters of euspermatozoa and paraspermatozoa in Tubifex tubifex.

Sperm components Acrosome Acrosome tube Acrosome vesicle Acrosome rod Secondary tube Membrane particles on plasma membrane Nucleus Length Shape DNA content Chromatin Shape in spermatozeugmata Cytoplasm Mitochondria Number Volume Tail Basal body Basal cylinder Axoneme Glycogen Flagellar plasma membrane Particles arrangement on flagellar plasma membrane

Eusperm

Parasperm

containing the acrosome vesicle with dense contents present present regular array

thinner: acrosome vesicle external apparently empty absent absent absent

about 30 µm cylinder basally straight and apically twisted-column-shaped 1C fully condensed

about 3 µm comma-shaped, elliptical in crosssections approx. one eighth of that of the eusperm with uncondensed areas

maintained virtually absent

lost small amount present

two, hemispherical normal

two, hemispherical double

extremely short; no microtubules visible present with ‘tetragon fibers’ in the central apparatus 18 β-glycogen granules around the axoneme close to the axoneme

longer; doublets visible

interrupted zipper-lines

irregular or absent with a conventional central apparatus large amount of γ-glycogen between axoneme and plasma membrane largely separated from the axoneme forming septate and scalariform junctions in spermatozeugmata

We do not know the biological meaning of the differences observed between the parasperm of the tubificine and limnodriloidine species, but, equally, this is not known for eusperm. It may be speculated that the reduction or absence of the acrosome and the reduced size of the nuclei and, at least as far as Tubifex tubifex is concerned, the reduction of their DNA content shown by Ferraguti et al. (1987) is related to their not being built for fertilization. Genesis of the two sperm types in Tubificinae. In tubificids, as in all the oligochaetes studied (see 8.4.1), the cysts consist of a central cytoplasmic mass, the cytophore, to which the cells are connected through a narrow collar

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(zonula collaris) (Fig. 8.51A). The cysts pass into the seminal vesicles where, in the eusperm line they undergo a series of nuclear divisions without cytoplasmic divisions (Fig. 8.52A), and finally undergo meiosis. Cysts at different developmental stages are mixed in the seminal vesicles thus the problem of distinguishing between cysts belonging to the two spermatogenic lines arises. At the spermatid stage the task is easy: the cysts of paraspermatids are much more numerous than those of euspermatids (see a discussion in Braidotti and Ferraguti 1982). Secondly, the parasperm cysts consist of several hundreds (1250 ± 900; n=114 in Tubifex tubifex, Ferraguti et al. 2002a) of small cells (Fig. 8.52G-H), whereas the eusperm cysts contain a smaller number (128 in T. tubifex: Fig. 8.52B-E) of larger spermatids (Ferraguti et al. 1983). Furthermore, pycnotic nuclei are always present in the central cytophore of the paraspermatid cysts (Fig. 8.51) whereas nuclei are absent from the eusperm cytophore (reviewed in Ferraguti 1999). Many ultrastructural details of spermatids and spermiogenesis also differ: in particular the parasperm nuclei have, from the early spermiogenetic stages, an irregular shape; chromatin condensation is also irregular; the microtubular manchette is incomplete (pointing to some kind of relationship between microtubular manchette, chromatin condensation and nuclear morphogenesis, see Ferraguti and Ruprecht 1992). The tail shows, from early spermiogenesis, wide separation of the plasma membrane from the axonemes. The early spermatogenetic stages of both lines are easily recognized by counting the nuclei in each cyst (Fig. 8.52A). However, we could not distinguish between the two lines by their DNA content, whether by measuring single cells per cyst with a traditional method, Feulgen stain and densitometry (Ferraguti et al. 1987), or by measuring DNA content of the whole cysts under the confocal microscope (Boi et al. 2001). In other words, it was not possible to identify, in terms of DNA content, a line of spermatogenic cells containing a DNA amount being two or four times that of parasperm. Furthermore, DNA content of parasperm cysts (i.e. those having more than 128 cells), although extremely variable, is less or equal to that of the euspermatid cysts (i.e. those with 128 cells) (Boi et al. 2001; different figures were reported in Ferraguti 1999: for a discussion of the discrepancy see Boi et al. 2001). This rules out the possibility that parasperm are produced through an increased number of cell divisions. How is the commitment of the two sperm lines achieved? Two explanations were possible: either there is an early commitment of the two developmental lines, but this is not in terms of DNA content, or the spermatogenetic pathway is common in the two lines until the spermatocytes I stage (32 cell cysts), then some peculiar process occurs to produce paraspermatids. Laboratory cohort cultures of Tubifex tubifex with a constant check of spermatogenesis have revealed that the production of parasperm begins before that of eusperm (Boi and Ferraguti 2001). However, since “... euspermatid and paraspermatid cysts and their precursors (i.e. meiotic cysts and fragmenting cysts) are present in the seminal vesicles at the same time,

!#& Reproductive Biology and Phylogeny of Annelida

Fig. 8.51. A-E. The process of fragmentation during paraspermiogenesis in T. tubifex is here represented for a small portion of the cyst. For a description, see text and Boi et al. 2001 from which this figure is reproduced (with permission). F. Contrasted spermiogenesis in an oligochaete with only one sperm line, Bythonomus lemani (Lumbriculidae). Modified after Ferraguti, M. and Jamieson, B. G. M. 1987. Hydrobiologia 155: 123-134, Fig. 1.

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Fig. 8.52. Spermiogenesis and paraspermiogenesis in Tubifex tubifex. A-G. Optical micrographs (fluorescent microscope) of Feulgen-stained whole mounts of sperm cysts from the seminal vesicles. A. Nine premeiotic cysts (16 and 32 cells) (× 390). B. Euspermatid cyst at the beginning of spermiohistogenesis: the 128 nuclei are still roundish (× 390). C-E. progressive elongation of nuclei during euspermiogenesis. In E a (probably) 64 cells cyst is visible in the lower right corner (× 390). F. A (seemingly) 32 cells cyst undergoes the fragmentation process. At the upper left a four cells cyst (× 390). G. A paraspermatid cyst during the elongation process (× 280). H. A paraspermatid cyst at the same stage as the one in G as seen under phase contrast microscope shows an enormous number of flagella (× 280). Arrowheads point to two eusperm; arrow points to a parasperm.

the two sperm productions overlap. This leads us to exclude a sequential commitment due to hormone production during development, as is the case, for instance, of Lepidoptera (Friedländer 1997)” (Boi and Ferraguti 2001). Independently from the presence of an early commitment, a peculiar process of cell division in Tubifex tubifex has been identified for which the term ‘fragmentation’ has been coined (Boi et al. 2001). The following account will be mainly based on these findings. Fragmentation is a nuclear division which does not entail the formation of a spindle and a regular migration of equal portions of DNA into the daughter cells. The process of fragmentation is extremely complex and only partly understood (Fig. 8.51A-E). However, we were able to identify a population of cysts resembling the 32 cell cysts (euspermatocytes I), but characterized by the collapse of the collars connecting the single cells to the central cytophore (Fig. 8.52F) and by a process of de novo mass production of centrioles. The collapse of the collars is probably caused by a

!$ Reproductive Biology and Phylogeny of Annelida depolymerization of the ring-forming actin (Boi et al. 2001) which in turn lets the nuclei ‘slide’ into the cytophore (Figs. 8.51B-D, 8.52F). This phenomenon is accompanied by an impressive multiplication of the centrioles (Ferraguti et al. 2002a) due to the high number of spermatids produced by each paraspermatid cysts: each newly-formed centriole will, in fact, become the basal body of a parasperm. Curiously enough, multiplication of centrioles occurs through the model of de novo formation (deuterosomal mode), a model never before observed in an uniflagellated spermatozoon, but followed in the production of basal bodies in the multiciliated spermatozoa, like that of the termite Mastotermes darwiniensis (Baccetti and Dallai 1978) and the paraspermatozoa of certain gastropod molluscs (Healy and Jamieson 1981). It is, however, interesting to remember that in tubificine spermatozeugmata the cortex of parasperm will, in fact, behave as a multiciliated cell (Ferraguti et al. 1988). The next stage of paraspermiogenesis is the formation of irregular chromatin lumps in each nucleus, which will become the paraspermatid nuclei (Fig. 8.51D-E). The last step of paraspermiogenesis is the migration of the newly-formed centrioles to the periphery of the cytophore where they will grow a flagellum (Figs. 8.51E, 8.52G-H). In the same area one of the ‘lumps’ of chromatin, now detached from the nuclei, migrates, accompanied by two mitochondria. Finally, actin re-polymerizes, the collars are re-formed, and the paraspermatid cyst assumes the typical final aspect with a large cytophore at the center, and hundreds of small paraspermatids at the periphery each with its own flagellum (Fig. 8.51E). The mechanism of irregular nuclear fragmentation produces a considerable variability of DNA content in the parasperm and explains the presence of degenerating nuclei in the common central cytoplasmic mass of the cytophore. We may suppose that the information for the production and working of the ‘functional’ parts of the parasperm (flagella, cell junctions, mitochondria) is already present in the common cytoplasm before fragmentation. It is difficult, in our present state of knowledge, to speculate on a possible evolutionary origin of the dichotomous spermatogenesis in the tubificinelimnodriloidine assemblage. It seems pertinent to report that alterations of cell divisions have been described in the past during spermiogenesis in the oligochaete Pheretima heterochaeta (Cognetti de Martiis 1925) and that in Tubifex tubifex parthenogenesis occurs through a deep alteration of meiosis, the premeiotic doubling model (Christensen 1984; Baldo and Ferraguti 2005) as in oogenesis in some earthworms (see 8.5, below). Among annelids there is only one other example of dichotomous spermiogenesis: that described in 13 Protodrilus species belonging to 22 different populations of the polychaete Protodrilus by von Nordheim (1987, 1989). Spermiogenesis has been followed with particular detail in Protodrilus oculifer (von Nordheim 1987). There were no evident differences between the two developmental lines at the stage of spermatogonia and spermatocytes, whereas euspermatids and paraspermatids were clearly recognizable.

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It is tempting to establish a parallel between the tubificine and protodriline dichotomous spermatogenesis. In both cases, parasperm and eusperm jointly form sperm bundles which may be interpreted as a transport device for the fertilizing eusperm. In both cases, the parasperm of the different species resemble each other much more than do eusperm, thus suggesting some sort of ‘arrest’ of spermiogenesis. Finally, in both cases there seem to be no differences between the two developmental lines at spermatogonia and spermatocyte stages, whereas spermatids of the two types are easily distinguishable. However, this similarity may be ascribed to homoplasy as a close relationship of tubificines and protodrilines cannot readily be postulated.

8.5 MATING AND COITION (BARRIE G. M. JAMIESON) Mating refers to the events surrounding insemination and in oligochaetes involves coition (copulation). The present account greatly augments the most recent review of coition in earthworms, that of Benham (1950). Although hermaphrodite, oligochaetes are usually amphimictic, with copulation. Uniparental reproduction is, however, known and involves either self-fertilization, as in the Enchytraeus buchholzi and E. bulbosus (Dozsa 1995) or parthenogenesis, widespread in the megascolecid Amynthas (Gates 1972). Some populations, at least, of the cosmopolitan enchytraeids, E. buchholzi, and E. bulbosus have obligate uniparental reproduction. Presence of sperm in the spermathecae is attributed to entry from the cocoon as it passes over the spermathecal pores. In contrast, E. coronatus and E. irregularis reproduce only biparentally (Dozsa 1995). Parthenogenesis, as in earthworms results in diploid zygotes, without fertilization, owing to premeiotic doubling of the chromosome number in primary oocytes. Elegant investigations of this phenomenon were reported for the lumbricids Eiseniella tetraedra, Allolobophora rosea, A. caliginosa and Octolasion lacteum by Omodeo (1951, 1952, 1955). Early limitation of knowledge of oligochaete anatomy to the Lumbricidae resulted in acceptance of the mode of copulation in lumbricids as the norm for oligochaetes. In lumbricids the male pores (usually on segment 15) are apposed in copulation to the clitellum of the partner and the exuded spermatozoa move in external seminal grooves to its spermathecae located anterior to the cliellum (Fig. 8.53). Seminal grooves are also utilized in some ocnerodriles and megascolecids. They are seen in the ocnerodrilid Eukerria which has two pairs of prostate pores connected by seminal grooves to the pair of male pores, the acanthodrilin arrangement (Figs. 8.9, 8.54A). They are also seen in the balantin reduction seen in the acanthodriline megascolecid Balanteodrilus (Fig. 8.54C) as in Torresiella Dyne (1997). In the vast majority of oligochaetes, however, the male pores are apposed directly to the spermathecal pores of the partner and are often located on permanent or transient protrusions, forming distinct porophores or ‘penes’, which are inserted into the spermathecal pores. This form of copulation is illustrated

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Reproductive Biology and Phylogeny of Annelida

Fig. 8.53. Lumbricus terrestris (Lumbricidae). A. External features of worm turned slightly to one side to show genital pores, seminal groove and clitellum. B. Two worms in coition. The slime tube encloses the clitellum and apposed spermathecal pores. Relabelled after Jepson, M. 1951. Biological Drawings. Part II. John Murray, London, p. 31.

diagrammatically for the megascolecid (Megascolecinae) Spenceriella (Fig. 8.54, top) and Metaphire (Fig. 8.54D), which have the megascolecin arrangement with male and prostate pores united on segment 18, and is also exemplified by Microscolex dubius (Acanthodrilinae, Megascolecidae) (Fig. 8.54C) which has the microscolecin arrangement, with male and prostatic pores united on segment 17. Penetration may be aided by insertion of penial chaetae into the spermathecal orifices. This mode of insemination, with or without penial chaetae, is normal for the Megascolecidae. Coition has been described in detail for the megascolecid Eutyphoeus waltoni by Bahl (1927). This species is unusual for its family in copulating above ground. The most striking feature is the ‘male cup’ on segment 17 in the centre of which is a true penis from the tip of which protrudes a penial chaeta. Further forwards, on segment 7?, is a pair of spermathecal pores. In coition the two worms appose their ventral surfaces, with anterior ends pointing in opposite directions. Each male cup fits over the spermathecal papilla of the partner and the penis, and penial chaeta, is inserted into the duct of the spermatheca. Benham (1950) recognized four external organs or structures employed in coition:

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(1) Genital markings, including the tubercula pubertatis of lumbricids. These have been discussed in 8.2.4 above. (2) ‘Coupling chaetae’, here termed genital chaetae and including penial and spermathecal chaetae; have been discussed in 8.2.3 above. Lumbricus terrestris uses 40 needle-like copulatory chaetae situated ventrally on segments 10, 26, and the clitellar segments, 31 to 38, to inject a substance into the mating partner from the chaetal glands. Compared to the normal (crawling) chaetae, these chaetae are longer and grooved. It has been proposed that the chaetal glands may produce an allohormone that manipulates the reproductive physiology of the mating partner (Koene et al. 2002). Some penial chaetae of megascolecids are illustrated in Fig. 8.55. (3) Claspers, with which we may include the alae of Glyphidrilus and Lutodrilus. Claspers are best developed in the almid genus Alma for which their interspecific variation is illustrated in Fig. 8.11A,B. They are discussed in 8.2.4. Detail for Alma tazelaari is shown in Fig. 8.56. (4) A true penis. Male porophores which act as what may be considered true penes are discussed for Spenceriella and Eutyphoeus in this section, above. The large penis like structure, accompanied by a seminal groove, of Stuhlmannia variabilis is illustrated in Fig. 8.10; it is presumed that this is inserted into the spermathecal aperture. (5) Seminal grooves (added here) of various types. These include the semnal grooves of lumbricids, which run from the male pore, usually on segment 15 to the far posterior clitellum (Fig. 8.53) and that of the eudrilid Stuhlmannia variabilis, which runs from the male pore to the tip of the penis (Fig. 8.10). Other examples are the seminal grooves of Eukerria (Ocnerodrilinae) (Fig. 8.9) and of most Acanthodrilinae (Megascolecidae). Precopulatory behaviour. Mating may be preceded by precopulatory behaviour. Thus, mating of Lumbricus terrestris involves a pre-copulation behaviour sequence during which prospective partners visit each others burrows. Mate searching involves trail-following on the soil surface. This is followed by a series of, usually reciprocated, burrow visits. A burrow visit typically consisted of anterior segments insertion, for a period of 30 to 50 seconds, but also deeper burrow-penetrations, which sometimes lasted several minutes. Resident worms, when visited, either withdraw below ground completely or remain at the surface, with the first few anterior segments in view. Visiting worms normally retain their posterior segments in their own burrows. Partners often maintain close contact while moving back and forth between the burrow openings and the pre-copulation phase appears to be a specific courtship behaviour. Uninterrupted, the pre-copulation behaviour sequences lasted from 11 to 90 minutes. After a pre-copulation sequence, pairs adopt a static ‘s’-shaped copulation position of close ventral contact. Copulations lasts from 69 to 200 minutes (median 135 minutes). If other individuals touch the copulating pair, matings are shorter (Nuutinen and Butt 1997).

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Colour Figure

Fig. 8.54 contd

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8.55. Genital chaetae. A-I. Australian Megascolecinae. A-C. Heteroporodrilus mediterreus. D. Notoscolex camdenensis. E-F. Cryptodrilus polynephricus polynephricus. G-I. Digaster armifera. From Jamieson, B. G. M. 2000. The Native Earthworms of Australia (Megascolecidae Megascolecinae). Science Publishers, Inc.: Enfield, New Hamphshire, Fig. 0.12, after various papers of Jamieson. J-N. New Caledonian Acanthodrilus (Megascoelcidae, Acanthodrilinae). J, K. A. cavaticus. Spermathecal setae. L. A. cavaticus. Penial chaeta. M, N. A. chevalieri. Penial chaeta. Unpublished, from the study of Jamieson, B. G. M. and Bennett, J. D. 1979. Bulletin du Muséum National d’Histoire Naturelle Zoologie Ser 1: 353403. Fig. 8.54 contd

Fig. 8.54. Top. Coition in Spenceriella (Megascolecidae). After Jamieson, B.G.M. 1994. Chapter 39. Annelids, Arthropods and Molluscs. Pp. 855-878. In B. Knox, P. Ladiges and B. Evans (eds), Biology, McGraw-Hill Book Company, Sydney, Fig. 39.8. Bottom. The copulatory apparatus of a mating couple of Ocnerodrilidae and Megascolecidae. A. The acanthodrilin condition in Eukerria saltensis (Ocnerodrilidae) B. The microscolecin reduction in Microscolex (Acanthodrilinae). C. The balantin reduction in Balantodrilus (Acanthodrilinae). D. The megascolecin reduction in Metaphire saigonensis (Megascolecidae). Note that only in the megascolecin reduction do the vasa deferentia open into the prostate ducts, whereas in other cases the male pores are independent of the prostate pores, to which they are connected by narrow seminal grooves. A-D slightly modified after Omodeo, P. 2000. Italian Journal of Zoology 67: 179-201, Fig. 9.

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Fig. 8.56. Alma tazelaari (Almidae). Detail of claspers. After Jamieson, B. G. M. 1971. Glossoscolecidae. Pp. 41-72. In R. O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World. Oliver and Boyd, Edinburgh, Toronto, Fig. 15.8G (lapsus for H).

In the Spanish endemic earthworm Hormogaster elisae, spermatogenesis ceases during summer months but spermatozoa are retained in the seminal funnels and spermathecae through the year, allowing copulation at any time, whenever conditions allow. H. elisae has two pairs of spermathecae variable in both shape and size; sperm storage is mainly in the second pair (Garvin et al. 1999). Mated individuals of Lumbricus terrestris produced cocoons for up to 12 months after mating, while unmated individuals produced no cocoons. Hatchability of cocoons decreased to 11% in the sixth month after mating and zero thereafter. Median total production of viable cocoons is 5 per individual (range 0-21). There is no discernible relationship between cocoon production and length of copulation, individual longevity, or individual mass at mating. Both partners usually contribute to the production of viable cocoons, but within mating pairs there was a median difference of 4 cocoons. Median survival time after an experimental mating period was 9 and 11 months for mated and unmated earthworms, respectively (Butt and Nuutinen 1998).

8.6 FERTILIZATION, CLEAVAGE AND DEVELOPMENT 8.6.1 Sperm Entry, Polarity and Meiosis For a molecular perspective on development, see Chapter 5. In Tubifex, sperm entry is restricted to the vegetal hemisphere, especially near the vegetal pole. A fertilization cone is formed (Shimizu 1982, and

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%$literature therein). Two deformation movements then result in extrusion of the two successive polar bodies. The second deformation movement results in the formation of optically dense, yolk-free cytoplasm at the animal and vegetal poles, the so-called pole plasms mentioned below (see review by Jamieson 1981c; 1988a). The meiotic apparatus (MA) of meiosis I is located away from the egg surface at the time of oviposition. The position of the animal pole is subsequently marked by attachment of one pole of the MA to the Tubifex egg cortex as an optically bright spot (Shimizu 1981, 1982). A centriole is found in the inner aster of the MA. Later a bulge develops at this site and is extruded as the first polar body. The second meiotic apparatus if formed at the site of extrusion and is tethered to the surface by the microfilamentous cortical layer. Eggs remain at metaphase II (for 90 minutes) and then enter anaphase, followed by extrusion of the second polar body. The deformation movements which occur during polar body formation are dependent on actin-containing microfilaments (Shimizu 1982).

8.6.2 Evidence for Polarity in the Primary Oocyte A central problem in Tubifex development is the relationship between the pole plasm localization and cell determination. It is known that the pole plasms are segregated first to the D-cell, subsequently to the 2d- and 4d-cell (see 8.6.3, Fate maps, below), and finally to germ band cells (Shimizu 1982). During meiosis of the Tubifex primary oocyte, biosomatic elements (endoplasmic vesicles and mitochondria) and nutritive elements (lipid droplets and yolk granules) migrate within the cell by streaming movements (see review by Jamieson 1981c). Yolk granules may be concerned with organization as much as nutrition (Eckelbarger 1988). As a result of these segregational movements the components become arranged before the first cleavage division in a specific morphogenetic pattern of which the most conspicuous element is the concentration of mitochondria and endoplasmic vesicles at the animal and vegetal poles to form the two pole plasms. There is evidence that actin-like microfilaments in the cortex are responsible for accumulation of pole plasm (see review by Shimizu 1982).

8.6.3

Embryogenesis

The embryology of oligochaetes was reviewed and reinterpreted by Anderson (1971, 1973) and development in Tubifex was comprehensively studied and reviewed by Shimizu (1982). These studies form the basis of the present, albeit brief, review. Additional studies on the development of teloblasts in Tubifex (Goto et al. 1999a,b; Arai et al. 2000; Kitamura and Shimizu 2000a,b; Nakamoto et al. 2000; Shimizu et al. 2001) will also be summarized. Fate maps. The fate maps of the blastulae of oligochaetes and polychaetes are basically similar (Anderson 1971, 1973) (Fig. 8.57) (see also chapter 5). In both groups the 1st, 2nd and 3rd quartets of micromeres give ectoderm, with adult epidermis developing from 2d; 4d gives mesoderm; and macromeres 3 A, B and D give the midgut. In oligochaetes the process of

!$& Reproductive Biology and Phylogeny of Annelida cleavage, in terms of cell lineages, which segregate the areas from one another and deformation movements, varies greatly according to the degree of yolkiness of the eggs which is least in the crassiclitellates (Anderson 1971, 1973). Yolk content and cleavage. In the Enchytraeidae, Tubificidae sensu stricto and Lumbriculidae the egg is large, ranging in diameter from approximately 300 to 500 µm in Enchytraeus albidus and Tubifex to 1 mm in Rhynchelmis (references in Anderson 1973; Jamieson 1988a). The albumen which fills the cocoon and bathes the embryos appears to be of little significance as a source of food for development as this is provided by the internal yolk. The large (yolky) egg is deduced to be basic (plesiomorphic) in euclitellates. The size

Fig. 8.57. Embryology of an oligochaete (Tubifex) embryo. A. 8-cell stage, anterodorsal view. B. Presumptive areas, lateral view. After Anderson, D.T. In R.O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World. Oliver and Boyd, Edinburgh, Toronto, Figs. 2.1C, 2.2C.

and degree of yolkiness of the egg are imperfectly known for many oligochaete families but there is a reduction of yolk in the naids (now placed, as a subfamily, in the Tubificidae) and it is further reduced, presumably independently, in crassiclitellate families. In crassiclitellates, and to a lesser extent in naids, the eggs are smaller and the ambient albumen in the cocoon is exploited, in what is termed albumenotrophy, as a major source of nutrition during embryonic development. The mesolecithal egg in microdriles gastrulates by epiboly and the lateforming archenteron is empty whereas the oligolecithal egg of megadriles (crassiclitellates) gastrulates by emboly and forms a large archenteron which engulfs the albumen contained in the cocoon (see Anderson 1971; Omodeo 2000). Whether the egg be large or small, total, spiral cleavage, leading to a spherical blastula persists, as in polychaetes, but unlike the latter, the trochophore and, therefore, such regions as the presumptive prototroch are absent and there is no trace of larval organs or of metamorphosis.

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Gastrulation is prolonged and is accompanied by formation of numerous segments from the posterior growth zone, leading to direct development of the adult organization. Cleavage in the naidines Stylaria and Chaetogaster shows major modifications although they retain eggs of considerable diameter, 350 to 400 µm in Stylaria, and 400 to 500 µm in Chaetogaster. The blastula is attained through fewer cell divisions than in Tubifex and consequently has a large 3d cell instead of a large 4d cell (Anderson 1971, 1973). Earthworms (crassiclitellates) have small eggs (in lumbricids 70 µm in Dendrobaena subrubicunda and 100 to 120 µm in Eisenia fetida; references in Jamieson (1988a) exhibiting a more marked reduction in yolk than that seen in naidids. Their cleavage is correspondingly modified relative to that of Tubifex. The D quadrant is emphasized to an even greater degree than in naids but resemblance of the blastula to that of oligochaetes with yolky eggs is retained and the usual 4d cell is present posteroventrally (Anderson 1971, 1973). Cleavage is least modified in the large yolky eggs of tubificids and lumbriculids and we will take development in Tubifex as described by Shimizu (1982) as an example. Developmental stages in Tubifex are illustrated in Fig. 8.58. The Tubifex egg is fertilized at metaphase of the first meiosis, that is, as a primary oocyte. After extrusion of polar bodies, pole plasms comprising endoplasmic reticulum and mitochondria accumulate around the animal and vegetal poles. The developmental stages of polar body formation are characterized by a dynamic shape change called deformation movement. The first cleavage produces the smaller AB- and larger CD-cell. The second cleavage gives rise to four cells: A, B, C and D; the D-cell is larger than the other three cells. Thereafter, these four cells divide in a spiral cleavage pattern, producing micromeres and yolky macromeres. The pole plasms are segregated into 2dand 4d-cells resulting from divisions of the D-cell. The descendant cell (2d111) of 2d-cell and 4d-cell exclusively participate in teloblastogenesis. The stages of teloblastogenesis as seen by by SEM are illustrated in Fig. 8.59. Four ectoblasts and a mesoblast are located on either side of the embryo, and bud off small cells forming germ bands. Gastrulation movement consists of two events: 1) ventral shift and ensuing coalescence of germ bands, and 2) epibolic expansion of a micromere-derived epithelial sheet over the endodermal cells (Shimizu 1982). In naidines gastrulation is wholly eliminated; the presumptive midgut, M cells and the ectoteloblast cells attain their definitive positions directly as a result of cleavage and proceed directly into organogenetic activity. In earthworms, in contrast, gastrulation by invagination (emboly) occurs as a secondary development relative to gastrulation by overgrowth of the large presumptive midgut by definitive ectoderm which occurs in yolky clitellate embryos (Anderson 1971, 1973). Organogenesis begins during the gastrula stage. The ectodermal germ bands are responsible for the ventral nerve cord and circular muscle layer. The mesodermal organs are exclusively derived from the stem cells produced by

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Fig. 8.58 contd

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the mesoblasts. Gastrulation is followed by elongation of the embryo. It finally changes its shape to become vermiform. The embryonic period lasts for two weeks at 18°C, and is divided into 19 stages (Shimizu 1982). Significance of pole plasm. It appears that the acquisition of both pole plasms accounts for the totipotency of the D-cell quadrant and for formation of organs, as opposed merely to endoderm and ectodermal epithelium produced by isolated A, B and C cells. The pole plasm is devoid of yolk granules. Its major components are mitochondria and its different metabolism may be responsible for the asynchronous Tubifex cleavage which is led by the D-quadrant. Differences in cleavage patterns of D cells compared with other cells may be governed by the presence of pole plasm as this determines the position of the MA. The somatoblasts of the early embryo thus differ from each other in the different proportions of primary constituents of the egg cytoplasm (mitochondria and ER) and in the distinct patterns of spatial distribution of these (see review by Shimizu 1982). Origin of ectoderm and mesoderm (Teloblasts and their fate). The following account of the fate of teloblasts in Tubifex is based chiefly on that of Shimizu et al. (2001) and the findings of Arai et al. (2000); Goto et al. (1999a,b); Kitamura and Shimizu (2000a,b) and Nakamoto et al. (2000) which they review. The relevant events of development, as summarized by Shimizu et al. (2001) are illustrated in Fig 8.60A-H. As in other clitellates, embryonic development in Tubifex is characterized by the generation of five bilateral pairs of teloblasts (designated M, N, O, P and Q), which serve as embryonic stem cells to produce germ bands on either side of the embryo (Goto et al. 1999a; Shimizu et al. 2001 ) (Fig. 8.60C,D). Each teloblast divides repeatedly to produce primary blast cells which are arranged in a coherent longitudinal column or bandlet (Fig. 8.60D). Four of the five bandlets on each side of the embryo join together to form an ectodermal germ band, while the remaining bandlet becomes a mesodermal germ band underlying that of the ectoderm (Fig. 8.60H). A large part of the tissues comprising body segments has been assigned to the progenies of the teloblasts. Goto et al. (1999a) followed the fate of the progenies of each teloblast using horseradish peroxidase tracers. M teloblasts give rise to nearly all of the mesodermal tissues, which included circular and longitudinal muscles, coelomic walls, nephridia (in segments VII and VIII) and primordial Fig. 8.58 contd

Fig. 8.58. Diagrammatic illustration of developmental stages in Tubifex. Stages 1a-12c, animal pole view; stages 13-15, side and ventral view; stages 16-18, side view. N, O, P, and Q denote ectoblasts; NOPQ, OPQ, and OP denote precursor cells of ectoblasts. Mesoblasts (M) are located posteriorly behind ectoblasts (stage 12). Anteroposterior (Ar-Pr) and dorsoventral (DI-VI) axes are indicated for stages 13-15. Double arrowheads (stages 16-18) point to the anterior end of the embryo. E, endodermal cell; GB, ectodermal germ band; PB, Polar body; PP, pole plasm; S, chaeta; Sg, segment. After Shimizu, T. 1982. Development in the freshwater oligochaete Tubifex. Pp. 283-316. In F. W. Harrison, and R. R. Cowden (eds), Developmental Biology of Freshwater Invertebrates. Alan R. Liss, Inc., New York, Fig. 3.

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Fig. 8.59 contd

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germ cells (in segments X and XI). Although few in number, M teloblasts also contributed cells to the ventral ganglion. Similarly, each of the ectoteloblasts, N, O, P and Q, made a topographically characteristic contribution to the ectodermal tissues such as the nervous system (i.e. ganglionic cells and peripheral neurones) and epidermis, all of which exhibited a segmentally repeated distribution pattern (Goto et al. 1999a). In tubificids and enchytraeids, products of the proliferation of primordial germ cells (PGCs) spread forward through the mass of yolky midgut cells during gastrulation; in each genital segment they proliferate to form the testes and ovaries projecting into the coelom and covered by somatic peritoneum. However, it is reported that in Eisenia the primordial germ cells which, as in other earthworms, first become recognizable in the walls of the genital segments, cannot be traced to the products of division of the pair of cells cut off as the first products of the mesoteloblasts as these products regress late in gastrulation (see Anderson 1971, 1973). As stated by Shimizu (1982) it has yet to be firmly established that PGCs undergo migration. Ectodermal bands and segmentation. Segmentation of the ectoderm in Tubifex is a process of separation of 50-µm-wide blocks of cells from the initially continuous ectodermal germ band (GB), a cell sheet consisting of four bandlets of blast cells derived from ectoteloblasts (N, O, P and Q). The initially linear array of blast cells in each ectodermal bandlet gradually changes its shape in a lineage-specific manner. These morphogenetic changes result in the formation of distinct cell clumps, which are separated from the bandlet to serve as segmental elements (SEs). SEs in the N and Q lineages each consist of clones of two consecutive primary blast cells. In contrast, in the O and P lineages, individual blast cell clones are distributed across SE boundaries; each SE is, therefore, a mixture of a part of the preceding anterior clone and a part of the next posterior clone (Shimizu et al. 2001). The P and Q teloblasts uniquely give rise to additional ectodermal tissues, namely ventral and dorsal chaetal sacs, respectively. Furthermore, O teloblasts make a contribution to the nephridiopores in segments VII and VIII as well. Ectoteloblasts and mesoteloblasts are the main source of ectodermal and mesodermal segmental tissues, respectively, but all of the teloblasts produce more types of tissue than has previously been thought.

Fig. 8.59 contd

Fig. 8.59. A-D. CSFM Micrographs illustrating the sequence of teloblastogenesis. A. Stage 11. B. Stage 12a. C. Stage 12b. D. Stage 12c. Ectoblasts are designated N, O, P, and Q; NOPQ, and OP denote Precursor cells of ectoblasts. M, mesoblast. Mesoblasts in C and D are covered with epithelium and indicated by ‘M.’ ed, endoderm; inic, cells derived from micromeres. × 120. E,F. Higher magnification of C and D, showing details of initiation of ectodermal gerrn band formation. Bars indicate relations between ectoblasts and their offspring. × 210. After Shimizu, T. 1982. Development in the freshwater oligochaete Tubifex. Pp. 283-316. In F. W. Harrison, and R. R. Cowden (eds), Developmental Biology of Freshwater Invertebrates. Alan R. Liss, Inc., New York, Fig. 10.

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Fig. 8.60. Summary of Tubifex development. A and B. Posterior view with dorsal to the top. C and D. Dorsal view with anterior to the top. E-G. Side view with anterior to the left and dorsal to the top. A. A 22cell stage embryo. Cells 2d11, 4d and 4D all come to lie in the future midline. B. 4d divides bilaterally into left and right mesoteloblasts (MI and Mr); 2d111 derived from 2d11 divides into a bilateral pair of ectoteloblast precursors (NOPQl and NOPQr), and 4D divides into a pair of endodermal precursor cells ED. C. An embryo at about 30 h after the bilateral division of 4d. Only teloblasts are depicted. NOPQ on each side of the embryo has produced ectoteloblasts N, O, P and Q. D. A two-day-old embryo following the bilateral division of 4d. Only teloblasts and associated structures are depicted. At this stage, a short ectoderrnal germband (EGB) extending from the ectoteloblasts N, O, P and Q is seen on either side of the embryo. A mesodermal germ band (MGB) extending from the M teloblast is located under the ectodermal germ band. E-G. Morphogenesis of the ectodermal germ band. Embryos are shown in E 2.5, F 4, and G 6 days after the 4d cell division. E. The germ band (EGB) is associated, at its anterior end, with an anteriorly located cluster of micromeres (called a micromere cap; MC), and it is initially located at the dorsal side of the embryo. F. The germ bands (EGB) on both sides of the embryo elongate and gradually curve round toward the ventral midline and finally coalesce with each other along the ventral midline. G. The coalescence is soon followed by dorsalward expansion of the edge of the germ band, Pr, prostomium. H. Longitudinal section showing the relative positions of the endoderm (end) and bandlets extending from teloblasts M and O. Anterior is to the left and posterior is to the right. The bandlet (germ band) derived from the M teloblast is overlain by the O-bandlet and is underlain by the endoderm. Asterisks indicate the presence of a single primary blast cell in each block of the bandlet. The remaining blocks individually represent a cell cluster, which is derived from a single primary blast cell. After Shimizu, T. et al. 2001. Hydrobiologia 463(123): 123-131, Fig. 1.

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Without the underlying mesoderm, separated SEs fail to space themselves at regular intervals along the anteroposterior axis (Nakamoto et al. 2000; Shimizu et al. 2001). Nakamoto et al. (2000) suggest that ectodermal segmentation in Tubifex consists of two stages: first, autonomous morphogenesis of each bandlet leading to generation of segmental elements and, secondly, the ensuing mesoderm-dependent alignment of separated segmental elements (Nakamoto et al. 2000; Shimizu et al. 2001). Some of the epidermis of the Tubifex embryo is reported to derive from the temporary yolk sac (Anderson 1971, 1973). Mesodermal segmentation. Using lineage tracers, in Tubifex Goto et al. (1999b) showed that segmental organization arises sequentially in the anterior-to-posterior direction along the longitudinal axis of the mesodermal germ band, a coherent column of primary blast cells that are produced from the mesodermal teloblast. Shortly after its origin, each primary blast cell undergoes a spatiotemporally stereotyped sequence of cell divisions to generate three classes of cells (in terms of cell size), which together give rise to a distinct cell cluster, a mesodermal compartment (Fig. 8.61). Each cluster is composed of descendants of a single primary blast cell; there is no intermingling of cells between adjacent clusters. Relatively small-sized cells in each cluster become localized at its periphery to form coelomic walls including an intersegmental septum thus establishing individuality of segments. Ablation experiments show that these features of mesodermal segmentation are not affected by the absence of the overlying ectodermal germ band, that each primary blast cell serves as a founder cell of each mesodermal

Fig. 8.61. Schematic summary of pattern and sequence of divisions in mesodermal blast cells. Inequality and direction of divisions are reflected by position and orientation of mitotic spindles in dividing cells. The mesodermal germ band (GB) extending from the mesoteloblast (M) cell is illustrated in the lower part of the figure; each block in the GB represents a cell cluster. Arrows indicate the approximate position, along the GB, where each division occurs. A—anterior; D—dorsal; P—posterior; V—ventral. After Shimizu, T. et al. 2001. Hydrobiologia 463(123): 123-131, Fig. 4, adapted from Goto A. et al. 1999. International Journal of Developmental Biology. July 43(4): 317-327, Fig. 4.

!%$ Reproductive Biology and Phylogeny of Annelida segment and that the boundary between segments is determined autonomously. In contrast with development of the ectoderm (see below), the metameric body plan of Tubifex thus arises from an initially simple organization (i.e., a linear series) of a segmental founder cell for each segment (Goto et al. 1999b; Shimizu et al. 2001). Using alkaline phosphatase activity as a biochemical marker for segments VII and VIII it appears that segmental identities in primary M-blast cells are determined according to the genealogical position in the M lineage and that the M teloblast possesses a developmental program through which the sequence of blast cell identities is determined (Shimizu et al. 2001).

8.6.4

Organogenesis

Somite formation. On each side of the embryo a somite derives from a single mesodermal stem cell produced by the mesoblast (Shimizu 1982). The mode of segmentation of the mesoderm, and the overlying ectoderm has been outlined above. Coelomic walls. The coelomic walls of the somite are differentiated into lateral somatopleure (somatic mesoderm), median splanchnopleure (splanchnic mesoderm) and the transverse epithelium covering the septa. The longitudinal muscle of the body wall differentiates from the somatic mesoderm. The splanchnic mesoderm is differentiated into the gut musculature and the overlying splanchnic peritoneum which on the midgut (and elsewhere) usually forms the cholagogen tissue (Shimizu et al. 2001). Prostomium. Shimizu et al. (2001) state, for Tubifex, that the prostomium and the cerebral ganglia (brain) originate from cells that are not the progeny of teloblasts. The prostomium, at least, derives from a ‘micromere cap’ (Fig. 8.60E-G). This contrasts with the view (Anderson 1971, 1973) that the prostomium originates from the anterior ends of the ectoblast bands and their underlying mesodermal bands and that the cerebral ganglia develop from the ventral neuroblast components at the extreme anterior end of each ectoblast band. Peristomium. The oligochaete peristomium appears to be a single segment formed by fusion of the first, bilateral pair of segmental somites, and overlying segmental ectoderm, immediately behind the prostomial rudiment. The neuroblast components of the segmental ectoderm giving rise within it to a single pair of ventral ganglia (Anderson 1971, 1973). Stomodeum. In yolky (lecithotrophic) euclitellate embryos, as part of the development of the gut, the stomodeum eventually grows back through the first segments of the segmenting embryo, develops a lumen and differentiates as the lining epithelium of the pharynx. Continuity between the buccal and pharyngeal lumen is established. Albumenotrophic embryos, in contrast, develop a precociously functional embryonic pharynx lined by cilia. In earthworms this is developed during gastrulation while that of naidines arises as an independent albumenotrophic invagination. It is later transformed into or (earthworms) replaced by the definitive pharynx. In Tubifex, the large mass of yolky cells, or in Rhynchelmis a syncytium, filling

Non-leech Clitellata

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the interior of the embryo develops more or less directly into a midgut epithelium by developing a central split. In naidines a provisional midgut sac becomes connected with the provisional pharynx and presumably acts temporarily in feeding on the ambient albumen. The walls of this sac later merge and become syncytial before resorption of the central part of the syncytium and differentiation of the definitive midgut. In earthworms the midgut sac formed as a result of invaginate gastrulation is already connected with the provisional pharynx and takes over the albumenotrophic role played in cleavage by albumenotrophic cells. The proctodeum forms, like the pygidial cells which surround it, from cells of the temporary yolk sac ectoderm. The formation of the proctodeum in naidines, in which the provisional ectoderm does not contribute to the definitive ectoderm, requires elucidation (Anderson 1973). Blood vessels. The ventral blood vessel (VBV) develops in Tubifex by separation of the apposed walls of the ventral mesentery and the segmental commissural vessels by separation of the apposed walls of the intersegmental septa; thus the blood vascular system occupies the site of the former blastocoel. In Criodrilus and Eisenia, however, the VBV is first apparent between the ventral mesentery and the floor of the midgut. In Tubifex the dorsal vessel develops precociously, as a paired vessel between the upper edges of the somites and the lateral surfaces of the yolky midgut. Later, as the edges of the somites come together in the dorsal midline, the two half-vessels combine into a single, dorsal longitudinal vessel in the resulting mesentery (Anderson 1971, 1973). In contrast, Shimizu (1982) states that all blood vessels develop between the endoderm and splanchnopleure of the somites. Nephridia. That nephridia are ectodermal derivatives, as is often stated, has been controversial. Origin from a single nephridioblast is generally accepted (Anderson 1971, 1973). Anderson noted that the septal location of the nephridioblast had been taken as evidence that the cell has a mesodermal origin but he considered the possibility of an earlier derivation from ectoderm. Goto et al. (1999a) demonstrated that embryonic nephridia of Tubifex are mesodermal, being descended from M teloblasts. These provisional protonephridia develop anteriorly in Tubifex, Rhynchelmis, and earthworms but were considered to be absent in naidines (Anderson 1973). However, Bunke (2003) has convincingly demonstrated origin of metanephridia of the naidine Dero digitata from three nephroblast cells in the frontal epithelium of a septum, suggesting its mesodermal origin. One cell produces the rudiment of the canal; another the ciliated mantle cell, and the third produces a flame of cilia that beats into the canal lumen, the so-called flame cell. In Tubifex hattai each nephridium-like structure was traced back, by histochemical staining, to a single cell (detected by alkaline phosphatase staining) that emerges in the mesodermal territory in the ventrolateral region of each of segments 7 and 8 at the late gastrula stage (Kitamura and Shimizu 2000b). Gonoducts. Where development of the gonoducts has been investigated in clitellate embryos, they have been identified as coelomoducts. In

!%& Reproductive Biology and Phylogeny of Annelida oligochaetes each gonoduct arises as a thickening of the coelomic epithelium opposite a gonad. The thickening develops as a funnel while the base of the thickening grows out as a duct. A small ectodermal invagination establishes the opening to the exterior. A detailed study of the development of the gonoducts and prostates in Tubificidae by Gustavsson and Erséus (1997, 1999) has been discussed in 8.2.11.2.

8.7

ACKNOWLEDGMENTS

The work of BGMJ was made possible by previous grants from the Australian Research Council, Australian Biological Resources Study, and University of Queensland Research Grants. Provision of facilities by the School of Integrative Biology, University of Queensland is also gratefully acknowledged. Mrs. Lina Daddow is especially to be thanked for aid in ultrastructural studies. Andrew Hugall is thanked for invaluable collaboration in molecular analyses and for kindly making his unpublished MCMC analyses available. Christer Erséus and Emilia Rota kindly provided some literature. Two PhD students of MF, Silvia Boi and Roberto Marotta, have done most of the work on the double sperm line respectively of Tubifex, and the limnodriloidines; Giulia Mazzoleni, and Samuele Cerea have done most of the cutting; Christer Erséus supplied the marine species; Patrick Martin contributed the Lake Baikal species; Umberto Fascio has helped with the confocal microscope. The research was supported by a grant from MIUR (Rome).

8.8 LITERATURE CITED Adiyodi, K. G. 1988. Annelida. Pp. 189-250. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates, John Wiley and Sons, New York. Anderson, D. T. 1971. Embryology. Pp. 73-103. In R. O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh. Anderson, D. T. 1973. Embryology and Phylogeny in Annelids and Arthropods. Pergamon Press, New York. Anderson, W. A. and Curgy, J. J. 1969. RNA synthesis in nuclei and mitochondria during spermiogenesis of Lumbricus terrestris. Journal of Submicroscopic Cytology 1: 25-34. Anderson, W. A. and Ellis, R. A. 1968. Acrosome morphogenesis in Lumbricus terrestris. Zeitschrift fur Zellforschung und Mikroskopische Anatomie 85: 398407. Anderson, W. A., Weissman, A. and Ellis, R. A. 1967. Cytodifferentiation during spermiogenesis in Lumbricus terrestris. Journal of Cell Biology 32: 11-26. Anderson, W. A., Weissman, A. and Ellis, R. A. 1968. Intranuclear crystalline inclusions in spermatozoa of Lumbricus terrestris. Protoplasma 66: 21-26. Arai, A., Nakamoto, A. and Shimizu, T. 2000. Specification of ectodermal teloblast lineages in embryos of the oligochaete annelid Tubifex: Involvement of novel cell-cell interactions. Development Cambridge 128(7): 1211-1219. Baccetti, B. and Dallai, R. 1978. The spermatozoon in the termite Mastotermes darwiniensis. Journal of Cell Biology 76: 569-576.

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Baldo, L. and Ferraguti, M. 2005. Mixed reproductive strategies in Tubifex tubifex (Oligochaeta, Tubificidae)?. Journal of Experimental Zoology, 303A, 168-177. Bahl, K. N. 1927. On the reproductive processes of earthworms: Pt. I. The process of copulation and exchange of sperms in Eutyphoeus waltoni Mich. Quarterly Journal of Microscopical Science 71: 481-502. Beauchamp, K. A., Kathman, R. D., McDowell, T. S., Hedrick, R. P., Braun, E. L. and Ligon, J. D. 2001. Molecular phylogeny of tubificid oligochaetes with special emphasis on Tubifex tubifex (Tubificidae). Molecular Phylogenetics and Evolution 19(2): 216-224. Beddard, F. E. 1889. On the structure of three new species of earthworms, with remarks on certain points in the morphology of the Oligochaeta. Quarterly Journal of Microscopical Science 29: 103-131. Beddard, F. E. 1901. On some earthworms from British East Africa; and on spermatophores of Polytoreutus and Stuhlmannia. Proceedings of the Zoological Society of London 1: 336-365. Benham, W. B. 1887. Studies on earthworms. III. Criodrilus lacuum, Hoffmeister. Quarterly Journal of Microscopical Science 27: 561-#. Benham, W. B. 1950. A review of certain external structures employed in coition of earthworms. Proceedings of the Zoological Society of London 119(4): 905-916. Bergstrom, B. H. and Henley, C. 1973. Flagellar necklaces: freeze-etch observations. Journal of Ultrastructure Research 42: 551-553. Block, E. M. and Goodnight, C. J. 1980. Spermatogenesis in Limnodrilus hoffmeisteri (Annelida, Tubificidae): a morphological study of the development of two sperm types. Transactions of the American Microscopical Society 99: 368-384. Boi, S., and Ferraguti, M. 2001. Temporal pattern of the double sperm line production in Tubifex tubifex (Annelida, Oligochaeta). Hydrobiologia 463: 103106. Boi, S., Fascio, U. and Ferraguti, M. 2001. Nuclear fragmentation characterises paraspermiogenesis in Tubifex tubifex (Annelida, Oligochaeta). Molecular Reproduction and Development 59(4): 442-450. Bondi, M., Ferraguti, M. and Scari, G. 1993. Sperm ultrastructure in Nematogenia panamensis (Annelida, Oligochaeta, Ocnerodrilidae): A phylogenetic approach. Invertebrate Reproduction and Development 23(1): 51-57. Bouché, M. B. 1972. Lombriciens de France: Écologie et Systématique, Institut National de la Recherche Agronomique, Vol. 72. 671 pp. Braidotti, P. and Ferraguti, M. 1982. Two sperm types in the spermatozeugmata of Tubifex tubifex (Annelida, Oligochaeta). Journal of Morphology 171: 123-136. Braidotti, P., Ferraguti, M. and Fleming, T. P. 1980. Cell junctions between spermatozoa flagella within the spermatozeugmata of Tubifex tubifex (Annelida: Oligochaeta). Journal of Ultrastructure Research 74: 299-309. Brinkhurst, R. O. 1964. A taxonomic revision of the Alluroididae (Oligochaeta). Proceedings of the Zoological Society of London 142: 527-536. Brinkhurst, R. O. 1982. Evolution in the Annelida. Canadian Journal of Zoology 60(5): 1043-1059. Brinkhurst, R. O. 1988. A taxonomic analysis of the Haplotaxidae (Annelida, Oligochaeta). Canadian Journal of Zoology 66(10): 2243-2252. Brinkhurst, R. O. 1989. A phylogenetic analysis of the Lumbriculidae (Annelida, Oligochaeta). Canadian Journal of Zoology 67(11): 2731-2739. Brinkhurst, R. O. and Gelder, S. R. 1989. Did the lumbriculids provide the ancestors of the branchiobdellidans, acanthobdellidans and leeches? Hydrobiologia 180: 7-15.

!& Reproductive Biology and Phylogeny of Annelida Brinkhurst, R. O. and Jamieson, B. G. M. 1971. Aquatic Oligochaeta of the World. Oliver and Boyd, Edinburgh. 462 pp. Brinkhurst, R. O. and Nemec, F. L. 1986. A comparison of phenetic and phylogenetic methods applied to the systematics of the Oligochaeta. Hydrobiologia 155: 65-74. Bunke, D. 1985. Ultrastructure of the spermatozoon and spermiogenesis in the interstitial annelid Potamodrilus fluviatilis. Journal of Morphology 185(2): 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 Ultrastructure and Molecular Structure Research 95(1-3): 113-130. Bunke, D. 2003. Early development of metanephridia in the caudal budding zone of a clitellate annelid, Dero digitata (Naidida): an electron microscopical study. Acta Zoologica 84(2): 87-97. Butt, K. R. and Nuutinen, V. 1998. Reproduction of the earthworm Lumbricus terrestris Linné after the first mating. Canadian Journal of Zoology 76(1): 104109. Cameron, M. L. and Fogal, W. H. 1963. The development and structure of the acrosome in the sperm of Lumbricus terrestris. Canadian Journal of Zoology 41: 753-761. Chekanovskaya, O. V. 1966. English translation 1981. Aquatic Oligochaeta of the USSR, United States Department of the Interior and the National Science Foundation, Washington, D. C., Amerind Publishing Co. Pvt. Ltd, New Delhi. 513. pp. Christensen, B. 1980. Annelida. Pp. 1-81. In B. John (ed), Animal Cytogenetics, Borntraeger, Berlin, Stuttgart. Christensen B. 1984. Asexual propagation and reproductive strategies in aquatic Oligochaeta. Hydrobiologia 115: 91-95. Christensen, B. and Theisen, B. F. 1998. Phylogenetic status of the family Naididae (Oligochaeta, Annelida) as inferred from DNA analyses. Journal of Zoological Systematics and Evolutionary Research 36(4): 169-172. Claparède, E. 1861. Études anatomiques sur les annelides, turbellariés, opalines et grégarines observés dans les Hébrides. Mémoires de la Sociéte de Physique et d’Histoire Naturelle de Genève 16: 71-164. Coates, K. A. 1986. Redescription of the oligochaete genus Propappus and diagnosis of the new family Propappidae (Annelida: Oligochaeta). Proceedings of the Biological Society of Washington 99(3): 417-428. Cognetti de Martiis, L. 1910. Contibuto all conoscenza della fecondazione negli Oligocheti. Atti Academia della Scienze, Torino 45: 493-506. Cognetti de Martiis, L. 1925. La prespermatogenesi negli oligocheti (Terricoli) e la formazione del citoforo. Bollettino dei Musei di Zoologia ed Anatomia comparata della R. Università di Torino 40: 1-38. Cook, D. G. 1968. The genera of the family Lumbriculidae and the genus Dorydrilus (Annelida, Oligochaeta). Journal of Zoology (London) 156: 273-289. Cook, D. G. 1971a. Family Dorydrilidae. Pp. 647-653. In R. O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto. Cook, D. G. 1971b. Family Opistocystidae. Pp. 640-653. In R. O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto. Cuadrado, S. and Martinéz-Ansemil, E. 2001. External structures used during attachment and sperm transfer in tubificids (Annelida, Oligochaeta). Hydrobiologia 463: 107-113.

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de Majo, A. M. A. 2002a. Gametogenesis masculina en Oligochaeta, Megascolecidae. I: Espermatogenesis normal de Amynthas hawayanus (Rosa, 1891), Amynthas morrisi (Beddard, 1892) y Metaphire californica (Kinberg, 1867) de la provincia de Tucuman, Argentina. Acta Zoologica Lilloana 2002 46: 5-21. de Majo A. M. A. 2002b. Gametogenesis masculina en Oligochaeta, Megascolecidae. II: Espermatogenesis en ejemplares no clitelados de Amynthas hawayanus (Rosa, 1891), Amynthas morrisi (Beddard, 1892) y Metaphire californica (Kinberg, 1867) de la provincia de Tucuman, Argentina. Acta Zoologica Lilloana 2002 46: 67-70. Dixon, G. C. 1915. Tubifex. Liverpool Marine Biological Committee Memoirs on Typical British Marine Plants and Animals 33: 1-93. Dozsa, F. K. 1995. Self-fertilization: An adaptive strategy in widespread enchytraeids. European Journal of Soil Biology 31(4): 207-215. 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. Dyne, G. R. 1997. Two new genera of Acanthodrilinae (Megascolecidae, Oligochaeta) from Cape York Peninsula and the Torres Strait. Memoirs of the Queensland Museum 42(1): 139-157. Eckelbarger, K. J. 1988. Oogenesis and female gametes. Pp. 281-307. In W. Westheide and C. O. Hermans (eds), The Ultrastructure of Polychaeta, Microfauna Marina, P. Ax (ed.) 4, Gustav Fischer, Stuttgart, New York. Erséus, C. 1997. A record of Randiella from New Caledonia, the first known occurrence of the marine interstitial family Randiellidae (Annelida; Oligochaeta) in the South Pacific Ocean. Journal of Natural History. Dec. 31(12): 1745-1750. Erséus, C. 1979. Bermudrilus peniatus n.g., n.sp. (Oligochaeta, Tubificidae) and two new species of Adelodrilus from the North-west Atlantic. Transactions of the American Microscopical Society 98: 418-427. Erséus, C. 1981. Taxonomic studies of Phallodrilinae (Oligochaeta, Tubificidae) from the Great barrier Reef and Comoro Island with descriptions of ten new species and one new genus. Zoologica Scripta 10: 15-31. Erséus, C. 1990. Cladistic analysis of the subfamilies within the Tubificidae (Oligochaeta). Zoologica Scripta 19: 57-63. Erséus, C. 1997. A record of Randiella from New Caledonia, the first known occurrence of the marine interstitial family Randiellidae (Annelida; Oligochaeta) in the South Pacific Ocean. Journal of Natural History. 31(12): 1745-1750. Erséus, C. 1999. Parvidrilus strayeri, a new genus and species, an enigmatic interstitial clitellate from underground waters in Alabama. Proceedings of the Biological Society of Washington 112(2): 327-337. Erséus, C. 2003. Phylogeny of oligochaetous Clitellata. Hydrobiologia 535(1): 357372. Erséus, C. and Baker, H. R. 1982. New species of the gutless marine genus Inanidrilus (Oligochaeta, Tubificidae) from the Gulf of Mexico and Barbados. Canadian Journal of Zoology 60: 3063-3067. Erséus, C. and Ferraguti, M. 1995. The use of spermatozoal ultrastructure in phylogenetic studies of Tubificidae (Oligochaeta). Mémoires du Muséum National d’Histoire Naturelle 166(0): 189-201. Erséus, C. and Gustavsson, L. 2002. A proposal to regard the former family Naididae as a subfamily within the Tubificidae (Annelida, Clitellata). Hydrobiologia 485: 253-256. Erséus, C. and Källersjö, M. 2003. 18S rDNA phylogeny of basal groups of Clitellata (Annelida). Zoologica Scripta: 33(2): 187-196.

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Erséus, C., Kallersjo, M., Ekman, M. and Hovmoler, R. 2002. 18S rDNA phylogeny of the Tubificidae (Clitellata) and its constituent taxa: dismissal of the Naididae. Molecular Phylogenetic and Evolution 22(3): 414-422. Erséus, C. and Strehlow, D. R. 1986. Four new interstitial species of Oligochaeta representing a new family. Zoologica Scripta 15: 53-60. Erséus, C., Prestegaard, T. and Kallersjo, M. 2000. Phylogenetic Analysis of Tubificidae (Annelida, Clitellata) Based on 18S rDNA Sequences. Molecular Phylogenetics and Evolution 15(3): 381-389. Fain-Maurel, M. A. 1966. Acquisitions récentes sur les spermatogenèses atypiques. Année Biologique 5: 513-564. Fawcett, D. W., Anderson, W. A. and Phillips, D. M. 1971. Morphogenetic factors influencing the shape of the sperm head. Developmental Biology 26: 220-251. Feldkamp, J. 1924. Untersuchungen über die Geschlechtsmerkmale und die Begattung der Regenwürmer. Zoologisches Jahrbücher, Anatomie 46: 609632. Fernandez, I. and Benito, J. 1987. Fine structure of the clitellum of Lumbricus friendi Cognetti 1904 (Annelida, Oligochaeta). Boletin De La Real Sociedad Espanola De Historia Natural Seccion Biologica 83(1-4): 79-92. Fernandez, J., Goater, T. M. and Esch, G. W. 1991. Population dynamics of Chaetogaster limnaei limnaei (Oligochaeta) as affected by a trematode parasite in Helisoma anceps (Gastropoda). American Midland Naturalist 125(2): 195-205. Ferraguti, M. 1983. Annelida-Clitellata. Pp. 343-376. In K. G. Adiyodi and R. G. Adiyodi (eds), Reproductive Biology of Invertebrates, John Wiley and Sons, New York. Ferraguti, M. 1984a. Slanted centriole and transient anchoring apparatus during spermiogenesis of an oligochaete (Annelida). Biology of the Cell 52(2): 175-180. Ferraguti, M. 1984b. The comparative ultrastructure of sperm flagella central sheath in Clitellata reveals a new Autapomorphy of the group. Zoologica Scripta 13: 201-207. Ferraguti, M. 1999. Euclitellata. Pp. 125-182. In B. G. M. Jamieson (ed), Reproductive Biology of Invertebrates. Series editors K. G. and R. G. Adiyodi, John Wiley, Chichester. Ferraguti, M., Balsamo, M., and Fregni, E. 1995. The spermatozoa of three species of Xenotrichulidae (Gastrotricha: Chaetonotida). The two “dunne Nebengeisseln” of spermatozoa in Heteroxenotrichula squamosa are peculiar para-acrosomal bodies. Zoomorphology 115(2): 151-160. Ferraguti, M., Bernardini, G., Melone, G. and Dallai, E. 1988. Structure and function of the metachronal wave in Tubifex tubifex spermatozeugmata (Annelida, Oligochaeta). Journal of Ultrastructure and Molecular Structure Research 99(1): 79-95. Ferraguti, M., Bondi, M. and Lanzavecchia, S. 1991. Ultrastructural analysis of sperm flagella in two clitellates (Annelida), plasma membrane and periaxonemal area. Tissue and Cell 23(4): 537-546. Ferraguti, M., Braidotti, P. and Trigari, A. 1983. Two different lines in Tubifex tubifex spermiogenesis. Pp. 446-449. In J. André (ed), The Sperm Cell, M. Nijhoff, The Hague. Ferraguti, M. and Erséus, C. 1999. Sperm types and their use for a phylogenetic analysis of aquatic clitellates. Hydrobiologia. May 402(0): 225-237. Ferraguti, M., Erséus, C. and Pinder, A. 1996. The spermatozoon of Capilloventer australis and the systemic position of the Capilloventridae (Annelida: Oligochaeta). Australian Journal of Zoology 44(5): 469-478.

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Jaana, H. 1982. The ultrastructure of the epithelial lining of the male genital tract and its role in spermatozeugma formation in Tubifex hattai Nomura (Annelida, Oligochaeta). Zoologisches Anzeiger 209: 159-176. Jamieson, B. G. M. 1957. Some species of Pygmaeodrilus (Oligochaeta) from East Africa. Annals and Magazine of Natural History, Series 12(10): 449-470. Jamieson, B. G. M. 1958. Taxonomic Studies on the Ocnerodrilinae and of the Pareudriline Genus Stuhlmannia. Unpublished Ph.D. Thesis. Bristol, UK, 691 pp. Jamieson, B. G. M. 1965. Two Ocnerodrilinae (Megascolecidae, Oligochaeta) from East Africa: Nematogenia lacuum (Beddard 1893) and Pygmaeodrilus montiskenyae sp. n. Annals and Magazine of Natural History 8: 95-107. Jamieson, B. G. M. 1967. A taxonomic review of the African megadrile genus Stuhlmannia (Eudrilidae, Oligochaeta). Journal of Zoology (London) 152: 79-126. Jamieson, B. G. M. 1968a. Macquaridrilus: a new genus of Tubificidae (Oligochaeta) from Macquarie Island. University of Queensland papers. Department of Zoology 3: 55-69. Jamieson, B. G. M. 1968b. A taxonometric investigation of the Alluroididae (Oligochaeta). Journal of Zoology (Lond.) 155: 55-86. Jamieson, B. G. M. 1969. A new Egyptian species of Chuniodrilus (Eudrilidae, Oligochaeta) with observations on internal fertilization and parallelism with the genus Stuhlmannia. Journal of Natural History 3: 41-51. Jamieson, B. G. M. 1970. A taxonomic revision of the oligochaete genus Eukerria Michaelsen 1935 (Ocnerodrilinae, Megascolecidae). Bulletin of the British Museum (Natural History) Zoology 20: 131-172. Jamieson, B. G. M. 1971a. Alluroididae. Pp. 708-722. In R. O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto. Jamieson, B. G. M. 1971b. Anatomy: Glossoscolecidae. Pp. 41-72. In R. O. Brinkhurst and B. G. M. Jamieson (eds), Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto. Jamieson, B. G. M. 1971c. Glossoscolecidae. Pp. 147-199. In R. O. Brinkhurst and B. G. M. Jamieson (eds), The Aquatic Oligochaeta of the World, Oliver and Boyd, Edinburgh, Toronto. Jamieson, B. G. M. 1971d. A review of the megascolecoid earthworm genera (Oligochaeta) of Australia. Part I—Reclassification and checklist of the megascolecoid genera of the world. Proceedings of the Royal Society of Queensland 82: 75-86. Jamieson, B. G. M. 1971e. A review of the megascolecoid earthworm genera (Oligochaeta) of Australia. Part II—The subfamilies Ocnerodrilinae and Acanthodrilinae. Proceedings of the Royal Society of Queensland 82(8): 95-108. Jamieson, B. G. M. 1971f. A review of the megascolecoid earthworm genera (Oligochaeta) of Australia. Part III—The subfamily Megascolecinae. Memoirs of the Queensland Museum 16(1): 69-102. Jamieson, B. G. M. 1974. The zoogeography and evolution of Tasmanian Oligochaeta. Pp. 195-228. In W. Williams (ed), Biogeography and Ecology in Tasmania, Dr. W. Junk, The Hague. Jamieson, B. G. M. 1977a. Marine meiobenthic Oligochaeta from Heron and Wistari Reefs (Great Barrier Reef) of the genera Clitellio, Limnodriloides and Phallodrilus (Tubificidae) and Grania (Enchytraeidae). Zoological Journal of the Linnean Society 61: 329-349. Jamieson, B. G. M. 1977b. On the phylogeny of the Moniligastridae with description of a new species of Moniligaster (Oligochaeta, Annelida). Evolutionary Theory 2: 95-114.

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Michaelsen, W. 1928. Oligochaeta. Pp. 1-118. In W. Kukenthal and T. Krumbach (eds), Handbuch der Zoologie. 2. (Plus appendices dated 1930 1932), Walter de Gruyter and Co, Berlin. Moon, S. Y., Kim, C. B., Gelder, S. R. and Kim, W. 1996. Phylogenetic positions of the aberrant branchiobdellidans and aphanoneurans within the Annelida as derived from 18S ribosomal RNA gene sequences. Hydrobiologia 324(3): 229-236. Nagase, I. and Nomura, E. 1937. On the Japanese aquatic Oligochaeta Criodrilus miyashitai n. sp. Scientific Reports of Tohoku Imperial University 11: 361-402. Nakamoto, A., Arai, A. and Shimizu, T. 2000. Cell lineage analysis of pattern formation in the Tubifex embryo. II. Segmentation in the ectoderm. International Journal of Developmental Biology 44(7): 797-805. Nordheim, H. von 1989. Vergleichende Ultrastrukturuntersuchungen der Eu- und Paraspermien von 13 Protodrilus-Arten (Polychaeta, Annelida) und ihre taxonomische und phylogenetische Bedeutung. Helgoländer Meersuntersuchungen 43:113-156. Nordheim, H. von 1990. Anatomie, Ultrastruktur und Systematik der gattung Protodrilus (Annelida, Polychaeta). Ph. D. Thesis, Universität Osnabrück, Osnabrück, Germany. Nuutinen, V. and Butt, K. R. 1997. The mating behaviour of the earthworm Lumbricus terrestris (Oligochaeta: Lumbricidae). Journal of Zoology London 242(4): 783-798. Nylander, J. A. A., Erséus, C. and Kallersjo, M. 1999. A test of monophyly of the gutless Phallodrilinae (Oligochaeta, Tubificidae) and the use of a 573-bp region of the mitochondrial cytochrome oxidase I gene in analysis of annelid phylogeny. Zoologica Scripta 28(3-4): 305-315. Omodeo, P. 1951. Raddoppiamento del corredo cromosomico nelle cellule germinali femminili di alcuni lombrichi. Atti Accademia Fisiocritici Siena 19: 14. Omodeo, P. 1952. Cariologia dei Lumbricidae. Caryologia 4: 173-275. Omodeo, P. 1955. Cariologia dei Lumbricidae. II contributo. Caryologia 8: 135-178. Omodeo, P. 1958. Oligochètes, In: La réserve intégrale du Mont Nimba. Mémoires de l’Institut Français d’Afrique Noire 53: 1-109. Omodeo, P. 1996. Kathrynella, a new oligochaete genus from Guyana. Hydrobiologia 334(1-3): 11-15. Omodeo, P. 1998. History of Clitellata. Italian Journal of Zoology 65: 51-73. Omodeo, P. 2000. Evolution and biogeography of megadriles (Annelida, Clitellata). Italian Journal of Zoology 67: 179-201. Omodeo, P. and Coates, K. A. New alluroidids (Annelida, Clitellata) from Guyana. Hydrobiologia 463(39): 39-47. Pickford, G. E. 1937. A Monograph of the Acanthodriline Earthworms of South Africa, Cambridge, UK. Pinder, A. M. and Brinkhurst, R. O. 1994. A preliminary guide to the identification of the microdrile Oligochaeta of Australian inland waters Cooperative Research Centre for Freshwater Ecology, Ellis Street, Thurgoona, Albury, New South Wales 2640. Plisko, J. D. 1996a. Michalakus, a remarkable new genus of microchaetid earthworm from South Africa (Oligochaeta: Microchaetidae). Annals of the Natal Museum 37: 287-293, Plisko, J. D. 1996b. Six new earthworm species of the southern African genus Proandricus Plisko 1992 (Oligochaeta: Microchaetidae). Annals of the Natal Museum 37: 295-307.

!' Reproductive Biology and Phylogeny of Annelida Pop, A. A., Wink, M. and Pop, V. V. 2003. Use of 18S, 16S rDNA and cytochrome c oxidase sequences in earthworm taxonomy (Oligochaeta, Lumbricidae). Pedobiologia 47: 428-433. Purschke, G., Westheide, W., Rohde, D. and Brinkhurst, R. O. 1993. Morphological reinvestigation and phylogenetic relationship of Acanthobdella peledina (Annelida, Clitellata). Zoomorphology Berlin 113(2): 91-101. Qiu, J. -P. and Bouché, M. B. 1998. Classified list of earthworms (Oligochaeta: Lumbricoidea), after the study of three fifth[s] of them. Documents Pédozoologiques et Intégrologiques 4(17): 181-200. Ramisch, H. and Graff, O. 1985. The cocoon chambers of some earthworms (Lumbricidae: Oligochaeta) from the Brunswick area (Lower Saxony) (West Germany). Braunschweiger Naturkundliche Schriften 2(2): 299-308. Righi, G. 1995. Colombian earthworms. Studies on Tropical Andean Ecosystems 4: 485-607. Righi, G., Ayres, I. and Bittencourt, C. R. 1978. Oligochaeta (Annelida) do Insituto nacional de Pesquisas da Amazonia. Acta Amazonica 8(3 Supplement 1): 1-49. Righi, G. and Varela, M. E. 1983. Narapa bonettoi, gen. nov. sp. nov. (Oligochaeta, Narapidae, fam. nov.) de agua doce da Argentina. Revista De La Asociacion De Ciencias Naturales Del Litoral 14(1): 7-15. Rota, E. and Brinkhurst, R. O. 2000. Mesenchytraeus antaeus, a new giant enchytraeid (Annelida, Clitellata) from the temperate rainforest of British Columbia, Canada, with a revised diagnosis of the genus Mesenchytraeus. Journal of Zoology London 252(1): 27-40. Rota, E. and Omodeo, P. 1992. Phylogeny of Lumbricina: re-examination by an authoritative process. Soil Biology and Biochemistry 24: 1263-1277. Rota, E., Martin, P. and Erséus, C. 2001. Soil-dwelling polychates: enigmatic as ever? Some hints on their phylognetic relationships as suggested by a maximum parsimony analysis of 18S rDNA gene sequences. Contributions to Zoology 70: 127-138. Rouabah Sadaoui, L. and Marcel, R. 1995a. Analysis of proteinic nutrients in clitellum and cocoon’s albumen in Eisenia fetida Sav (Annelida Oligochaeta). Evidence for a vitellogenin-like glycolipoprotein. Reproduction Nutrition Development 35(5): 491-501. Rouabah Sadaoui, L. and Marcel, R. 1995. Glucids and lipids of clitellum and cocoon’s albumen in Eisenia fetida Sav (Annelida Oligochaeta). Reproduction Nutrition Development 35(5): 537-548. Rouse, G. W. and Jamieson, B. G. M. 1987. An ultrastructural study of the spermatozoa of the polychaetes Eurythoe complanata (Amphinomidae), and Clymenella sp., and Micromaldane sp. (Maldanidae), with definition of sperm types in relation to reproductive biology. Journal of Submicroscopic Cytology 19: 573-584. Schmelz, R. M. 2003. Taxonomy of Fridericia (Oligochaeta, Enchytraeidae). Revision of species with morphological and biochemical methods. Abhandlungen des Naturwissenschaftlichen Vereins in Hamburg. Vol. 38. O. Kraus (ed.). Goecke and Evers, Keltern-Weiler, Germany. 415+73 plates. Shay, J. W. 1972. Ultrastructural observations on the acrosome of Lumbricus terrestris. Journal of Ultrastructure Research 41: 572-578. Shimizu, T. 1976. The fine structure of the Tubifex egg before and after fertilization. Journal of the Faculty of Science of Hokkaido University Series IV 20: 253-262.

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Shimizu, T. 1981. Cortical differentiation of the animal pole during maturation division in fertilized eggs of Tubifex (Annelida, Oligochaeta): I. Meiotic apparatus formation. Developmental Biology 85: 65-76. Shimizu, T. 1982. Development in the freshwater oligochaete Tubifex. Pp. 283-316. In F. W. Harrison and R. R. Cowden (eds), Developmental Biology of Freshwater Invertebrates. Alan R. Liss, Inc., New York. Shimizu, T., Kitamura, K., Arai, A. and Nakamoto, A. 2001. Pattern formation in embryos of the oligochaete annelid Tubifex: Cellular basis for segmentation and specification of segmental identity. Hydrobiologia 463(123): 123-131. Siddall, M. E. 1998. Success of parsimony in the four-taxon case: Long-branch repulsion by likelihood in the Farris Zone. Cladistics, 14: 209-220. Siddall, M. E. and Burreson, E. M. 1998. Phylogeny of leeches (Hirudinea) based on mitochondrial cytochrome c oxidase subunit I. Molecular Phylogenetics and Evolution. Feb. 9(1): 156-162. Siddall, M. E., Apakupakul, K., Burreson, E. M., Coates, K. A., Erséus, C., Gelder, S. R., Kallersjo, M. and Trapido, R. H. 2001. Validating Livanow: Molecular data agree that leeches, branchiobdellidans, and Acanthobdella peledina form a monophyletic group of oligochaetes. Molecular Phylogenetics and Evolution 21(3): 346-351. Siekierska, E. 2003. The structure of the ovary and oogenesis in the earthworm, Dendrobaena veneta (Annelida, Clitellata).Tissue and Cell. 35(4): 252-259. Silberglied, R. E., Shepherd, J. G. and Dickinson, J. L. 1984. Eunuchs: the role of apyrene sperm in Lepidoptera. American Naturalist 123: 255-265. Sims, R. W. 1966. The classification of the megascolecid earthworms: an investigation of oligochaete systematics by computer techniques. Proceedings of the Linnean Society of London 177: 125-141. Sims, R. W. 1967. Earthworms (Acanthodrilidae and Eudrilidae) from Gambia. Bulletin of the British Museum of Natural History 16: 1-43. Sims, R. W. 1980. A classification and the distribution of earthworms, suborder Lumbricina (Haplotaxida: Oligochaeta). Bulletin of the British Museum of Natural History (Zoology) 39(2): 103-124. Sims, R. W. 1982. Lumbricina. Pp. 55-61. In S. P. Parker (ed), Synopsis and Classification of Living Organisms. McGraw-Hill, New York. Stacey, D. F. and Coates, K. A. 1996. Oligochaetes (Naididae, Tubificidae, Opistocystidae, Enchytraeidae, Sparganophilidae and Alluroididae) of Guyana. Hydrobiologia 334(1-3): 17-29. Stephenson, J. 1922. Contributions to the morphology classification, and zoogeography of Indian Oligochaeta. Proceedings of the Zoological Society of London 1922: 109-148. Stephenson, J. 1930. The Oligochaeta. Clarendon Press, Oxford. Struck, T., Hessling, R. and Purschke, G. 2002. The systematic position of two enigmatic oligochaete-like taxa of the Polychaeta based on molecular data: Aeolosomatidae and Parergodrilidae. Journal of Zoological Systematics and Evolutionary Research 40(3): 155-163. Takashima, Y. and Mawatari Shunsuke, F. 1998. Mitinokuidrilus excavatus n. g., n. sp., a marine tubificid (Oligochaeta) with a unique mode of reproduction. Zoological Science Tokyo 15(4): 593-597. Teisaire, E. S. and Del, C. Q. M. 1989. Determination of the ATP concentration in spermatozoa stored in the spermathecae of Amynthas hawayanus (Oligochaeta: Megascolecidae). Comunicaciones Biologicas 8(2): 115-120.

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Teisaire, E. S. and Roldan, I. A. 1995. Ultrastructure and histochemistry of the spermatheca of Amynthas Kinberg and Metaphire Sims and Easton (Oligochaeta: Megascolecidae). Comunicaciones Biologicas 13(2): 169-181. Tétry, A. 1934. Description d’une espèce française du genre Pelodrilus (Oligochètes). Contes Rendues Hebdomadaires des Séances de l’Académie, Paris 199: 322. Troyer, D. 1980. Spermiogensis in Lumbricid earthworms revisited. II—Elongation and shortening of the spermatid nucleus and the roles of microtubules and chromatin in organelle shaping. Biologie Cellulaire 37: 287-292. Troyer, D. and Cameron, M. L. 1980. Spermiogensis in Lumbricid earthworms revisited. I—Function and fate of centrioles, fusion of organelles and organelle movements. Biologie Cellulaire 37: 273-286. Valvassori, R., de Eguileor, E. M., Grimaldi, A. and Lanzavecchia, G. 1999. Nematomorpha. Pp. 271. In B. G. M. Jamieson (ed), Reproductive Biology of Invertebrates. Progress in Male Gamete Ultrastructure and Phylogeny, John Wiley & Sons, Chichester. Van Praagh, B. D. 1995. Reproductive biology of Megascolides australis McCoy (Oligochaeta : Megascolecidae). Australian Journal of Zoology 43: 489-507. Walsh, M. P. 1954. Spermatogenesis of Lumbricus terrestris (L.). Transactions of the American Microscopical Society 73: 59-65. Wasawo, D. and Omodeo, P. 1963. Some arboricolous Oligochaeta from Ivory Coast. Memorie del Museo Civico di Storia Naturale di Verona 11: 211-223. Webster, P. M. and Richards, K. S. 1977. Spermiogenesis in the enchytraeid Lumbricillus rivalis (Oligochaeta: Annelida). Journal of Ultrastructure Research 61: 62-77. Welsch, U., Storch, V. and Richards, K. S. 1984. Annelida: Epidermal cells. Pp. 269296. In J. Bereiter-Hahn, A. G. Matoltsy and K. S. Richards (eds), Biology of the Integument, Springer-Verlag, New York. Westheide, W. 1988. Genital organs. Pp 263-279. In W. Westheide and C. O. Hermans (eds), The Ultrastructure of the Polychaeta, Microfauna Marina (ed. P. Ax), 4. Gustav Fischer, Stuttgart, New York. Westheide, W. 1999. Ultrastructure and functional significance of intestinojunctional spermathecae in enchytraeids (Oligochaeta, Annelida). Hydrobiologia 406: 199211 Westheide, W., Purschke, G. and Middendorf, K. 1991. Spermatozoal ultrastructure of the taxon Enchytraeus (Annelida, Oligochaeta) and its significance for species discrimination and identification. Zeitschrift Fuer Zoologische Systematik und Evolutionsforschung 29(5-6): 323-342. Winnepenninckx, B. M. H., Van, D. P. Y. and Backeljau, T. 1998. Metazoan relationships on the basis of 18S rRNA sequences: A few years later. American Zoologist. Dec. 38(6): 888-906. Yamaguchi, H. 1953. Studies on the aquatic Oligochaeta of Japan VI. A systematic report with some remarks on the classification and phylogeny of the Oligochaeta. Journal of the Faculty of Science of Hokkaido Imperial University Series VI. Zoology 11: 277-342. Zicsi, A. 1997. Contribution to the knowledge to the earthworm fauna of East Africa (Oligochaeta: Eudrilidae), with description of a new species of Polytoreutus. Revue Suisse de Zoologie. 104 (4): 807-820.

CHAPTER

9

Hirudinida Mark E. Siddall1, Alexandra E. Bely2, and Elizabeth Borda1

9.1 PHYLOGENY AND SYSTEMATICS Leech phylogenetic relationships and, consequently, classification of its constituents has seen considerable attention in the last decade particularly as leeches have been the subject of analyses at several taxonomic levels using morphological characters and DNA sequence data. The origin of leeches and other symbiotic clitellate annelids was at one time an issue rather hotly debated by annelid systematists. As with many annelids, leeches are soft-bodied and do not regularly leave a fossil record. There are two putative Jurrasic fossils from Bavarian deposits, Epitrachys rugosus and Palaeohirudo eichstaettensis, but neither has both the caudal sucker and annular subdivisions that together would definitively suggest a leech (Ehlers 1869; Kozur 1970). Nonetheless there have long been anatomical clues regarding hirudinidan origins. Leeches have a constant number of somites and a posterior sucker used for attachment to hosts, but so too do the tiny branchiobdellidan crayfish worms and the Arctic salmon worm Acanthobdella peledina. The latter has oligochaete-like chaetae and a constant number of 29 somites but exhibits leech-like coelomic and reproductive structures. In contrast, the branchiobdellidans have a more oligochaete-like reproductive organization, a constant number of 15 body somites and yet lack chaetae altogether. Not surprisingly there have been several historical suggestions of a close relationship amongst these groups (Odier 1823; Livanow 1931; Jamieson 1988; Brinkhurst and Gelder 1989; Siddall and Burreson 1996) but others worried that the similiarities were mere convergence (Holt 1989; Purschke et al. 1993; Brinkhurst,1994). Ferraguti and Erséus (1999) suggested several synapomorphies in sperm ultrastructure corroborating a sister-group relationship of leeches and Acanthobdella, whereas they found no evidence in support of an exact position for Branchiobdellida within the Clitellata (but see formation of the preacrosomal vesicle in spermatogenesis below). Siddall et al. (2001) demonstrated with nuclear and mitochondrial gene 1 2

American Museum of Natural History, New York, New York 10024, USA Department of Biology, University of Maryland, College Park, Maryland 20742, USA

!'" Reproductive Biology and Phylogeny of Annelida sequences that leeches (Hirudinida), branchiobdellidans and acanthobdellidans are a monophyletic “oligochaete” group that shares a common ancestor with Lumbriculida (Fig. 9.1). Consequently each should have equivalent ordinal rank (i.e., Hirudinida, Branchiobdellida and Acanthobdellida) in the class Oligochaeta or Clitellata. Ultimately the choice of “Oligochaeta” or “Clitellata” for the class is an arbitrary one since the taxa are synonymous in a phylogenetic sense. Use of “Clitellata” herein reinforces the actual synapomorphy shared by leeches and oligochaetes inasmuch as oligochaetousness is paraphyletic. On the other hand, use of “Oligochaeta” would reinforce the notion that leeches are members of that clade, not separate from it: a notion many find irksome—in itself a reason to insist upon it. Leeches are themselves subdivided first into suborders based on anatomical adaptations for feeding. “Rhynchobdellida”, as the name implies, is a group possessing a muscular proboscis to effect bloodfeeding from vascularized subdermal tissues. The Giant Amazonian leech, Haementeria ghilianii, which grows to a tremendous 16 inches (= 40 cm), has a proboscis that is nearly half its body length. Three families of proboscisbearing leeches are: the strongly dorsoventrally flattened freshwater Glossiphoniidae (Fig. 9.2A); the mostly marine Piscicolidae feeding seasonally on fishes, and the Ozobranchidae specializing on (usually) marine turtles (Fig. 9.2b). Arhynchobdellida, of which Hirudo medicinalis (Fig. 9.2C) is typical, are larger, vermiform and usually have three muscular jaws, each of which may be armed with a row of teeth creating a serrated cutting edge that allow them to feed through the skin on capillary-rich

Fig. 9.1 Ordinal level phylogeny of Clitellata with symbiotic lineages represented by thick lines. After Siddall et al. 2001 and Jamieson et al. 2002.

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Colour Figure

Fig. 9.2 A. The glossiphoniid leech Placobdelloides jaegerskioeldi feeding from its preferred attachment site, the rectal tissues of Hippopotamus amphibius. B. The marine turtle leech Ozobranchus margoi sporting lateral appendages of uncertain function. C. A very colorful species of Hirudo of blood-letting infamy engaging in that most infamous act on the third author’s finger. D. A Malagasy terrestrial haemadipsid in the genus Malagabdella making an incision on the first author’s hand. Original.

tissues. The large aquatic Hirudinidae (“medicinal leeches”) and the smaller terrestrial Haemadipsidae (“jungle leeches”, Fig. 9.2D) are perhaps the best known blood feeding arhynchobdellids. Both of these groups are equipped with a parabolic arc of 10 eyespots that detect movement in three dimensions. Terrestrial leeches have the additional adaptation of respiratory auricles near their caudal sucker that permit gas exchange without excessive loss of fluid and well-developed sensory systems for detecting vibrations, carbon dioxide and heat. As well there are several predatory arhynchobdellids like the slender Erpobdelliformes (families Erpobdellidae, “Salifidae” and Americobdellidae), the larger amphibious Haemopidae and various other poorly understood families like the Cylicobdellidae and Semiscolescidae. The evolutionary relationships of leeches have been investigated using morphological data (Siddall and Burreson 1995), life history characters (Siddall and Burreson 1996), nuclear and mitochondrial gene sequences (Siddall and Burreson 1998; Trontelj et al. 1999; Siddall et al. 2001), as well as combinations of these data sets at the familial level (Apakupakul et al. 1999; Light and Siddall 1999; Siddall 2002; Borda and Siddall 2004) and for

!'$ Reproductive Biology and Phylogeny of Annelida individual genera (Siddall and Borda 2003). A concatenation of recent phylogenetic datasets (Fig. 9.3) reveals the artificiality of many traditional groupings. Accepting the presence of a proboscis as an unreversed synapomorphy for the Rhynchobdellida (e.g. Livanow 1931; Mann 1962; Sawyer 1986; Siddall and Burreson 1995) had not been controversial. However, analyses that have used DNA sequence data consistently suggest these rhynchobdellids are paraphyletic (Siddall and Burreson 1998; Apakupakul et al. 1999; Trontelj et al. 1999) with Glossiphoniidae diverging from other leeches first, leaving Piscicolidae as sister to the arhynchobdellids (Fig. 9.3). In retrospect there previously have been several suggestions of loss of the proboscis en route to the more “advanced” medicinal leeches (e.g. Apathy 1888) as well as a basal position for Glossiphoniidae (Selensky 1907; Autrum 1939). Moreover there are several aspects of development and developmental regulation that indicate a suite of plesiomorphies retained in Glossiphoniidae (see section 9.5). Among the various families of leeches (Fig. 9.3), characteristics that typically have been considered diagnostic often prove to not be so. The notion that five pairs of eyespots arranged in a parabolic arc is a synapomorphy for Hirudiniformes is refuted by its presence in Linta be and (possibly) Americobdella valdiviana that constitute the basal most taxa of Erpobdelliformes. Small terrestrial jungle leeches likewise form two independent clades corroborating the notion that a portion of Haemadipsidae stands as a family in its own right: Xerobdellidae. Even the so-called medicinal leeches are not monophyletic. The predominantly New World Macrobdellidae stands apart from an exclusively Old World Hirudinidae, the latter of which is more closely related to the nonbloodfeeding genus Haemopis (for which, however, the parent family, Haemopidae, is polyphyletic insofar as the Semiscolescinae group with the new world hirudinids). Obviously reliance on bloodfeeding bahaviour has muddled leech systematics for some time as seen from the cladogram in Fig. 9.3 that requires the loss of sanguivory at least six times. Awkwardly, other characteristics that were deemed unreliable in the past prove to be extremely consistent in analyses of combined data sets. For example, the possession of two pairs of compact salivary cells at the base of the proboscis unites Haementeria species with Placobdella species within Glossiphoniidae notwithstanding their previously having been placed in separate subfamilies (Sawyer 1986). Similarly the arrangement of eyespots in erpobdellids was the only consistent morphological character for that group (Siddall 2002). Evolutionary relationships among leeches (Fig. 9.3) demonstrate that the ancestral hirudinid was a blood feeder in a freshwater environment suggesting that they are no older than vertebrates and probably are no older than the amphibian lineage. Corroborating this are the dietary preferences of basal lineages in Glossiphoniidae and Piscicolidae with Marsupiobdella africana still feeding on pipid frogs and ozobranchids on turtles. Moreover, several leech species have been examined for

Hirudinida

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Fig. 9.3 Phylogeny of leeches using the combined information from morphology and the genetic loci 28S rDNA, 18S rDNA, mitochondrial 12S rDNA CO-I, and ND-I, in which black lineages are most parsimoniously optimized as historically blood-feeding. Original.

!'& Reproductive Biology and Phylogeny of Annelida anticoagulants and, in terms of the phylogeny of the group (Fig. 9.3), it is clear that three coagulation inhibitors must have been inherited from the common ancestor. That is, a broad range of leeches have all inherited from the ancestral leech, genes coding for anti-platelet, anti-thrombin and antimetastatic anti-Xa factors. Probably many of the lineages that later gave up blood feeding have as well. Already this has been corroborated by the discovery of anti-Xa guamerins in the macrophagous hirudinid, Whitmania edentula. What lies undiscovered in other non-bloodfeeding groups remains an exciting prospect. Notably, even though the well-known medicinal leeches are associated with freshwater habitats, Fig. 9.3 implies that that the ancestral hirudiniform unequivocally was terrestrial. Aquatic hirudinid sexual biology would seem to corroborate this terrestrial ancestry. Unlike most rhynchobdellid and erpobdellid leeches that mate by way of traumatic insemination (hypodermic implantation of a membrane-bound spermatophore that injects sperm in response to an osmotic pressure change), the haemadipsids and other hirudiniforms are characterized by internal fertilization (gonopore to gonopore copulation with a protrusible penis and a compensatory vagina). As well, aquatic hirudinids and macrobdellids still deposit their cocoons on land; hatchling leeches must find their way to nearby water when they emerge. In terms of biogeography, the evolutionary history of leeches is perhaps most remarkable for the anomalies implied by Fig. 9.3, though several items offer easy interpretation. The majority of the basal-most members of Glossiphoniidae are African (i.e., Marsupiobdella africana, Placobdelloides multistriatus, Batracobdelloides tricarinata, Oosthuizobdella garoui, and Placobdelloides jagerskioeldi). Furthermore, subsequent diversification, proximal in time, involves the overwhelmingly South American genus Helobdella (though many North American taxa are known) and the exclusively South American Haementeria. The foregoing suggests a Gondwanan origin for that family, and this is somewhat consistent elsewhere with finding the Malagasy Linta species and South American Cylicobdella species at the base of their respective groups (i.e., Erobdelliformes and Hirudiniformes). However, several arrangements mitigate against such an easy interpretation. The North American Placobdella radiation stands wholly as sister to the South American group. A post-Gondwanan Panamanian isthmus explanation for the origin of the former would only be sensible only if Placobdella were nestled within Haementeria (much as how Macrobdella species are few and nestled within a paraphyletic South American clade of Oxyptychus, Semiscolex and Patagoniobdella species, and that the North American species of Helobdella arise in several places from a basal South American stock); and would require the complete speciation of the group plus a leap to Europe (for Placobdella costata) in a very short 3.5 million years. There are in fact several relatively recent sister group relationships consistent with a Laurasian connection. The European pair Haemopis

Hirudinida

!''

sanguisuga and Haemopis caeca are sister to the North American clade of Haemopis comprising the balance of that genus. The European pair Erpobdella lineata and Erpobdella mestrovi are sister to a North American Erpobdella clade. Similarly, the balance of that genus comprises only North American, European and Asian taxa. Also, the North American Glossiphonia elegans (frequently misnamed G. complanata which is European) is sister to the Eurasian fauna in this genus. If this is to be believed, one might find it awkward to postulate a series of Laurasian associates grouping within (as opposed to adjacent to) Gondwanan associations in so far as both land masses are supposed to have been late Paleozoic contemporaries. Other relationships defy explanation at all, save perhaps through ad hoc invocations of massive extinction or extremely long distance dispersal by leeches that do not seem capable of such a feat. In Xerobdellidae in particular, taxa included in this analysis are the Alpine Xerobdella lecomtei and the Chilean Mesobdella gemmata. Lest taxon sampling alone be thought to explain this extreme disjunction, the remaining taxa in the family are known only from Mesoamerica.

9.2 ANATOMY WITH REFERENCE TO THE REPRODUCTIVE SYSTEM Among the various diagnostic morphological characteristics for leeches, the reproductive system has been particularly important in leech taxonomy and phylogenetics (Richardson 1969 1976; Ringuelet 1985; Sawyer 1986; Siddall and Burreson 1995; Apakupakul et al. 1999; Siddall 2001a, b; Siddall and Borda 2003; Borda and Siddall 2004). The reproductive anatomy of leeches is variable across the group, and can be quite useful for recognizing some higher taxa and even (in some families) to distinguish among species. Like other clitellate oligochaetes, leeches possess a clitellum, the saddlelike glandular region associated with cocoon deposition in the anterior portion of the body. The prominence of the clitellum is variable in leeches (usually somites X to XIII; that is, the eighth through eleventh true segments, or M4-M7) and often is not evident externally in most species, compared to the pronounced swollen clitellar region typified by the lumbricid oligochaetes (Fig. 9.4). Leeches are hermaphrodites, with simultaneous possession of independent male and female reproductive systems. The male median reproductive apparatus is found anterior to the female median reproductive apparatus and each possesses a separate ventral opening to the exterior, or gonopore, on clitellar somites XI and XII respectively. In general, the male reproductive system consists of an atrium (or bursa), either fused (unpaired) or paired (bilobed), and sperm ducts of varying thickness. Some species are characterized by the presence of ejaculatory bulbs, coiled “epididymes”, and a protrusible penis. The testisacs are derived from coelomic epithelial sacs and usually are paired

" Reproductive Biology and Phylogeny of Annelida

Fig. 9.4 A. A large glossoscolescid oligochaete from the Andes in Boliva equipped with a large and prominent clitellum. B. The erpobdelliform leech Barbronia gwalagwalensis from South Africa with a noticeable clitellum. C. Hirudo medicinalis in which the clitellum is cryptic as it is in most species of leech. Original.

posteriorly along distal sperm ducts (one testisac on each side per somite). The female reproductive system consists of a pair of ovisacs. In some groups this is elaborated into a vagina and oviducts that connect the ovisacs to the vagina. Glossiphoniidae. The reproductive anatomy of glossiphoniid leeches (e.g. Helobdella spp., Haementeria spp., Placobdella spp.) is relatively simple in form (Fig. 9.5A, B), typically with laterally directed bilobed (paired) atria, which give rise to simple descending and ascending sperm ducts. Some glossiphoniids (i.e. Theromyzon spp.) have a male apparatus with a fused (unpaired) atrium and elongation of an eversible atrium (Wilkialis and Davies 1980; Siddall and Burreson 1995). The sperm ducts descend from the atrial lobes and ascend towards the atria before entering the parenchyma with a posterio-lateral arrangement of one pair of testisacs per somite (five to eight pairs total). The female reproductive system only has a pair of posteriorly directed tubular ovisacs. The posterior extents of ovisacs and sperm ducts in the central coelomic space are species-specific. Piscicolidae and Ozobranchidae. As with Glossiphoniidae, the fish leeches (freshwater and marine piscicolids) and the turtle leeches (ozobranchids) also have a relatively simple reproductive apparatus. Their male reproductive systems generally have an anteriorly directed bilobed bursa, with sperm ducts and ejaculatory bulbs curving posteriorly connected to one pair of testisacs per somite in a manner similar to glossiphoniids. The female system is also similar to that of the glossiphoniids in having long tubular ovaries. Most notably, piscicolids possess parenchymal conducting (vector) tissue that is utilized for the

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passage of sperm from the copulatory area to the female reproductive tissue (See Fig. 9.5C; Sawyer 1986; Siddall and Burreson 1995), whereas ozobranchids have bilateral coelomic tubes connecting the male and female reproductive systems (MacCallum and MacCallum 1918; Raj and Penner 1962) Erpobdelliformes. The reproductive anatomies of erpobdellid (Erpobdellidae) and salifid (Salifidae) leeches are quite similar to those already discussed for Glossiphoniidae, Piscicolidae, and Ozobranchidae. The male atrium is bilobed and can have sperm ducts with or without a preatrial loops (‘U’ shaped) that extend anteriorly from the atria, then descend posteriorly; ejaculatory bulbs are not present. As for the female system, they too have simple tubular ovaries. What is unique to Erpobdellidae and Salifidae, is the testisac arrangement. Contrasting the typically discrete bilateral paired arrangement per somite, salifid species have testisacs that are discretely arranged, but in somatic tetrads (two pairs of testisacs on each sperm duct) per somite, whereas erpobdellid leeches have an atypical arrangement of the testisacs in multiple grape-like clusters profusely arranged along the length of the sperm ducts (Sawyer 1986; Siddall and Burreson 1995; Siddall 2002; Borda and Siddall 2004). Americobdellidae. Some of the historical problems in classifying the anomalous Americobdella valdiviana were due to similarities in reproductive structures to erpobdellids, glossiphoniids and piscicolids (Moore 1924; Sawyer 1986; Siddall and Burreson 1995), though the leech is hirudiniform both in gross morphology and in feeding habit. Americobdella valdiviana has a reproductive anatomy that is unique in its own right (Fig. 9.5D). Americobdella valdiviana has a fused male atrium with an anterior seminal receptacle, awkwardly referred to as the “female” bursa (Moore 1924; Sawyer 1986). The sperm ducts are preceded by tightly coiled epididymes, followed by pairs of testisacs in nine somites. The female ovaries are tubular, much like in erpobdellids and the rhynchobdellid leeches. The most striking features, however, are the intergonadal coelomic tubules connecting the ovisacs with the female bursa (copulatory area), which may facilitate the passage of sperm (Fig. 9.5D). These have been considered homologous in form and function to the vector tissue known from piscicolid leeches (Moore 1924; Sawyer 1986; Siddall and Burreson 1995) and approach a morphology very similar to those in Ozobranchidae. Predictably, monotypic Americobdellidae is the basal-most lineage of erpobdelliform leeches. It is tempting to consider intergonodal conducting tissues to be a mechanism that allows self-fertilization, particularly because both the marine leeches and Americobdella valdiviana can exist in quite rarified population structures. Hirudiniformes. The so-called ‘medicinal’ leeches (e.g. Hirudo medicinalis and Macrobdella decora), other aquatic sanguivorous species (e.g. South American Oxyptychus spp.) and several macrophagous groups (species of Haemopis and Patagoniobdella) have all been classified under the family Hirudinidae (Blanchard 1896). Similarly, all terrestrial leeches were

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Reproductive Biology and Phylogeny of Annelida

Fig. 9.5 A. Dissected Helobdella wodzickiorum revealing typical glossiphoniid anterior paired ejaculatory atria from which descend and loop the male sperm ducts along the midline of the coelomic cavity. B. Dissected Helobdella wodzickiorum with the male sperm ducts removed revealing typical elongate glossiphoniid ovisacs (arrows). C. Diagrammatic representation of the median reproductive structures of piscicolids with conducting tissues (arrows) between the anterior bursa and the paired ovisacs. D. Dissected Americobdella valdiviana revealing the pair of intergonadal coelomic tubules (arrows) connecting the ovisacs (to the right) with the female bursa. E. Median reproductive anatomy of Aliolimnatis africana exhibiting globular oviscacs terminal to the paired and common oviduct, and the typically hirudiniform “epididymes” one either side of the protrsuible penis. F. Median reproductive anatomy of the terrestrial Mesobdella gemmata exhibiting the left globular oviscac paired “epididymes” and protrsuible penis. G. Median male atria of Cylicobdella coccinea. Abbreviations: e, epididymes; o, ovisacs; p, penis. Original.

once placed in Haemadipsidae. This classification was primarily based on feeding habit, habitat preference and/or gross external morphology. Dissection and examination of the reproductive anatomy of members of these families initiated a re-examination of their relationships, as well as a re-evaluation of their taxonomic status (Moore 1927; Richardson 1969, 1971, 1976; Ringuelet 1985; Sawyer 1986; Siddall and Burreson 1995; Apakupakul et al. 1999; Borda and Siddall 2004). Compared to the relatively simple reproductive anatomies already discussed, the hirudiniform leeches (Haemadipsidae, Haemopidae, Hirudinidae, Macrobdellidae, Semiscolecidae, and Xerobdellidae) have a considerably more ‘complex’

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anatomy, coincident with their copulatory mating behaviour and with as much variation in structure as they are ecologically diverse. The male reproductive apparatus of hirunidform leeches generally has a fused atrium with posterior elongation of the atrium into a protrusible penis that recurves and ascends towards the male gonopore (Fig. 9.5E). The sperm ducts usually are equipped with bilateral ejaculatory bulbs adpressed to which are coiled epididymes. Typically there is one pair of testisacs in each of nine somites though species in Semiscolecinae may have two pairs of testisacs per somite much as was described for the Salifidae. The female reproductive apparatus is equipped with a pair of small bulbous ovisacs subtended by oviducts to the vagina, either independently (e.g. Mesobdella gemmata, Fig. 9.5F) or as a single common duct from the ovisacs (e.g. Aliolimnatis africana, Fig. 9.5D). Cylicobdellidae. Considering they are sister group to the rest of the Hirudiniformes, cylicobdellid leeches have a remarkably plesiomorphic reproductive anatomy. Borda and Siddall (2004) found that Cylicobdellidae was the basal-most lineage in Hirudiniformes and their anatomy seems to be consistent with this phylogenetic position. Cylicobdellid leeches appear to retain the reproductive anatomy found in Erpobdellidae. They have anteriorly directed, paired male atria with cornua (Fig. 9.5G), and primary coiling of the sperm ducts that extend posteriorly, looping anteriorly towards the atria before descending laterally. Unlike erpobdellid leeches (which this arrangement resembles) they have a single pair of testisacs in nine to 12 somites (much as described for Glossiphoniidae above). The female apparatus consists of a pair of simple tubular ovisacs as is typical for Erpobdelliformes and “Rhynchobdellida.”

9.3 GAMETOGENESIS Oogenesis and spermatogenesis in leeches each begins with the release of epithelial cells from the walls of specialized coelomic sacs. Testisacs are found intersegmentally on either side. Ovisacs typically are ventrally arranged along the midline. The released spermatogonia and oogonia float freely in the lumen of the coelomic sacs wherein they undergo mitotic divisions followed by meiosis and the haploid nuclei eventually are found arranged around the periphery of an anucleate cytophore to which the gametes remain connected. Unlike many other annelids, hirudinidan meiosis reportedly produces only a single viable oocyte receiving the bulk of the cytoplasm and remaining associated with three adjacent cells (Aisenstadt et al. 1967). The latter are true “nurse cells” inasmuch as there is considerable passage both of rRNA and proteinaceous material from them to the oocyte first by cytoplasmic streaming prior to cytokinesis and later pinocytically (Aisenstadt et al. 1967). Hirudinidan meiosis to spermatocytes typically follows six mitotic divisions resulting in 256 immature spermatocytes attached to the

"" Reproductive Biology and Phylogeny of Annelida cytophore (Malécha 1970; Lechenault and Pastisson 1973). There appears to be an additional mitotic event among erpobdelliformes (Bonet and Molinas 1988) though the precise number is unknown for most taxa. Maturation of the sperm in leeches, Acanthobdella and branchiobdellidans reflects their close phylogenetic affinities. All three groups possess an unusual dextrogyrously twisted acrosomal tube, the leading edge of which protrudes to form a marginal ridge (Ferraguti and Gelder 1991; Franzén 1991; Westheide and Purschke 1996; Malécha 1975; Bonet and Molinas 1988). Branchiobdellidans have either seven or four mitochondria whereas leeches and Acanthobdella each have only one per spermatozoon. Among leeches there is considerable consistency in the shape and maturation of the spermatozoon (Malécha 1975; Sawyer 1986, Bonet and Molinas 1988). In addition to the helical acrosome, all leeches have a dextrogyrously twisted nucleus with two or more helical ridges a well as a helical acrosomal tip (or anterior acrosome) that forms anterior to the invagination point of the proacrosomal vesicle. Following the second meiotic event, the large multilaminate U-shaped Golgi apparatus (from which the proacrosomal cap forms) is positioned between the nucleus and the cytophore with a single large mitochondrion and basal body at the distal pole. Elongation of the flagellum and the single mitochondrion proceeds unremarkably while the acrosomal protube begins to form in the collar region between the cap and the nucleus attachment zone. The protube elongates in association with a palisade of microtubules extending from a distinctive plate region where the tube contacts the nucleus. At the point in which the acrosomal tube is fully formed, several transformations unique to leeches occur. The terminal electron-dense cap is forced to one side as a fibrous sheath extends anteriorly to form the anterior acrosome while the surrounding microtubules change conformation inducing a rotational force. This rotation ultimately twists each of the acrosomal tip, the acrosome itself, the nucleus and, in some taxa (Wissocq and Malécha 1975; Bonet and Molinas 1988), the mitochondrion as well. The proacrosomal vesicle appears to consist of a single highly coiled Golgi tubule (Fig. 9.6A, B) both in leeches and in branchiobdellidans. The vesicle then invaginates into the acrosomal tube posteriorly with the acrosomal tip not receiving any acrosomal material. Finally a perforatorium-like structure assembles at the base. Maturation of the nucleus entails condensation of chromatin followed (at the rotational stage) by the longitudinal formation of two or more ridges such that the mature nucleus is either a double helix in the case of Hirudinidae and Erpobdelliformes or a triple helix among piscicolids (Fig. 9.6A, C) whereas the mature glossiphoniid spermatozoon has a more complex and geometric unequal double helical nucleus (Fig. 9.6D). Oocytes generally remain in an arrested state until fertilized while still in the ovisacs. Spermatozoa travel up the sperm ducts towards the male median structures where they are bundled with agglutinating secretions from associated glands and are packaged into spermatophores.

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Fig. 9.6 Hirudinidan-like spermatogenesis. A. Spermatozoa of Malmiana scorpii in mid-stage of maturation arranged around a central cytophore and exhibiting a coiled nucleus, an empty acrosomal tube, an early acrosomal tip as well as the highly coiled proacrosomal vesicle prior to invagination. B. Coiled proacrosomal vesicle of the branchiobdellidan Cronodrilus ogygius. C. Transverse section of spermatozoa of Malmiana scorpii in mid-stage of maturation exhibiting the flagellum, single mitochondrion, trihelical nucleus and empty acrosomal tube. D. Mature spermatozoa of Theromyzon tessulatum in longitudinal section revealing the complex geometry of the coiled nucleus. E. Diagrammatic representation of the relative structure and sizes of the five principal zones of a mature leech spermatozoon. Abbreviations: c, central cytophore; n, nucleus; pv, proacrosomal vesicle. B and D courtesy of Marco Ferraguti, others original.

"$ Reproductive Biology and Phylogeny of Annelida

9.4 MATING, FERTILIZATION AND PARENTAL CARE The most common form of sex in leeches (observed for most Glossiphoniidae and Erpobdelliformes) is traumatic insemination by way of packaged spermatophores implanted indiscriminately through the cuticle of a recipient mate (Fig. 9.7A). Presumably hydrostatic changes induce the spermatophore to empty its contents whereupon the spermatozoa cork-screw their way through the parenchyma and the coelomic spaces ultimately fertilizing oocytes in the ovisacs. Among piscicolids, spermatophore implantation is often site-specific to the ventral region of the clitellum where conducting (or “vector”) tissues (Fig. 9.5C) guide the sperm directly to the ovisacs. Among Hirudinformes mating is by copulation and insertion of a protrusible penis into a vaginal sac, the sizes and shapes of which tend to be species-specific (Fig. 9.5E, F). Parental care has two basic forms among leeches. The fish leeches, or Piscicolidae, exhibit an adaptation that promotes their offspring achieving an early blood meal. Rather than abandoning a secreted “cocoon” as the oligochaetes and arhynchobdellid leeches do (Fig. 9.7B), the piscicolids cement their egg cases to the surface of crustaceans (Fig. 9.7C, D). When that animal is later eaten, young leeches readily attach to the fish host’s buccal surfaces and migrate to the gills. Glossiphoniidae, such as Haementeria ghilianii, are broad and flattened, and normally found feeding on turtles or amphibians. Species in this family secrete a membranous bag holding their eggs on their underside in a brooding position underneath rocks and other debris (Fig. 9.7E). When the brood hatches, the young will turn and attach to the venter of their parent and, when the parent finds its next blood meal, they are carried to their first.

9.5 DEVELOPMENT Leeches stand out for being the best characterized annelid group, and arguably the best characterized lophotrochozoan group, with respect to cellular and molecular aspects of embryogenesis. As such, leeches have featured prominently in recent discussions of the evolution of development (Shankland and Seaver 2000). Clitellate embryogenesis is clearly derived in many respects, so it is fortunate that additional annelid groups are attracting the attention of a growing number of molecular and experimental developmental biologists. Still, the currently unparalleled experimental tractability of leech model systems and the deep base of knowledge already available for this group will doubtless keep leeches in the developmental spotlight for years to come. Leeches, like all other clitellates, develop directly from yolky eggs: there is no trace of a trochophore larva. All 32 segments are formed during embryogenesis and after hatching juveniles grow only in segment size, not segment number, in contrast to many other annelids which continue to add segments throughout their lives. (Note that the prostomium and

Fig. 9.7 A. Dorsal surface of Placobdella parasitica with three spermatophores embedded in its integument. B. Spongy cocoon of the hirudinid Hirudo medicinalis which that species deposits on land usually in damp vegetation. C. Scanning electron micrographs of the spheroid cocoon of the piscicolid leech Oxytonostoma typica showing the posterior operculum. D. Scanning electron micrographs of the flattened cocoon of the piscicolid leech Oceanobdella microstoma revealing both opercula (arrows). E. Scanning electron micrograph of Helodbella elongata revealing the membranous cocoon brooded on the venter. Original.

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"& Reproductive Biology and Phylogeny of Annelida peristomium are traditionally denoted somites I and II by leech systematists even though they do not constitute true developmental segments.) Embryonic development in the large yolky embryos of glossiphoniids (e.g. Helobdella, Theromyzon) closely resembles that in the large yolky embryos of several oligochaete groups (e.g. tubificids and lumbriculids). This suggests that glossiphoniids may show the plesiomorphic form of embryogenesis for Hirudinida. Albumenotrophy, in which embryos ingest the albumen provisioned in the cocoon rather than relying solely on yolk stores of the egg, occurs in several groups including the hirudiniforms, erpobdellids, and piscicolids. Early development in albumenotrophic species involves the formation of temporary embryonic feeding structures and other modifications that are clearly derived. However, later development in these groups is largely similar to development in yolky-egged species. The small non-blood feeding glossiphoniid leeches in the genus Helobdella produce large (~450 µm diameter) embryos that are amenable to a range of manipulations (e.g. intracellular injections of lineage tracers, intracellular injections of drugs and reagents to manipulate geneexpression, targeted cell ablations) offering tremendous advantages for experimental investigations of development. Studies of leech development have focused heavily on species of Helobdella, though considerable work has also been done on glossiphoniid Theromyzon species and on the hirudiniform Hirudo medicinalis.

9.5.1 Cellular Aspects of Leech Development: Cell Lineages and Cell Fates As in other annelids, cleavages in leeches are highly stereotyped with respect to timing, orientation, and cell size, making it possible to reproducibly identify many cells of the embryo and follow many of their cleavages. Cell lineages and cell fates have been most extensively investigated in species of Helobdella, primarily using injections of intracellular lineage tracers (see Weisblat and Huang 2001 for a recent review). After fertilization, meiosis of the egg is arrested at metaphase I until the zygote is deposited into the cocoon. Meiosis then resumes, two polar bodies are produced, and the two pronuclei fuse. Prior to the first cleavage, a series of cytoplasmic reorganizations assemble regions of yolk-free cytoplasm, called teloplasm, at the animal and vegetal poles of the zygote. This teloplasm contains cell-fate determinants and is enriched in maternal mRNAs, mitochondria and ribosomes (Astrow et al. 1987; Fernandez et al. 1987; Nelson and Weisblat 1992; Holton et al. 1994). Early cleavages in Helobdella give rise to three classes of cells: three large, yolk-rich vegetal cells called macromeres (the main endodermal precursors), 25 small animal cells called micromeres (which contribute primarily to asegmental head structures and a provisional embryonic epithelium), and 10 large cells called teloblasts (the precursors of all segmental mesoderm and ectoderm) (Figs. 9.8, 9.9).

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The spiralian cell division pattern is discernible in leech embryos through the first few cleavages, although important modifications are apparent even early on. The first two cleavages are roughly meridional and orthogonal to each other, dividing the animal into four large cells, A, B, C, and D, of which D is the largest and inherits the bulk of the teloplasm (Fig. 9.10A). The third cleavage is equatorial and highly unequal in all four quadrants, generating four large vegetal macromeres and four small animal micromeres (Fig. 9.8). Although this and several subsequent cell divisions show the spiralian pattern of alternation of sinistral and dextral cleavages, with micromeres produced towards the animal pole, glossiphoniid leech embryos show an important deviation beginning at this third division: divisions in the B quadrant are in the opposite direction from the typical spiralian pattern (Sandig and Dohle 1988; Weisblat and Huang 2001). Thus, the third and fourth B-quadrant divisions are sinistral and dextral, respectively, while they are respectively dextral and sinistral (the typical spiralian pattern) in the A, B, and C quadrants. The A and B quadrants are therefore mirror images of each other. Endoderm. The macromeres of the A, B, and C quadrants are the main precursors of the endoderm. In Helobdella, these three macromeres produce three micromeres each and then cease cleaving, arresting in the G2 phase of the cell cycle (Bissen and Weisblat 1989). Later in development, these macromeres fuse together in a stepwise manner to form a syncytial yolk cell

Fig. 9.8 Diagram of glossiphoniid leech development. A. Eight-cell embryo (animal view). B. 20-cell embryo (animal view). C. Beginning of germinal-plate formation (animal view). The left and right germinal bands coalesce along the future ventral midline to form the germinal plate in an anterior to posterior direction. The syncytial yolk cell (prospective endoderm) is shaded light gray; the micromerederived provisional epithelium which covers the germinal bands and germinal plate is shaded dark gray. D. Completion of germinal band and beginning of segment morphogenesis (lateral view). E. Juvenile (dorsal view). The midgut (endoderm) is shaded gray; the foregut (micromere-derived) is outlined in gray. Original.

" Reproductive Biology and Phylogeny of Annelida (Fig. 9.9) (Liu et al. 1998), a process at least partly dependent on signals from cells in the D quadrant (Isaksen et al. 1999). Late-stage teloblasts and some of their recent progeny (supernumerary blast cells) also eventually fuse with the syncytial yolk cell (Liu et al. 1998; Desjeux and Price 1999). The syncytial yolk cell ultimately cellularizes to form the midgut endoderm (crop, intestine, and rectum) surrounding the remaining yolk (Whitman 1878; Nardelli-Haefliger and Shankland 1993), which is digested prior to the juvenile’s first meal. Local signals from the mesoderm to the endoderm are critical for normal leech gut morphogenesis (Wedeen and Shankland 1997), as they also appear to be in the lumbriculid Eisenia (Devriès 1974). Unlike the situation in Helobdella, however, in several other leeches and other clitellates the D macromere contributes substantially to the presumptive endoderm (in addition to generating the teloblasts). Furthermore the macromeres do not cease cleaving early in development but rather continue dividing to form a multi-celled endodermal mass (Anderson 1973; Shimizu 1982). Thus, Helobdella may represent a derived condition in these two aspects of gut morphogenesis. Segmental ectoderm and mesoderm. All segmental ectoderm and mesoderm is derived from 10 large embryonic stem cells, called teloblasts, which are produced by the large D’ macromere (Fig. 9.9). D’ generates the 10 teloblasts and 15 additional micromeres through a unique series of cell divisions which are teloplasm-dependent (Astrow et al. 1987; Nelson and Weisblat 1991). The division of D’ is obliquely equatorial and forms a (more vegetal) mesodermal precursor cell (DM) which ultimately produces a left/ right pair of mesodermal teloblasts called mesoteloblasts, and a (more animal) ectodermal precursor cell (DNOPQ) which ultimately produces four left/right pairs of ectodermal teloblasts called ectoteloblasts. These five pairs of teloblasts form the left and right halves of the leech body, each half developing largely independently of the other. The sequence of divisions producing the ectoteloblasts and the relative positions of teloblasts vary among leech species (Fernandez and Stent 1982), but those of Helobdella resemble those of other glossiphoniids and the oligochaete Tubifex (Shimizu 1982; Sandig and Dohle 1988), suggesting that these may represent the plesiomorphic patterns for leeches, a notion consistent with the relationships in Fig. 9.3. On each side of the embryo, then, there is one mesodermal teloblast, designated M, and four ectoteloblasts, designated N, O/P, O/P, and Q. Each teloblast repeatedly divides asymmetrically producing small progeny cells, called primary blast cells, which are organized into a discrete column, or bandlet (Figs. 9.8, 9.10C). The M, N, and Q teloblasts generate blast cells that are from birth fated to be of the M, N, or Q type, respectively, but the O/P teloblasts are equivalent (Weisblat and Blair 1984; Zackson 1984). They generate bandlets which are only secondarily induced to adopt an O or P fate through interactions with neighbouring bandlets and the micromerederived provisional epithelium (Shankland and Weisblat 1984; Ho and Weisblat 1987; Huang and Weisblat 1996).

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Fig. 9.9 Summary of early cell divisions and ultimate cell fates in the glossiphoniid Helobdella robusta. For diagram clarity, cell divisions of only one of the two DNOPQ cells is included. O/P teloblasts produce o/p blast cells which acquire O and P fates through interactions with other bandlets. Dotted lines represent continued blast cell production by teloblasts. Supernumerary blast cells, like teloblasts, fuse with the syncytial yolk cell (not shown). The relative timing of fusion of teloblasts O, P, and Q and of supernumerary blast cells with the syncytial yolk cell is not known. Divisions of micromeres and blast cells are not included in this diagram. Leech cell nomenclature is used throughout; standard spiralian notation is indicated in parentheses for the first few cleavages. Diagram based on Weisblat, D.A. and Huang, F.Z. 2001. Canadian Journal of Zoology 79: 218–232, Fig. 2.

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Segment formation occurs in an anterior-to-posterior progression: the first blast cells produced give rise to the most anterior segmental tissue and subsequent blast cells give rise to progressively more posterior tissue. The segmental identities of blast cells are largely established at birth (Martindale and Shankland 1990; Gleizer and Stent 1993; Nardelli-Haefliger et al. 1994). Possibly reflecting the plesiomorphic clitellate phenomenon of indeterminate segment addition, leech teloblasts generate more than the number of blast cells required to build the 32 segments of the leech. These “supernumerary” blast cells fuse with the syncytial yolk cell late in development (Desjeux and Price 1999). The five bandlets on each side of the embryo come together to form left and right germinal bands, with the n, o, p, and q bandlets lying side by side and overlying the m-bandlets. The left and right germinal bands progressively move ventrally over the surface of the macromeres and coalesce to form the germinal plate (Figs. 9.8, 9.10B), with the two nbandlets straddling the future ventral midline. Finally, the lateral edges of the germinal plate extend dorsally and the left and right q-bandlet progeny fuse at the dorsal midline. The mass of endoderm is thus internalized, completing the formation of the leech’s body tube. Germ band formation, germinal plate formation, and dorsal closure appear to be quite similar among various clitellate annelids (Anderson 1973). Injections of cell lineage tracers into teloblasts have beautifully revealed the ultimate fates of cells in each teloblast lineage (e.g. Weisblat et al. 1978; Weisblat et al. 1980; Weisblat and Shankland 1985). Cells in each of the five types of bandlets divide in a unique stereotypical fashion and give rise to segmentally iterated sets of definitive progeny (Fig. 9.10D). Each blast cell in the m, o, and q bandlets produces one hemisegmental compliment of definitive progeny, while in the n and q bandlets a pair of consecutive blast cells (nf/ns and qf/qs) together produces one hemisegmental complement (Fig. 9.10D, E). The M lineage gives rise to body wall muscle, visceral mesoderm, septa walls (which are later lost), nephridia, a few ganglionic neurons, and probably the germ line. Each of the four ectodermal lineages gives rise to neurons and epidermis, the N lineage contributing mostly to ventral tissue (including most of the ventral nerve cord neurons) and the O, P, and Q lineages contributing primarily to progressively more dorsal tissue. Asegmental tissue and provisional epithelium. Initially restricted to the “micromere cap” at the embryo’s animal pole, the micromeres produced during early cleavages ultimately give rise to asegmental tissues, primarily of the head, and a provisional embryonic epithelium (Weisblat et al. 1984; Nardelli-Haefliger and Shankland 1993; Smith and Weisblat 1994; Huang et al. 2002). Definitive micromere-derived structures include the proboscis (foregut) epithelium, proboscis muscles (Fig. 9.10F), proboscis sheath, neurons and connective tissue of the cerebral (supraesophageal) ganglion, putative glial cells and connective tissue of the fused rostral (subesophageal) ganglia, and anterior and posterior sucker epithelium.

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Interestingly, a recent detailed cell-lineage analysis of micromeres reveals that some micromeres give rise to different definitive progeny in two closely related Helobdella species, attesting to the evolutionary lability of micromere cell fates (Huang et al. 2002). The micromere-derived provisional epithelium initially covers the animal pole and progressively spreads over the germinal bands (Fig. 9.10B). Although it does not produce any definitive structures, it sends critical signals to the underlying developing germinal bands and is required for proper germ band migration over the endoderm surface (Ho and Weisblat 1987; Smith et al. 1996). The provisional epithelium expands ventrally as the germinal bands coalesce ventrally, and then contracts dorsally and ultimately disappears during dorsal closure.

9.5.2 Molecular Aspects of Leech Development: Developmental Regulatory Genes Since the late 1980’s, when the first molecular studies of leech development were published, a wide range of developmental regulatory genes have been isolated from and characterized in leeches (Table 9.1). These genes include members of many gene families, including homeodomain transcription factors (Hox, ParaHox, engrailed, even-skipped, msx, NK-2, orthodenticle), zincfinger transcription factors (hunchback, snail), basic helix-loop-helix transcription factors (hairy/Enhancer of split, twist), a rel-domain transcription factor (dorsal), secreted signaling molecules (wnt, hedgehog, netrin), a zinc-finger RNA-binding protein (nanos), a phosphatase (cdc25), and a cyclin (cyclin A) and investigations of many other genes are in progress. Several techniques to investigate mRNA and protein expression are now routine in leech studies (RT-PCR, in situ hybridization, immunolocalization, developmental Northern analyses) and techniques for manipulating normal expression of target genes are also becoming available (Nardelli-Haefliger et al. 1994; Pilon and Weisblat 1997; Baker and Macagno 2000; Song et al. 2002). Gene expression during early development. Maternal genes, the products of which are present in leech oocytes and in the early embryo, include Hro-wnt-A, Hro-nos, Lzf2, Le-msx, Hro-twi, cdc25 and cyclin A, and two additional genes, Hro-eve and Hro-hes, are expressed beginning in early cleavage stages, although they may not be maternally deposited (Table 9.1). Hro-nos, lfz2, and Le-msx show a similar early pattern of mRNA localization: transcripts are initially present throughout the oocyte and uncleaved zygote, become segregated to the pools of teloplasm that form prior to first cleavage, and ultimately become localized primarily to the D’ macromere, the precursor of all segmental tissues which inherits the bulk of the teloplasm. In line with these gene-specific findings, earlier studies showed that the majority of polyadenylated RNAs in leech oocytes become concentrated into the teloplasm (Holton et al. 1994). Hro-eve, Hro-hes, cdc25 and cyclin A are expressed throughout the embryo beginning in early cleavage stages. In some or all embryonic lineages they are regulated, or

"" Reproductive Biology and Phylogeny of Annelida

Colour Figure

Fig. 9.10 contd

Hirudinida

"#

become accessible, in a cell-cycle dependent manner suggesting that their expression helps regulate or responds to embryonic cell cycles. Hro-Wnt-A is one of the earliest genes detected in leeches and its protein product, Hro-wnt-A, displays a highly unexpected expression pattern for a species with a fixed cell lineage: stochastic expression between non-equivalent cells (Huang et al. 2001). Despite the many clear differences between cells AB and CD (e.g. in developmental potential, volume, cytoplasmic inheritance, and cell cycle duration), shortly after first cleavage Hro-wnt-A is expressed stochastically in either the AB or the CD cell. Experimental manipulations suggest this gene is involved in signaling from one cell to the other to regulate cell-cell adhesion, and is not involved in cell fate decisions at this stage. Hro-wnt-A is expressed at later stages as well, stochastically at the 4-5 cell stage, during micromere production, and apparently stochastically in provisional epithelium cells (Kostriken and Weisblat 1992; Huang et al. 2001). Hro-wnt-A is expressed in a similar way in micromeres and the provisional epithelium in several glossiphoniid species (Kostriken and Weisblat 1992). Another gene expressed early in leech development, Hro-nos, is a strong candidate for being involved in the cell-fate decision between segmental ectoderm and segmental mesoderm (Pilon and Weisblat 1997). Its protein is expressed primarily in cells which inherit teloplasm and its levels peak when macromere D’ divides to generate the ectodermal and mesodermal precursor cells. At this cleavage, although both daughter cells inherit teloplasm, the transcripts and protein of Hro-nos are preferentially Fig. 9.10 contd

Fig. 9.10 A. Helobdella embryos at the 4-cell stage. Note the large D macromere. Photo courtesy of M. Shankland. B. Helobdella embryo during germinal plate formation silver-stained to highlight superficial cell outlines. This is a ventral view, with anterior up. The provisional epithelium covers much of the embryo at this stage, but the left and right germinal bands and the germinal plate (formed by the coalescence of left/right germinal bands) are apparent as bulges below this epithelium. Courtesy of F. Huang. C. Fluorescently labeled O teloblast and bandlet of a Helobdella embryo. The labeled teloblast has produced a column of approximately 20 primary blast cells since being injected. Courtesy of M. Shankland. D. Late-stage Helobdella embryo (dissected away from yolk) in which two teloblasts (M on left, N on right) were injected with fluorescent tracers early in development. This is a ventral view, with anterior up. The M teloblast gives rise to extensive mesodermal tissues (red) and the N teloblast gives rise primarily to the ventral nerve cord ganglia (green) which lie along the ventral midline. The M and N teloblasts were injected at the same time and produce primary blast cells at approximately the same rate. However, the anterior boundaries of labeled M progeny and labeled N progeny differ because one m primary blast cell produces one M-lineage hemisegmental complement, while two consecutive n primary blast cells are required to produce one N-lineage hemisegmental complement. Courtesy of M. Shankland. E. Late stage Helobdella embryo (dissected away from yolk) with one hemisegmental complement of the o lineage fluorescently labeled (red). This is a ventral view, with anterior up. A single o primary blast cell was injected with tracer four days earlier. Nuclei are labeled with Hoechst 33258 stain (blue). Courtesy of M. Shankland. F. Late stage embryo in which two micromeres, dm’ and c’’’, were injected at an early stage. Anterior is to the left, and dorsal is up. Progeny of dm’ (red) and c’’’ (green) interdigitate to give rise to probable muscle fibers of the proboscis as well as to a temporary fiber network extending throughout the embryo. Courtesy of F. Huang and D. Weisblat.

Helobdella robusta

Helobdella robusta

Helobdella spp.

cdc25

Hro-dl

ht-en

Hro-eve

Hro-hes

Hro-hh

cdc25

dorsal (dl)

engrailed (en)

even-skipped (eve) hairy/Enhancer of split (hes) hedgehog (hh)

Helobdella triserialis

Hirudo medicinalis

Lox7 Lox6

Lox18

Lox1

Hox: labial (lb) Hox: Deformed (Dfd)

Hox: Deformed (Dfd) Hox:? (Scr?Antp?)

Helobdella spp. Helobdella spp. Hirudo medicinalis

Helobdella spp. Theromyzon rude

Helobdella robusta

Helobdella spp.

cyclin A

Helobdella spp.

Leech gene(s) Species

cyclin

Gene family

most (all?) cells in mitosis; neurons of segmental ganglia (mRNA) most (all?) cells in mitosis (mRNA); most (all?) cells in interphase (protein) foregut and midgut; reproductive organs; body wall; neurons of segmental ganglia (mRNA) neurons of segmental ganglia, all segments (mRNA) neurons of segmental ganglia from segment 2 posterior (strongest in segment 3); peripheral nervous system (sensillae and eyes) in Hirudo only (mRNA) longitudinal connectives and lateral nerve roots of all segmental ganglia (mRNA) neurons in all (early) or most (later) segmental ganglia, strongest from segment 3 posterior; nephridia; unidentified body wall cells (mRNA, protein)

all cells, mRNA accessibility invariant in early blastomeres and cell-cycle dependent in teloblasts and probably micromeres (mRNA) all cells, constitutive expression in blastomeres, macromeres, and teloblasts, cell-cycle dependent expression in micromeres all primary blast cells and their progeny; unidentified segmentally iterated stripes of cells (protein) specific cells of young blast cell clones in all teloblast lineages; neurons of segmental ganglia (protein)

Expression (mRNA/protein)

Aisemberg et al. (1993); Aisemberg and Macagno (1994)

Kourakis and Martindale (2001)

Kourakis et al. (1997) Kourakis et al. (1997); Wong and Macagno (1998)

Kang et al. (2003)

Song et al. (2004)

Wedeen and Weisblat (1991); Lans et al. (1993); Ramirez et al. (1995) Song et al. (2002)

Goldstein et al. (2001)

Bissen (1995)

Chen and Bissen (1997)

References

Table 9.1. Developmental regulatory genes investigated in leeches and their primary domains of expression. Where more than one species of Helobdella has been investigated, this is indicated by “spp.”

"$ Reproductive Biology and Phylogeny of Annelida

neurons of segmental ganglia from segment 4 posterior (mRNA). neurons of segmental ganglia from segment 10 posterior (posterior 2/3 of nerve cord); ovaries (segment 10); musculature (dorsoventral flattener muscles in posterior 2/3 of midbody); nephridia; body wall; digestive tract (mRNA, protein) neurons of segmental ganglia strongest from segment 14 posterior (though anterior limit is at segment 7); peripheral tissues from segment 17–25 (mRNA, protein) not expressed? undetectable at any stage (mRNA) oocytes; micromeres; macromeres (especially in teloplasm); throughout germinal bands and germinal plate; micromere cap; unidentified stripes lateral to ganglia; caudal ganglion of rear sucker; rostral region of ventral nerve cord; proboscis sheath; gut (intestine/rectum/anus); subesophageal ganglion; rear sucker (mRNA); one cell stage; micromeres; provisional epithelium; neurons of segmental ganglia; prostomium; intestine (caeca) and rectum (protein) oocyte; D quadrant (primarily in teloplasm); all teloblasts, blast cells, bandlets, and the germinal plate; neurons of segmental ganglia; nephridia (mRNA) midgut, in segmentally iterated spots/stripes in the crop and intestine, and throughout rectum; supraesophageal ganglion (mRNA)

Helobdella spp

Helobdella spp. Hirudo medicinalis

Hirudo medicinalis

Helobdella triserialis Helobdella triserialis

Lox4

Lzf1 Lzf2

Le-msx

Lox10

Hox: Ultrabithorax/ abdominal-A (Ubx/abd-A) hunchback (hb) hunchback (hb)

msx

NK-2

Helobdella triserialis

Helobdella spp.

neurons of segmental ganglia from segment 3–6 (mRNA)

Helobdella spp.

Hox: Sex combs Lox20 reduced (Scr) Hox: Lox5 Antennapedia (Antp) Hox: Lox2 Ultrabithorax/ abdominal-A (Ubx/abd-A)

Nardelli-Haefliger and Shankland (1993)

Master et al. (1996)

Savage and Shankland (1996) Savage and Shankland (1996); Iwasa et al. (2000)

Wong et al. (1995)

Wysocka-Diller et al. (1989); Nardelli-Haefliger and Shankland (1992); Aisemberg et al. (1993); Nardelli-Haefliger et al. (1994)

Kourakis et al. (1997)

Kourakis et al. (1997)

Hirudinida

"%

htr-wnt-A Hro-Wnt-A

wnt

twist (twi)

Hro-sna Hro-sna2 Hro-twi

snail (sna)

Helobdella spp.

Helobdella robusta

Helobdella robusta

Helobdella triserialis Hirudo medicinalis

Lox3 (A/B/C)

ParaHox: Xlox

Hirudo medicinalis

Helobdella triserialis

LNET-1

netrin

Helobdella robusta

orthodenticle (otx) Lox22-Otx

Hro-nos

Leech gene(s) species

nanos (nos)

Gene family

Table 9.1 contd

oocyte; D quadrant (primarily in teloplasm); strong in ectodermal precursor (DNOPQ) and low in mesodermal precursor (DM); primordial germ cells (putative) (mRNA, protein) neurons of segmental ganglia; ventral longitudinal muscles (mRNA, protein) anterior surface ectoderm, eventually encircling developing mouth; foregut; supraesophageal ganglion; neurons of segmental ganglia (mRNA) midgut, in segmentally iterated stripes of segment 6–17 corresponding to crop constrictions and intestinal caeca, and throughout rectum (mRNA) all primary blast cells and their progeny; unidentified segmentally iterated strips of cells (protein) oocyte; throughout development (no spatial distribution information) (mRNA) oocyte; dynamic expression in 2-cell stage, including stochastic expression; micromeres; provisional epithelium (mRNA, protein)

Expression (mRNA/protein)

Kostriken and Weisblat (1992); Huang et al. (2001)

Soto et al. (1997)

Goldstein et al. (2001)

Wysocka-Diller et al. (1995); Wedeen and Shankland (1997)

Gan et al. (1999); Aisemberg et al. (2001) Bruce and Shankland (1998)

Pilon and Weisblat (1997); Kang et al. (2002)

References

"& Reproductive Biology and Phylogeny of Annelida

Hirudinida

"'

segregated to the ectodermal precursor (DNOPQ) relative to the mesodermal precursor (DM). Hro-nos apparently plays no role in anterior/ posterior axis specification, in contrast to the important role the Drosophila homolog nanos plays in this specification (St. Johnston and NüssleinVolhard 1992). Lzf2, Hro-wnt-A, and Hro-nos (Table 1 and S. J. Agee and D. A. Weisblat, unpublished data on Hro-nos) are strongly and transiently expressed in subsets of micromeres shortly after their birth. It seems possible that Hrownt-A plays a role in cell adhesion during micromere production, but the role of Lzf2 and Hro-nos in micromeres at this stage is unknown. Gene expression during mid development. Despite great interest in understanding the molecular basis of leech segmentation, this aspect of leech embryogenesis remains largely obscure. Teloblasts clearly differ in size and developmental potential, and blast cells exhibit lineage-specific stereotypical cleavage patterns, possess lineage-specific developmental potentials, and inherit different axial identities. Nevertheless, nearly all of the characterized genes expressed in segmental precursor cells are expressed broadly and uniformly, with no obvious expression differences between teloblast lineages, blast cell types, or along the anterior/posterior axis. A central puzzle in leech development continues to be how, at the molecular level, differences between teloblasts and between blast cells are established. Genes broadly expressed in developing segmental tissue (i.e., in teloblasts, blast cells, germinal bands, and/or the early germinal plate) include Le-msx, Lzf2 (transcript though not protein), Hro-nos, Hro-eve, Hrohes, Hro-dl, and Hro-sna1/2 (Table 9.1). A homologue of the Drosophila segment polarity gene engrailed was one of the earliest genes characterized in leeches, and remains the only gene known to exhibit clear expression differences between teloblast lineages during germ-band stages of development. The protein of ht-en is expressed transiently in each of the five teloblast lineages, in an iterated, lineagespecific pattern in each blast-cell clone (Wedeen and Weisblat 1991; Lans et al. 1993). Although the ht-en expression pattern initially prompted speculation that it may be involved in establishing segmental boundaries or in compartmentalizing the ventral nerve tissue into ganglia (Wedeen and Weisblat 1991; Lans et al. 1993; Ramirez et al. 1995), further studies including targeted ablations of ht-en expressing cells or their precursors have clearly demonstrated that neither of these hypotheses is correct (Shain et al. 1998; Seaver and Shankland 2000; Seaver and Shankland 2001). Particularly telling, primary blast cell clones can develop normally even when anterior and/or posterior clones are experimentally removed (Seaver and Shankland 2000), indicating that signalling between neighboring clones (by ht-en or any other gene) is not required for normal segment polarity or delineation, in sharp contrast to the method of segment formation in arthropods (Kornberg and Tabata 1993; Davis and Patel 1999). Mid-stage leech embryos also exhibit expression of several genes in developing non-segmental tissue. The protein products of Hro-wnt-A and

"  Reproductive Biology and Phylogeny of Annelida Lzf2 are both expressed in the provisional epithelium, although in quite different patterns: Hro-wnt-A is expressed in an apparently stochastic pattern that varies from embryo to embryo (Kostriken and Weisblat 1992), while Lzf2 is expressed in a bilaterally symmetrical pattern (Iwasa et al. 2000). In the anterior asegmental tissue (prostomium), Lzf2, Lox10, and Lox22-Otx are expressed, each in a unique pattern (Table 9.1). Of note, Lox22-Otx is expressed at the extreme anterior end of the germinal plate, ultimately forming a circle of expression surrounding the developing mouth. Otx homologs are expressed in the developing head of many organisms, including other annelids (Arendt et al. 2001; Bely and Wray 2001), suggesting that this domain of expression in leeches represents a very conserved aspect of animal embryogenesis. Gene expression during late development. In the final stages of embryogenesis, during organogenesis and tissue differentiation, a whole suite of genes is expressed in the leech embryo. Leech neurogenesis is marked by particularly extensive expression of developmental regulatory genes: Ht-en, Hro-eve, Lox22-Otx, Hro-hh, Lzf2, Le-msx, Lnet-1, and all characterized Hox gene homologs are expressed in subsets of ventral nerve cord neurons and peripheral neurons (Table 9.1). Most of these genes are expressed in many contiguous segments or in all segments, in segmentally iterated subsets of neurons that are likely to be (or known to be) segmental neuronal homologs (many of which are identified in Hirudo). These data suggest that specific neuronal phenotypes may be established and/or maintained by a large number of regulatory genes possibly acting in a combinatorial way. In addition to genes expressed neuronally, Lnet-1 is expressed by ventral longitudinal muscles that lie just below the ventral nerve cord, and Lox18 and Lzf2 are expressed in interganglionic longitudinal connective tissue. As a member of the netrin gene family, Lnet-1 is expected to direct axonal growth, and the leech muscle expression likely serves as a guide for peripheral innervation. Expression in the central nervous system midline and expression in muscles appear to be ancient and conserved features of netrin in metazoans (Arendt and Nübler-Jung 1999). As for asegmental neuronal expression, peripheral neurons associated with sensillae and eyes express Lox6 in Hirudo (though not in Helobdella), and neurons of the asegmental, micromere-derived cerebral (supraesophageal) ganglion express Lox22-Otx and Lox10. With two exceptions (Lox18, which is one of two Dfd homologs, and Lox1, of uncertain affinity), all leech Hox genes display nested anterior boundaries of expression relative to one another in the segmental ganglia (reviewed in Kourakis et al. 1997), conforming to the Hox colinearity rule that has been observed in most animals investigated. The labial homolog Lox7 is the most anterior Hox member and is expressed in all segmental ganglia, the Dfd homolog Lox6 is the next most anterior Hox gene and is expressed in all but the anterior-most ganglion, the Scr homolog Lox20 is expressed beginning in the third rostral ganglion, the Antp homolog Lox5 is expressed beginning in the fourth rostral ganglion, and the two Ubx/abd-A

Hirudinida

" 

homologs Lox2 and Lox4 are expressed primarily in the midbody region, in segments 10-25 (somites XII-XXVII) and segments 14-25 (somites XVIXXVII), respectively. Anterior limits of expression tend to be sharp, while posteriorly expression generally fades gradually over multiple segments. All leech Hox genes are expressed considerably after segmental identities are conferred to the segmental ganglia (Martindale and Shankland 1990; Nardelli-Haefliger et al. 1994), and thus cannot be involved in establishing segmental identities, in contrast to the role of Hox genes in Drosophila (McGinnis and Krumlauf 1992). Instead, most leech Hox genes may be involved only in late stages of segmental diversification by helping to establish and maintain different terminal cell fates of neurons. Hox genes in a polychaete (Chaetopterus) are expressed during segment formation and thus could be involved in conferring segmental identity (Irvine and Martindale 2000). Broader sampling within the annelids is needed to determine the ancestral expression patterns and inferred functions of annelid Hox genes. Several genes are also detected in developing mesodermal structures of late-stage embryos (Table 9.1). Lox1, Lox2, and Le-msx are expressed in developing nephridia and Lox20 and Lox5 are expressed in developing segmentally iterated mesodermal structures, such as septa. Dorsoventral flattener muscles (which cause segmental constrictions of the gut) in the posterior two-thirds of the body express Lox2 (the same body segments that express Lox2 in the ventral nerve ganglia), and as mentioned previously ventral longitudinal muscles express LNET-1, presumably to direct peripheral nerve axonal growth. The developing gonads express Lox2 and Hro-hh, and putative primordial germ cells express Hro-nos. Expression of nanos-class genes in germ cells appears to be a widely conserved feature of metazoans (Matova and Cooley 2001). Several genes, including Lzf2, Hro-dl and Hro-sna1/2, are expressed in segmentally iterated domains that may be mesodermal, but the exact origin and ultimate fates of these expressing cells are not known. (Expression in visceral mesoderm is described below, along with expression in the gut lining.) Although a number of genes are expressed in the body wall epidermis, expression tends to be sparse, occurring in only a few, mostly unidentified, cells per segment. A notable exception is Lzf2, for which transcripts, though not protein, are detected specifically in the caudal sucker. The developing gut expresses several genes, either in the true gut lining or in the visceral mesoderm (Table 9.1). The endodermal gut lining initially appears unsegmented, but the crop and intestine (though not the rectum) ultimately develop segmentally iterated bulges and constrictions, the caeca of the crop and intestine. Interestingly, two genes, Lox10 and Lox3, are expressed in the endodermal gut lining in segmentally iterated patterns of spots or stripes that largely prefigure the segmental periodicity and regionalization of the leech gut architecture (Nardelli-Haefliger and Shankland 1993; Wysocka-Diller et al. 1995). The expression of at least one of these (Lox3) is dependent on signals from the tightly apposed visceral

"

Reproductive Biology and Phylogeny of Annelida

mesoderm (Wedeen and Shankland 1997). Both ParaHox and NK-2-class genes are expressed in the developing guts of disparate animals, suggesting that these roles may be widely conserved (Brooke et al. 1998; Ristoratore et al. 1999; Venkatesh et al. 1999). Lzf2 transcripts and protein are also expressed in the intestine and rectum, but in broad, aperiodic patterns. At late embryonic stages Lox22-Otx continues to be expressed in a ring around the developing mouth and also becomes highly transcribed in the developing foregut, specifically in muscles of the proboscis (Bruce and Shankland 1998). Hro-hh is also expressed in proboscis muscles, as well as in all three regions of the midgut (crop, intestine, and rectum), including in rings of segmentally iterated visceral mesoderm (Kang et al. 2003). Experimental disruptions of Hro-hh expression demonstrate that Hro-hh signalling is required for normal gut morphogenesis. Together, molecular studies of Lox3 and Hro-hh reinforce findings from cell ablation studies in implicating endoderm-visceral mesoderm interactions as critical for normal gut morphogenesis. While the number of genes investigated during leech embryogenesis is still relatively limited, at least a few genes involved (or likely to be involved) in many aspects of development have now been identified. Important similarities between leeches and other animal models (Drosophila, vertebrates) have helped to highlight the extreme conservation of the involvement of some of these genes in specific processes: Hox genes in generating axial diversity (especially of the central nervous system), ParaHox genes, NK-2-class genes, and hedgehog-class genes in gut development and regionalization, Otx-class genes in head development, nanos-class genes in germ line development, and wnt-genes in cell adhesion. However, it is clear that the molecular basis of many aspects of leech development differs profoundly from that in other developmental model systems. In sharp contrast to Drosophila, for example, the establishment of the anterior-posterior axis in leeches does not appear to involve nanos or hunchback homologues, establishment of the dorsal-ventral axis and specification of the mesoderm does not appear to involve dorsal or snail homologues, and leech segmentation apparently does not involve homologues of several important Drosophila segmentation genes, including engrailed, even-skipped, hairy/Enhancer of split, hunchback, and hedgehog. Clearly, though much has been learned, leech development still presents many puzzles that remain to be solved.

9.6 LITERATURE CITED Aisemberg, G. O. and Macagno, E. R. 1994. Lox 1, an Antennapedia-class homeobox gene is expressed during leech gangliogenesis in both transient and stable central neurons. Developmental Biology 161: 455–465. Aisemberg, G. O., Kuhn, J. and Macagno, E. R. 2001. Netrin signal is produced in leech embryos by segmentally iterated sets of central neurons and longitudinal muscle cells. Development Genes and Evolution 211: 589–596.

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Aisemberg, G. O., Wysocka-Diller, J., Wong, V. Y. and Macagno, E. R. 1993. Antennapedia-class homeobox genes define diverse neuronal sets in the embryonic CNS of the leech. Journal of Neurobiology 24: 1423–1432. Aisenstadt, T. B., Brodsky, V. J. and Gazarjan, K. G. 1967. An autoradiographic study of the RNA and protein synthesis in gonads of animals with different types of oogenesis. Tsitologiia 9: 397–406. Anderson, D. T. 1973. Embryology and Phylogeny in Annelids and Arthropods, Pergamon Press, Braunschweig. Apakupakul, K., Siddall, M. E. and Burreson, E. M. 1999. Higher-level relationships of leeches (Annelida: Clitellata: Euhirudinea) based on morphology and gene sequences. Molecular Phylogenetics and Evolution 12: 350–359. Apathy, S. V. 1888. Analyse der äusseren Körperform der Hirudineen. Mitteilungen aus der Zoologischen Station zu Neapel 8: 153–232. Arendt, D. and Nübler-Jung, K. 1999. Comparison of early nerve cord development in insects and vertebrates. Development 126: 2309–2325. Arendt, D., Technau, U. and Wittbrodt, J. 2001. Evolution of the bilaterian 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. Autrum, H. 1939. Hirudineen. Geographische Verbreitung. Pp 497–520. In. H. S. Bronns (ed.). Klassen und Ordnungen des Tierreichs, Vol. 4. Section 3, Book 4, No. 2. Akademische Verlagsgesellschaft, Leipzig. Baker, M. W. and Macagno, E. R. 2000. RNAi of the receptor tyrosine phosphatase HmLAR2 in a single cell of an intact leech embryo leads to growth-cone collapse. Current Biology 10: 1071–1074. 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: 2781–2791. Bissen, S. T. 1995. Expression of the cell cycle control gene, cdc25, is constitutive in the segmental founder cells but is cell-cycle-regulated in the micromeres of leech embryos. Development 121: 3035–3043. Bissen, S. T. and Weisblat, D. A. 1989. The durations and compositions of cell cycles in embryos of the leech, Helobdella triserialis. Development 106: 105–118. Blanchard, R. 1896. Viaggio del Dott. A. Borelli nella Republica Argentina e nel Paraguay. 21. Hirudinées. Bollettino del Museo di zoologia dell’Università di Torino 11: 1–24. Blanchard, R. 1917. Monographie des Hémadipsines (Sangsues terrestres). Bulletin de la Société de Pathologie Exotique 10: 640–675. Bonet, S. and Molinas, M. 1988. Ultrastructure of the sperm and spermatogenesis and spermiogenesis of Dina lineata (Hirudinea, Erpobdellidae). Gamete Research 19: 177–190. Borda E. and Siddall, M. E. 2004. Arhynchobdellida (Annelida: Oligochaeta: Hirudinida): Phylogenetic relationships and evolution. Molecular Phylogenetics and Evolution 30: 213–225. Brinkhurst, R. O. 1994. Evolutionary relationships within the Clitellata: An update. Megadrilogica 5: 109–112. Brinkhurst, R. O. and Gelder, S. R., 1989. Did the lumbriculids provide the ancestors of the branchiobdellidans, acanthobdellidans and leeches? Hydrobiologia 180: 7– 15.

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" $ Reproductive Biology and Phylogeny of Annelida Livanow, N. 1931. Die organisation der Hirudineen und die Bezie-hungen dieser Gruppe zu den Oligochäten. Ergebnisse und Fortschritte der Zoologie 7: 378–484. MacCallum, W. G. and MacCallum, G. A. 1918. On the anatomy of Ozobranchus branchiatus (Menzies). Bulletin of the American Museum of Natural History 38: 395–408. Malécha, J. 1975. Étude ultrastructurale de la spermiogenese de Piscicola geometra (Hirudinée, Rhynchobdelle). Journal of Ultrastructural Research 51: 188–203. Mann, K. H. 1962. Leeches (Hirudinea) their structure, physiology, ecology and embryology, Pergamon Press, New York. Martindale, M. Q. and Shankland, M. 1990. Intrinsic segmental identity of segmental founder cells of the leech embryo. Nature 347: 672–674. 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. Matova, N. and Cooley, L. 2001. Comparative aspects of animal oogenesis. Developmental Biology 231: 291–320. McGinnis, W. and Krumlauf, R. 1992. Homeobox genes and axial patterning. Cell 68: 283–302. Moore, J. P. 1924. The anatomy and systematic position of the Chilean terrestrial leech, Cardea valdiviana (Philippi). Proceedings of the Academy of Natural Sciences of Philadelphia 76: 29–48. Moore, J. P. 1946. Leeches (Hirudinea) from the Hawaiian Islands, and two new species from the Pacific region in the Bishop Museum collection. Occasional Papers of Bernice P. Bishop Museum 18: 171–191. 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 in the cephalic nervous system of the leech Helobdella. Development 118: 877–892. 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. 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. Odier, A. 1823. Memoire sur le Branchiobdelle nouveau genre d’Annelides de la famille des Hirudiner. Mémoires de la Societé d’Histoire naturelle, Paris 1: 69–78. Pilon, M. and Weisblat, D. A. 1997. A nanos homolog in leech. Development 124: 1771–1780. Purschke, G., Westheide, W., Rohde, D. and Brinkhurst, R. O. 1993. Morphological reinvestigation and phylogenetic relationship of Acanthobdella peledina (Annelida, Clitellata). Zoomorphology (Berlin) 113: 91–101. Raj, P. J. S. and Penner, L. R. 1962. Concerning Ozobranchus branchiatus (Menzies, 1791) (Piscicolidae: Hirudinea) from Florida and Sarawak. Transactions of the American Microscopical Society 81: 364–371. Ramirez, F. A., Wedeen, C. J., Stuart, D. K., Lans, D. and Weisblat, D. A. 1995. Identification of a neurogenic sublineage required for CNS segmentation in an annelid. Development 121: 2091–2097.

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" & Reproductive Biology and Phylogeny of Annelida Siddall, M. E., Apakupakul K., Burreson, E. M., Coates, K. A., Erseus, C., Gelder, S. R., Kallersjö, M. and Trapido-Rosenthal, H., 2001. Validating Livanow: Molecular data agrees that leeches, branchiobdellidans, and Acanthobdella peledina form a monophyletic group of oligochaetes. Molecular Phylogenetics and Evolution 21: 346–251. Siddall, M. E. and Burreson, E. M. 1995. Phylogeny of the Euhirudinea: Independent evolution of blood feeding by leeches? Canadian Journal of Zoology 73: 1048– 1064. Siddall, M. E. and Burreson, E. M. 1996. Leeches (Oligochaeta?: Euhirudinea), their phylogeny and the evolution of life history strategies. Hydrobiologia 334: 277– 285. Siddall, M. E. and Burreson, E. M. 1998. Phylogeny of leeches (Hirudinea) based on mitochondrial cytochrome c oxidase subunit I. Molecular Phylogenetics and Evolution 9: 156–162. Siddall, M. E. 2002. Phylogeny of the leech family Erpobdellidae (Hirudinida: Oligochaeta). Inverterbrate Systematics 16: 1–6. Siddall, M. E. and Borda, E. 2003. Phylogeny and revision of the leech genus Helobdella (Glossiphoniidae) based on mitochondrial gene sequences and morphological data and a special consideration of the triserialis complex. Zoologica Scripta 32: 23–33. Smith, C. M. and Weisblat, D. A. 1994. Micromere fate maps in leech embryos— lineage-specific differences in rates of cell proliferation. Development 120: 3427– 3438. Smith, C. M., Lans, D. and Weisblat, D. A. 1996. Cellular mechanisms of epiboly in leech embryos. Development 122: 1885–1894. 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: 3681–3692. 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 69: 183–195 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. St. Johnston, D. and Nüsslein-Volhard, C. 1992. The origin of pattern and polarity in the Drosophila embryo. Cell 68: 201–219. Trontelj, P., Sket, B. and Steinbruck, G. 1999. Molecular phylogeny of leeches: congruence of nuclear and mitochondrial rDNA data sets and the origin of bloodsucking. Journal of Zoological Systematics and Evolutionary Research 37: 141–147. Venkatesh, T. V., Holland, N. D., Holland, L. Z., Su, M. -T. and Bodmer, R. 1999. Sequence and developmental expression of amphioxus AmphiNK2-1: insights into the evolutionary origin of the vertebrate thyroid gland and forebrain. Development, Genes and Evolution 209: 254–259. Wedeen, C. J. and Shankland, M. 1997. Mesoderm is required for the formation of a segmented endodermal cell layer in the leech Helobdella. Developmental Biology 191: 202–214. 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: 805–814. Weisblat, D. A. and Blair, S. S. 1984. Developmental indeterminacy in embryos of the leech Helobdella triserialis. Developmental Biology 101: 326–335.

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10

CHAPTER

Phyllodocida Fredrik Pleijel1 and Greg W. Rouse2

10.1 PHYLOGENY AND SYSTEMATICS There are two studies providing cladistic analyses of Phyllodocida, Rouse and Fauchald (1997) and Pleijel and Dahlgren (1998), both based on morphological data. There are also other studies that include only a very limited number of Phyllodocida, but these will not be further discussed here (see Rouse and Pleijel 2001). Although both resolution and terminals within Phyllodocida differ between the results of these two studies, the inclusiveness of the group is identical in both, and both also have Eunicida as sister group. Phyllodocida and Eunicida, together with Amphinomida, make up the major clade Aciculata, by and large corresponding to what was once upon time known as ‘Errantia’. One main difference between the two studies relates to the position of Pisionidae. Following Rouse and Fauchald (1997), they are situated within a large grade including e.g., chrysopetalids, syllids, nereidids, phyllodocids, nephtyids and glyceriforms, but according to Pleijel and Dahlgren (1998) they have the more traditionally (e.g. Åkesson 1961) favored position as sister to the scaleworms; the relationships of pisionids certainly warrant further studies. Another difference involves chrysopetalids, nereidids and hesionids, which in Rouse and Fauchald (1997) appear within the previously mentioned grade, but instead constitute a clade in Pleijel and Dahlgren (1998). This latter relationship was also previously suggested by Glasby (1993), as well as in a combined morphological and molecular analysis by Dahlgren et al. (2000). Based on these earlier studies, Rouse and Pleijel (2001) recognized the two major clades Aphroditiformia (scaleworms and Pisionidae) and Nereidiformia (Chrysopetalidae, Hesionidae, Nautiliniellidae, Nereididae,

1

Department of Marine Ecology, Tjärnö Marine Biological Laboratory, Göteborg University, SE-452 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 2 South Australian Museum Nth Terrace, Adelaide, SA 5000, and School of Earth and Environmental Sciences, University of Adelaide SA 5005, Australia

"!

Reproductive Biology and Phylogeny of Annelida

Pilargidae, Syllidae, plus Microphthalmus and Hesionides) within Phyllodocida. In addition they referred a number of taxa to ‘Phyllodocida unplaced’, namely Glyceriformia, Ichthyotomus, Lacydonia, Lopadorhynchidae, a group of ‘minor’ holopelagic Phyllodocida, Myzostomida, Nephtyidae, Paralacydonia, Phyllodocidae, Sphaerodoridae, and Tomopteridae. As for proposed phylogenies of less inclusive taxa within Phyllodocida, there are now a number of studies available. One study by Dahlgren and Pleijel (1995) included a small analysis of Chrysopetalidae. Hesionidae were revised by Pleijel (1998) (see also Pleijel and Rouse 2000), and within that group there are also smaller analyses carried out on Heteropodarke (Pleijel 1999) and on the relationships between Amphiduros and Gyptis (Pleijel 2001b). On nereidids there are two studies treating the whole group (Fitzhugh 1987; Glasby 1991) that both may be regarded as more preliminary, and one more extensive study on namanereidins (Glasby 1999), the latter including also a full revision of the group. The position of pilargids was investigated by Licher and Westheide (1994); however, the result that pilargids actually belong within hesionids has not obtained support in subsequent studies (Pleijel and Dahlgren 1998; Dahlgren et al. 2000). There are several recent studies on the phylogeny of syllids; one by Nygren (1999) which treats the whole group, but based on a limited number of taxa, and two much more extensive ones on autolytins (Nygren and Sundberg 2003; Nygren 2004). Glyceridae were recently analyzed and fully revised by Böggemann (2002). A series of analyses has been conducted on benthic Phyllodocidae by Pleijel (1991), Eibye-Jacobsen (1993), Orrhage and Eibye-Jacobsen (1998), and Kato and Pleijel (2003), with divergent results, especially regarding the more basal relationships and the monophyly of Eteoninae and Phyllodocinae; clearly the basal relationships among the phyllodocids are in need of further investigations. Analyses of subgroups of phyllodocids include revisions of Phyllodoce (Pleijel 1993), of Notophyllum (Kato and Pleijel 2002), and of Paranaitis (Kato and Pleijel 2003), and a more preliminary study on the phylogeny of Alciopini was published by Wu and Lu (1993). Somewhat surprisingly, to date there are no cladistic studies performed on scaleworms. Myzostomids are a story of their own and one that is still developing. Myzostomida, all obligate symbionts, mainly with crinoids but some with other echinoderms, comprises a small taxon of around 150 nominal species. There is an ongoing debate on the overall position of the group, with some authors favoring a position among polychaetes (Rouse and Pleijel 2001), while others favor a placement outside annelids and closer to platyhelminths (Eeckhaut et al. 2000) or rotifers (Zrzavy et al. 2001). Here we follow Rouse and Pleijel (2001) in placing them within Phyllodocida but further investigation is certainly required. With the exception of Dahlgren et al. (2000), Nygren and Sundberg (2003), Nygren and some of the Myzostomida analyses, the phylogenetic studies mentioned above are all based on morphological characters only, and since these tend to be limited in number there is an obvious lack of

Phyllodocida

"!!

stability in the trees; even minor changes of characters and taxa will tend to affect the results, sometimes in drastic ways. Furthermore, there is the issue with the unresolved root position of Phyllodocida. Although the monophyly of this group seems reasonably well supported, some authors (Storch 1968; Conway Morris and Peel 1995; McHugh 1997; Westheide 1997) have, implicitly or explicitly, suggested that it may constitute a basal group on the annelid tree, and that it therefore is not monophyletic (see Rouse and Pleijel 2001; 2003 for further references and discussion of these issues).

10.2 ANATOMY WITH REFERENCE TO THE REPRODUCTIVE SYSTEM Most of the body in annelids is segmental. In many cases some of the anterior segments may become associated with the head. Often the remainder of the body segments are basically similar in dimension and components. This is seen in most taxa of Phyllodocida, although a few anterior segments may become cephalized. All Phyllodocida have a spacious coelomic cavity. While a complete, closed circulatory system is present in most annelids, a limited circulatory system occurs in quite a number of Phyllodicida (e.g., Phyllodocidae, Glyceridae). In these groups, some of the major blood vessels are present but the distal capillary vessels are missing (see Rouse and Pleijel 2001 and references within). The effects of this in terms of gametogenesis have yet to be fully investigated, though it has have been discussed in terms of segmental organs (see below).

10.2.1 Epitoky Many benthic polychaetes swarm to the surface regions of the water and spawn. This widespread method is accomplished by epitoky (morphological modification that enables the animal to leave the bottom to reproduce). Epitokes can arise by two different processes. Epigamous epitokes are the result of the transformation of a whole individual into the epitoke (Fig. 10.1A). Examples occur across Phyllodocida including Glyceridae, Nephtyidae, Nereididae, Phyllodocidae and Syllidae (Clark 1961). Schizogamous epitoky arises by modification and separation from the posterior end of the worm. It is further separated into two forms. Scissiparity is where existing segments are transformed into stolons and this is known in Eunicida as well as in Phyllodocida such as Syllidae. The form of scissiparity known as gemmiparity is unique to syllids. This is where stolons are produced from newly produced segments, specialised for the purpose (see e.g. Garwood 1991; Franke 1999; Nygren 1999 and references within). The complex physiological changes associated with epitoky are reviewed by Schroeder and Hermans (1975) and Durchon (1984). Most work has been done on the epigamous epitoky found in Nereididae. Here there are major changes to the eyes, parapodia and the circulatory system and musculature. Wissoq (1978; 1977) described the changes in the longitudinal muscle fibers during the transformation of

"!" Reproductive Biology and Phylogeny of Annelida

Fig. 10.1 A –C. Laubierpholoe swedmarki from Carrie Bow Cay, Belize. A. Transmission electron micrograph of longitudinal section through late spermatid. B. Light microscope micrograph of spermatids apparently developing in tetrads. C. Light microscope micrograph of sperm. Same scale as B. Note the median insertion of flagella. D. Lepidonotus sp. from Heron Island, Great Barrier Reef. Transmission electron micrograph of longitudinal section through mature sperm. E. Psammolyce sp. from Carrie Bow Cay, Belize. Transmission electron micrograph of longitudinal section through mature sperm. Abbreviations: a, acrosome; d, distal centriole; f, nuclear fossa; m, mitochondrion; n, nucleus; p, proximal centriole. Original.

Phyllodocida

"!#

the benthic form to the epitoke where there is histolysis of the longitudinal and circular muscles and re-differentiation of longitudinal muscles where glycogen and the number of mitochondria in the muscle vastly increases. New ‘heteronereis’ muscles in the parapodia are also grown with masses of mitochondria and glycogen granules and these muscles are mainly responsible for the frantic swimming activity the epitokes are capable of.

10.2.2 Segmental Organs In most annelids there are excretory ducts to the exterior referred to as nephridia. Other ducts, known as gonoducts or coelomoducts, are also required for the transfer of gametes that develop in the coelom to the outside of the body on maturity. The two different kinds of ducts will be referred to as segmental organs here since determining what kind of duct is present is problematic, especially in Phyllodocida. Major studies by Goodrich (1895; 1945) on segmental organs are still influential, but are now being supplanted (reviewed by Bartolomaeus 1999). The basis of Goodrich’s (1945) theory and classification of annelid segmental organs rested on the hypothesis that mesodermal coelomoducts and ectodermal (or ectomesodermal) nephridia could fuse to form single structures. He proposed that the ancestral condition in polychaetes was a pair of excretory nephridia in each segment as well as a pair of gonoducts. In annelid taxa such as Clitellata and Capitellidae this condition exists, but in virtually all other polychaetes there is at most a single pair of ducts in a given segment. Therefore, Goodrich proposed that in the majority of polychaetes the nephridia and coelomoducts had fused to varying degrees to give a single pair of ducts per segment. Nephridia can have the coelomic end closed (= protonephridia), or have an open funnel (= metanephridia) termed a nephrostome. Coelomoducts can have funnels (coelomostomes) and gonoducts to the exterior but are often combined with nephridia. Goodrich (1945) defined such structures as nephromixia and classified them into three groups; protonephromixia, metanephromixia and mixonephridia. Protonephromixia have mesodermal components grafted onto a canal of the protonephridium and are found in Phyllodocida such as Glyceridae and Phyllodocidae. Metanephromixia was a term used when there was clear morphological evidence that mesodermal funnels have been added to open nephrostomes and were only found in segments with gametes. Goodrich stated that taxa in Phyllodocida such as Chrysopetalidae, Hesionidae and Syllidae had these organs. Mixonephridia was a term used for when the coelomic ends of the nephridia were completely fused with coelomostomes and were found in all segments irrespective of reproductive status. Goodrich (1945) described these as being present in most polychaetes outside Phyllodocida, though he regarded scale worms as having mixonephridia as well. However, the line between metanephromixia and mixonephridia is difficult to draw. Lately, Goodrich’s work has been subject to some criticism

"!$ Reproductive Biology and Phylogeny of Annelida (Bartolomaeus 1999) and his hypothesis and naming system seems to be inadequate. Bartolomaeus (1999) proposed that in polychaetes in general, the nephridia act as gonoducts as well as excretory organs. Segmental organs therefore represent structures that arise from a single tissue source and are not the result of fusion of two kinds of ducts. In Phyllodocida such as Pholoe (Pholoidae), Bartolomaeus has also shown that what Goodrich (1945) called metanephromixia do not arise from a mixture of two kinds of tissue, but develop from a single source. Also, Goodrich’s protonephromixia are in fact protonephridia that can develop a funnel coming off the side of the duct, and this is a reversible process. Bartolomaeus (1999) argues that Goodrich based his interpretation of two tissue sources for the segmental organs of many polychaetes on misleading staining of histological sections, not on ontological evidence. There is some evidence to support Goodrich’s hypothesis of segmental origins originating from two tissue sources and these are reviewed in Rouse and Fauchald (1997). It may be worthwhile to maintain Goodrich’s terms protonephromixia to distinguish them, from protonephridia, but justifying the use of metanephromixia and mixonephridia needs much more work. The distribution of segmental organs in annelids varies, but a common pattern is for segmental organs to be present along most of the body, and to be used for both excretion and gamete emission. This is the case across most of Phyllodocida and the major differences among taxa occur with reference to the occurrence of protonephridia or metanephridial based segmental organs (see Rouse and Fauchald 1997; Rouse and Pleijel 2001). Protonephromixia are found in Glyceridae, Goniadidae, Nephtyidae, Phyllodocidae, Pisionidae, Tomopteridae and Typhloscolecidae, metanephridia are present in Nereididae without apparent association with gamete discharge, which seems to be via body wall rupture (Goodrich 1945). Other Phyllodocida excrete and discharge gametes through what Goodrich (1945) would term metanephromixia or mixonephridia and Bartolomaeus (1999) would call metanephridia.

10.3

OOGENESIS

A general description of oogenesis in polychaetes is given by Eckelbarger in Chapter 2. Here we provide an entry into the literature on oogenesis in Phyllodocida in tabular format (Table 10.1), together with further details from case studies in the different subgroups. The more detailed studies on oogenesis in scaleworms are those on Harmothoe imbricata by Garwood (1981) and on Pholoe baltica by Heffernan and Keegan (1988b). The study on P. baltica contains both quantitative and qualitative data on the maturation of oocytes. The development is intraovarian, and it is unusual compared to other Phyllodocida in that oocytes with follicle cells remain in clusters joined by intercellular bridges throughout the major part of the oogenesis. A similar development is

Phyllodocida

"!%

present also in the only studied sphaerodorid, Sphaerodorum gracilis (Christie 1984). A female of S. gracilis with eggs is illustrated in Fig. 10.9A. In H. imbricata, the ovaries are associated to segmental blood vessels oogenesis, and the development is also intraovarian (sensu Eckelbarger 1988). The eggs are released into the coelom when reaching maturity at a size of 125–140 µm (Garwood 1981). Among hesionids there are three studies on oogenesis of which two on Psamathe fusca (Olive and Pillai 1983a, 1983b; as Kefersteinia cirrata). Ovaries are present in all segments from segment 11, as finger-like lobes situated around an axial blood vessel. Oogenesis is intraovarian and each oocyte is associated with a follicle. The oogenesis is well synchronized and influenced both by temperature and day-length. Gametogenesis for Sirsoe methanicola was described by Eckelbarger et al. (2001; as Hesiocaeca methanicola). It has intraovarian oogenesis taking place on the outside of blood vessels associated to the body wall and the oocytes are completely covered by follicle cells. Ovarian eggs are up to c. 60 µm in diameter, and coelomic ones c. 70 µm, meaning some postovarian growth does occur. The spawned eggs are about 80 µm in diameter. There is a large amount of information present on egg development in nereidids, and only some examples will be mentioned here: a comprehensive list is provided in Table 10.1 and molecular developmental studies are covered in Chapter 5. Fischer (1974; 1975) described the oogenesis in Platynereis dumerilii. Discrete gonads are absent and oogenesis extraovarian, as seems to be the case in all studied nereidids (e.g. Eckelbarger 1984), and the actual site of proliferation of the early oocytes remains unknown (Fischer 1984). Early coelomatic oocytes are connected by intercellular bridges and covered by sheath cells. In association with vitellogenesis, the bridges and the sheath cells are lost and the oocytes break apart and further development is solitary in the coelom. Whereas younger females contain oocytes that are highly heterogenous, they become uniform in size in association with maturity, and virtually all are fully mature at spawning. Nevertheless, how the actual final growth and maturation of the oocytes is controlled and timed to the somatic changes associated with the epitoky remains poorly understood (Fischer 1996). Pfannenstiel et al. (1987) described the gametogenesis in the two sibling species P. dumerilii and P. massiliensis and, in terms of eggs, the two differ in the size of mature oocytes, which are 160– 180 µm in diameter in the former, and 280–300 µm in the latter. They differ also in amount of yolk and in number of eggs; at the onset of spawning the P. dumerilii females are completly filled with oocytes, whereas a P. massiliensis female sheds only about 300 eggs. Another newly discovered sibling species complex in nereidids with (among other reproductive features) different egg sizes is represented by Hediste atoka, H. diadroma and H. japonica (Sato 1999; Sato and Nakashima 2003), where H. atoka has mature oocytes that measures 200–250 µm in diameter, H. diadroma 130–170 µm, and H. japonica 180–210 µm.

Chrysopetalum debile

Chrysopetalidae

Glyceriformia

planktotrophic (Rouse 2000); all larval stages (Cazaux 1968) juvenile (Bhaud and Cazaux 1987) planktotrophic (Pernet et al. 2001; Rouse 2000); early larvae (Blake 1975) juvenile (Bhaud and Cazaux 1987) metatrochophore (Kisseleva 1992)

Hermione hystrix

Aphroditidae

Glycera convoluta planktotrophic; all larval stages (Cazaux 1967)

Glycera capitata

Strepternos didymopyton Viktoriella zaikai

Dysponetus pygmaeus Paleanotus bellis

lecithotrophic (Drasche 1885; Rouse 2000)

Panthalis oerstedi

Acoetidae

development

species

major taxon

oogenesis

Table 10.1 Summary of reproductive features in Phyllodocida

ect-aquasperm (Franzén 1956)

sperm

epitoky

parental care

swarming (Berkeley and Berkeley 1948) swarming (Gravier and Dantan 1928; Lo Bianco 1909)

mating

"!& Reproductive Biology and Phylogeny of Annelida

nectochaete (Blake 1975)

planktotrophic; trochophore (Simpson 1962a)

Glycinde armigera lecithotrophic?; all larval stages (Blake 1975) Glycinde juveniles polygnatha (Blake 1975) Goniada emerita lecithotrophic?; all larval stages (Cazaux 1972; Rouse 2000) Goniada maculata late larva (Plate and Husemann 1994) Goniadella late larva (Plate bobretzky and Husemann 1994)

Glycera tesselata

Glycera oxycephala Glycera rouxii

Glycera lapidum

Glycera dibranchiata

ect-aquasperm (Franzén 1956)

ect-aquasperm (Simpson 1962a)

limited (Simpson 1962b)

none (Simpson 1962b)

Table 10.1 contd

swarming (Fage and Legendre 1927) swarming (Gravier and Dantan 1928)

swarming (Arwidsson 1899; Fage and Legendre 1927; Gravier and Dantan 1928; StøpBowitz 1941)

swarming (Simpson 1962b)

Phyllodocida

"!'

capricornia

Hesionidae

lecithotrophic? (Pleijel and Rouse 2000) Gyptis late larvae brevipalpa1 (Blake 1975) Gyptis ?brunnea late larvae and juveniles (Blake 1975) Gyptis late larvae propinqua (Bhaud 1971) Gyptis rosea planktotrophic; all larval stages and juveniles (Haaland and Schram 1982) Hesionid A (Blake 1975) Lizardia hirschi lecithotrophic? (Pleijel and Rouse 2005) Nereimyra punctata planktotrophic; all larval stages and juveniles (Banse 1956; Rasmussen 1973; Schram and Haaland 1984)

Hemipodia simplex late larvae (Blake 1975)

Glyceriformia

development

species

major taxon

Table 10.1 contd

oogenesis

copulation (Pleijel and Rouse 2005)

mating

introsperm (Pleijel and Rouse 2005)

parental care

copulation (Pleijel and Rouse 2000)

epitoky

introsperm (Pleijel and Rouse 2000)

sperm

"" Reproductive Biology and Phylogeny of Annelida

planktotrophic; all ect-aquasperm larval stages and (Franzén 1956) juveniles (Haaland and Schram 1983; Plate and Husemann 1994) lecithotrophic; all larval stages (Blake 1975; Strathmann 1987) planktotrophic; intraovarian embryology, (Eckelbarger et al. trochophore 2001) (Eckelbarger et al. 2001; Treadwell 1901) intraovarian (Eckelbarger et al. 2001; Olive and Pillai 1983a)

Sirsoe methanicola planktotrophic?; intraovarian ect-aquasperm none early stages (Eckelbarger et al. (Eckelbarger et al. (Eckelbarger et al. (Eckelbarger et al. 2001) 2001) 2001) 2001) Syllidia armata all larval stages (Casanova 1954; Rasmussen 1956)

Sinohesione genitaliphora

Psamathe fusca

Ophiodromus obscurus

Ophiodromus pugettensis

Ophiodromus flexuosus

introsperm (Westheide et al. 1994)

Table 10.1 contd

copulation (Westheide et al. 1994) broadcast spawner (Eckelbarger et al. 2001)

swarming (Strathmann 1987)

Phyllodocida

""

species

Nephtyidae

Myzostomida

development

late larva (Lacalli 1980)

two early stages (Lacalli 1980)

Aglaophamus neotenus

Nephtys caeca intraovarian (Eckelbarger 1984)

oogenesis

later larval stages (Thorson 1946) Nephtys hombergi all larval stages intraovarian (Rasmussen 1973; (Bentley et al. Wilson 1936) 1984) Nephtys incisa all larval stages (Lacalli 1980)

Nephtys ciliata

(Kato 1952)

(Eeckhaut and Jangoux 1993)

planktotrophic; all larval stages (Kleinenberg 1886; Meyer 1901; Åkesson 1967) late larva (Mileikovsky 1972)

Myzostoma sp.

Myzostoma sp.

Myzostoma cirriferum

Pelagobia longocirrata

Lopadorhynchidae Lopadorhynchus spp.

major taxon

Table 10.1 contd

introsperm (Afzelius 1983 1984) introsperm (Mattei and Marchand 1988)

sperm

none (external) (Bentley et al. 1984)

epitoky

none (Bentley et al. 1984)

parental care

broadcast spawner (Bentley et al. 1984) broadcast spawner (Bentley et al. 1984)

(Kato 1952)

(Eeckhaut and Jangoux 1993)

mating

"" Reproductive Biology and Phylogeny of Annelida

Nereididae

extraovarian (Eckelbarger 1984; Klesch 1970; Mazurkiewicz 1975)

extraovarian (Dales 1950; Dhainaut 1969 1984; Olive and Garwood 1981) all larval stages (Sato 1999; Sato (Sato and Tsuchiya and Tsuchiya 1991) 1991)

all larval stages (Cazaux 1969; Dales 1950)

Laeonereis culveri (Mazurkiewicz 1975)

Hediste japonica

Hediste diversicolor

Hediste diadroma

(Durchon 1956)

Ceratonereis costae Hediste atoka

(Sato 1999; Sato and Tsuchiya 1991) all larval stages (Sato 1999; Sato (Sato and Tsuchiya and Tsuchiya 1991) 1991)

late larva (Plate and Husemann 1994)

Nephtys sp.

Nephtys sp.

ect-aquasperm (Sato and Osanai 1986; Takashima and Takashima 1963)

ect-aquasperm limitted epigamy (Bertout 1976; (Dales 1950) Olive and Garwood 1981)

ect-aquasperm? (Rouse 1999)

limited swarming (Herpin 1925)

Table 10.1 contd

swarming (e.g. Sato and Nakashima 2003; Sato and Tsuchiya 1987) pseudocopulation (Dales 1950; Rasmussen 1973 and references within) swarming (Sato and Nakashima 2003)

Phyllodocida

""!

major taxon

Table 10.1 contd

oogenesis

3-chaetiger larva (Feuerborn 1931) lecithotrophic; all larval stages (Herpin 1925; Reish 1957) lecithotrophic; all larval stages (Gilpin-Brown 1959) all larval stages (Cazaux 1969; Herpin 1925)

(Rullier 1954)

development

ect-aquasperm (Defretin and Wissocq 1974)

sperm

epitoky

epigamy (Herpin 1925 and references within)

epigamy (Charrier 1920; Fage 1904)

epigamy (Herpin 1925 and references within)

swarming: egg mass (Herpin 1925 and references within)

parental care

Neanthes succinea late larva (Plate and Husemann 1994; Rasmussen 1973) Neanthes virens lecithotrophic; all ect-aquasperm ect-aquasperm limited epigamy none (Bass and larval stages (Bass and Brafield (Bass and Brafield (Bass and Brafield Brafield 1972) (Banse 1954; Bass 1972) 1972) 1972) and Brafield 1972)

Neanthes irrorata

Neanthes fucata

Micronereis variegata Namalycastis hawaiiensis Neanthes caudata

Micronereis nanaimoensis

Leptonereis glauca

species

swarming (Bass and Brafield 1972; Rasmussen 1973)

swarming (Rasmussen 1973)

swarming (Herpin 1925 and references within) copulation or pseudocopulation (Berkeley and Berkeley 1953; Paxton 1983)

mating

""" Reproductive Biology and Phylogeny of Annelida

(Reish 1954)

extraovarian (Eckelbarger 1984; Schroeder 1971) ect-aquasperm (Fallon and Austin 1967)

(Reish 1954)

viviparous; all (Smith 1950) larval stages (Smith 1950) Nereis pelagica all larval stages extraovarian ect-aquasperm epigamy (e.g. (Herpin 1925; (Dhainaut 1984; (Defretin and Herpin 1925 and Lacalli 1980; Eckelbarger 1984) Wissocq 1974) references within) Olive and Garwood 1981; Plate and Husemann 1994; Rasmussen 1956; Wilson 1932) Nereis procera all larval stages epigamy (Strathmann 1987) Nereis vexillosa all larval stages (Johnson 1943) epigamy (Johnson 1943; (Johnson 1943) Pernet et al. 2001; Strathmann 1987) Perinereis ect-aquasperm brevicirrus (Kubo and Sawada 1977; Sawada 1984) Perinereis cultrifera all larval stages extraovarian epigamy (e.g. (Cazaux 1969; (Dhainaut 1984; Herpin 1925; Herpin 1925) Herpin 1921; Rouabah and Rouabah and Scaps 2003) Scaps 2003)

Nereis limnicola

Nereis limbata

Nereis grubei

Table 10.1 contd

swarming (Johnson 1943)

egg mass (Johnson 1943)

swarming; egg mass (Gravier 1923; Herpin 1925)

swarming (e.g. Herpin 1925)

swarming (Reish 1954)

intracoelomic development (Smith 1950) egg mass (e.g. Herpin 1925)

none

Phyllodocida

""#

Nereidiformia incertae sedis

Nereididae

major taxon

Table 10.1 contd

development

Hesionides arenaria

Tylorrhynchus heterochaetus

Platynereis massiliensis

Platynereis dumerilii

Platynereis bicanaliculata

several stages (Westheide 1967)

lecithotrophic (putatively planktotrophic nectochaete); all larval stages and juveniles (Blake 1975) all larval stages (Cazaux 1969; Herpin 1925)

Perinereis marionii all larval stages (Herpin 1925)

species

extraovarian (Fischer and Dorresteijn 2004 and references within; Pfannenstiel et al. 1987) (Pfannenstiel et al. 1987)

extraovarian (Eckelbarger 1984; Herpin 1921)

oogenesis

parental care

mating

introsperm no (Westheide 1984b) (Westheide 1967)

ent-aqua- or no introsperm? (e.g. Pfannenstiel (Pfannenstiel et al. et al. 1987) 1987) ect-aquasperm (Sato and Osanai 1983, 1990) eggs deposited in sediment (Westheide 1967)

no

hypodermic injection (Westheide 1967)

swarming (Fischer and Dorresteijn 2004 and references within)

epigamy (Herpin swarming; egg mass 1925 and (Herpin 1925 and references within) references within) epigamy swarming (Strathmann 1987) (Strathmann 1987)

epitoky

ect-aquasperm epigamy (Fischer (Pfannenstiel et al. and Dorresteijn 1987) 2004 and references within)

sperm

""$ Reproductive Biology and Phylogeny of Annelida

Paralacydonia paradoxa

Laubierpholoe swedmarki Pholoe baltica

Pholoidae

Microphthalmus similis

Microphthalmus sczelkowii

Microphthalmus nahantensis

Microphthalmus listensis

planktotrophic; all larval stages (Christie 1982; Heffernan and Keegan 1988a)

(Heffernan and Keegan 1988b)

probably lecithotrophic (Rouse 2000); several larval stages (Bhaud 1967)

several stages (Rasmussen 1956; Westheide 1967)

all larval stages (Westheide and Rieger 1987) (Westheide and Rieger 1987)

Microphthalmus carolinensis

Paralacydonia

Nereidiformia incertae sedis

trochophore (Westheide 1967)

Microphthalmus aberrans

ect-aquasperm? (Heffernan and Keegan 1988b)

introsperm (Westheide and Rieger 1987) introsperm (Westheide and Rieger 1987) introsperm (Westheide and Rieger 1987) introsperm (Westheide 1967)

introsperm (Westheide 1967)

no (Westheide 1967)

no (Westheide 1967)

no (Westheide 1967)

no (Westheide 1967)

brooding in elytrae (Laubier 1975)

eggs deposited in sediment (Westheide 1967)

cocoon (Westheide 1967)

cocoon (Westheide and Rieger 1987)

cocoon (Westheide and Rieger 1987)

cocoon (Westheide 1967)

Table 10.1 contd

copulation (Westheide 1984a)

copulation (Westheide 1967)

copulation (Westheide 1967)

hypodermic injection (Westheide 1967)

Phyllodocida

""%

Phyllodocidae

major taxon

Table 10.1 contd

development

oogenesis

Alciopa sp.

Alciopa reynaudii

Alciopa parasitica

juvenile (Buchholz 1869)

(Eckelbarger and Rice 1988)

Pholoe cf inornata late larva (Plate and Husemann 1994) Pholoe minuta planktotrophic? all larval stages (Lacalli 1980; Thorson 1946) Pholoe pallida lecithotrophic (Christie 1982) Pholoe lecithotrophic; synopthalmica all larval stages (Cazaux 1968) Pholoides asperus (Blake 1975) (Blake 1975) Taylorpholoe sp.

species

ent-aquasperm (Rice 1987; Rice and Eckelbarger 1989) ent-aquasperm (Rice 1987; Rice and Eckelbarger 1989)

introsperm? (Rouse 1999)

ent-aquasperm? (Christie 1982)

sperm

epitoky

mating

embryos in body cavity, later brooding under elytra (Wolf 1984; Pettibone 1992)

parental care

""& Reproductive Biology and Phylogeny of Annelida

late larvae (Plate and Husemann 1994; Thorson 1946) later stages and juveniles (Blake 1975) all larval stages (Lacalli 1980; Plate and Husemann 1994; Rasmussen 1956; Thorson 1946) all larval stages (Cazaux 1985)

ect-aquasperm? (Rouse 1988) Eulalia viridis all larval stages extraovarian ect-aquasperm (Meyer 1938; Nolte (Eckelbarger 1984; (Olive 1975) 1938; Olive 1975; Olive 1975) Plate and Husemann 1994) Eumida sanguinea all larval stages (Cazaux 1969; Plate and Husemann 1994; Thorson 1946) Krohnia lepidota ent-aquasperm (Rice 1987; Rice and Eckelbarger 1989)

Eulalia sp

Eteone picta

Eteone longa

Eteone dilatae

Eteone barbata

none (e.g. Olive 1975)

mucous bag

Table 10.1 contd

Phyllodocida

""'

major taxon

Table 10.1 contd

development

oogenesis

planktotrophic?; all larval stages (Thorson 1946) Phyllodoce all stages (Bhaud laminosa and Cazaux 1987) Phyllodoce lineata several larval stages (Bhaud 1967) Phyllodoce early stages maculata (Lacalli 1980) Phyllodoce all larval stages mucosa (Cazaux 1969; Lacalli 1980; Plate and Husemann 1994) Phyllodoce rosea late larva (Plate and Husemann 1994) Phyllodoce sp. late stages (Blake 1975) Phyllodoce planktotrophic; williamsi all larval stages and juveniles (Blake 1975)

Notophyllum foliosum Phyllodoce groenlandica

species Naiades cantrainii

sperm ent-aquasperm (Franzén 1956; Rice 1987; Rice and Eckelbarger 1989)

epitoky

egg mass (Blake 1975)

mucous bag (Cazaux 1969; Lacalli 1980; Sach 1975)

mucous bag

brooding (Okada 1930)2 mucous bag (Thorson 1946)

parental care

pseudo-copulation in groups (Sach 1975)

mating

"# Reproductive Biology and Phylogeny of Annelida

Vanadis formosa

Torrea candida

Sige fusigera

Rhynchonerella moebii

Rhynchonerella angelini

Pseudomystides limbata

Plotohelmis tenuis

late larva (Plate and Husemann 1994)

late larval stages (Bhaud 1967; Plate and Husemann 1994) (Eckelbarger and Rice 1988)

ent-aquasperm? (Franzén 1956; Rice 1987; Rice and Eckelbarger 1989) ent-aquasperm (Rice 1987; Rice and Eckelbarger 1989)

ent-aquasperm (Rice 1987; Rice and Eckelbarger 1989) ent-aquasperm (Rice 1987; Rice and Eckelbarger 1989)

ent-aquasperm (Rice 1987; Rice and Eckelbarger 1989)

Table 10.1 contd

Phyllodocida

"#

planktotrophic; late larval stages (Plate and Husemann 1994; Åkesson 1961)

Pisione remota

Polynoidae

Pisionidae

late larval stage (Bhaud 1973) late larva (Blake 1975) late larvae (Britayev 1981)

Ancistrosyllis groenlandica Ancistrosyllis sp.

Pilargidae

Arctonoe vittata

planktotrophic; trochophore (Britayev 1991; Pernet et al. 2001; Phillips and Pernet 1996)

Acholoe astericola all larval stages (Cazaux 1968; Davenport 1954) Alentia gelatinosa

Pisionidens indica trochophore and juvenile (Aiyar and Alikunhi 1940)

Pisione spp.

Cabira cf bohajensis

development

species

major taxon

Table 10.1 contd

oogenesis

(Franzén and Rice 1988)

introsperm? (Aiyar and Alikunhi 1940)

introsperm (Westheide 1988)

sperm

none (Stecher 1968)

epitoky

none (Stecher 1968)

parental care

copulation (Yamanishi 1992, 1998) copulation (Aiyar and Alikunhi 1940)

copulation (Stecher 1968)

mating

"# Reproductive Biology and Phylogeny of Annelida

Harmothoe longisetis

Harmothoe impar

Harmothoe imbricata

Halosydna gelatinosa Halosydna johnsoni

Halosydna brevisetosa

Gattyana cirrosa

Enipo kinbergi

(Rossi 1976 in Phillips and Pernet 1996) planktotrophic; all intraovarian ent-aquasperm brood eggs under larval stages and (Eckelbarger 1984; (Bentley and elytrae (e.g. Bentley juveniles (Blake Garwood 1981) Serries 1992; Daly and Serries 1992) 1975; Cazaux 1972) 1968; Daly 1972; Holborow 1971; Holborow and Laverack 1969; Korn 1958, 1959; Lacalli 1980; Rasmussen 1956; Thorson 1946) all larval stages (Korn 1958, 1959; Rasmussen 1956) all larval stages ect-aquasperm (Cazaux 1968) (Bentley and Serries 1992)

late larval stage (Thorson 1946) all larval stages and juveniles (Blake 1975) (Cazaux 1968)

ect-aquasperm (Franzén 1956)

Table 10.1 contd

pair formation (Bentley and Serries 1992; Daly 1972)

Phyllodocida

"#!

Sigalionidae

Polynoidae

major taxon

Table 10.1 contd

development

Sigalion bandaensis

Neoleanira tetragona Psammolyce sp.

Lepidonotus (Simon 1965) sublevis Polynoe antarctica

Harmothoe lunulata all larval stages (Cazaux 1968) Harmothoe sarsi several stages (Korn 1958) Lagisca extenuata all larval stages (Cazaux 1968) Lepidonotosus all larval stages clava (Cazaux 1968) Lepidonotosus planktotrophic; squamatus all larval stages (Cazaux 1968; Lacalli 1980; Rasmussen 1973) Lepidonotus sp. early stages (Mead 1897) Lepidonotus sp.

species

oogenesis

ect-aquasperm (Franzén 1956) ect-aquasperm? (Rouse 1999) (Jamieson and Rouse 1989)

ect-aquasperm? (Rouse 1988) (Simon 1965)

ect-aquasperm (Franzén 1956)

sperm

epitoky

mating

brood eggs under elytrae (Fauvel 1916)

parental care

"#" Reproductive Biology and Phylogeny of Annelida

Amblyosyllis cincinnata Amblyosyllis speciosa

Syllidae

Calamyzas amphictenicola

Bollandia antipathicola Brania pusilla

Autolytus varians

Autolytus roseus

Autolytus maculata

Sphaerodorum

Sphaerodoridae

late larval stages (Malaquin 1893)

lecithotrophic; all larval stages (Pernet 1998)

lecithotrophic? (Christie 1984)

planktotrophic?; all larval stages (Cazaux 1968; Plate and Husemann 1994) Sthenelais limicola late larva (Plate and Husemann 1994)

Sthenelais boa

(Christie 1984)

introsperm? (Franzén and Rice 1988)

ent-aquasperm? (Pernet 1998)

ect-aquasperm (Christie 1984)

schizogamy (Potts 1913) schizogamy (Potts 1913) schizogamy (Mensch 1900; Potts 1913) atokous (Glasby 1994)

egg mass

atokous (Riser 1982)

none

Table 10.1 contd

Phyllodocida

"##

major taxon

Table 10.1 contd

development

Haplosyllis spongicola Myrianida edwarsi3 (Malaquin 1893)

Haplosyllis hamata

Epigamia ?magna3 late larva (Strathmann 1987) Eusyllis assimilis (Malaquin 1893) Eusyllis blomstrandi Exogone mediterranea Exogone naidina (Gravier 1923; Rasmussen 1973 and references within)

Epigamia alexandriý3

Dentatisyllis mangalis

species

extraovarian (Eckelbarger 1984; Gidholm 1963)

oogenesis

epitoky

ent-aquasperm? (Franzén 1956)

schizogamy (Potts 1913) schizogamy (Okada 1937) schizogamy (Gidholm 1963, 1965, 1967; Nygren 2004; Okada 1937; Potts 1913)

epigamy (Herpin 1925)

all larval stages schizogami (Okada 1930; Qian (Hamond 1969; and Chia 1989) Qian and Chia 1989)

sperm

swarming (Herpin 1925)

swarming (Hamond 1969; Qian and Chia 1989)

mating

viviparous (San Martín 1984) attached juveniles swarming? (Gravier 1923; (Rasmussen 1973) Rasmussen 1973 and references within)

brooding pouches (Qian and Chia 1989)

viviparous (Russell 1995)

parental care

"#$ Reproductive Biology and Phylogeny of Annelida

schizogamy (Okada 1937) schizogamy (Potts 1913; Rasmussen 1973)

atokous (Bührmann et al. 1996a)

Proceraea aurantiaca Proceraea cornuta

introsperm (Bührmann et al. 1996a)

epigamy (Herpin 1925)

all larval stages (Cazaux 1969)

attached embryos (Potts 1913)

epigamy (Galloway and Welch 1911; Wolken and Florida 1984) epigamy (Daly 1975)

schizogamy (Potts 1913) lecithotrophic; later schizogamy stages (Dales 1951) (Gidholm 1965 Rasmussen 1973f) ect-aquasperm? (Franzén 1982a)

Pionosyllis lamelligera Pionosyllis pulligera

Odontosyllis polycera Parapionosyllis gestans Petitia amphophthalma

Odontosyllis later stages ctenostoma (Herpin 1925) Odontosyllis enopla

Myrianida sp.

Myrianida prolifera3 (Malaquin 1893)

Myrianida pinnigera (Malaquin 1893)

Table 10.1 contd

copulation (Bührmann et al. 1996a) swarming (Herpin 1925)

Phyllodocida

"#%

major taxon

Table 10.1 contd

development

all larval stages (Herpin 1925)

Syllis amica

Syllis corruscans

Syllis armillaris

late larval stages (Cazaux 1972)

Sphaerosyllis hystrix Sphaerosyllis tetralix Syllides edentula

Salvatoria limbata5 all larval stages and juveniles (Cazaux 1972) Sphaerosyllis hermaphrodita

Salvatoria clavata4

Procerastea halleziana

Proceraea picta

species

oogenesis

ect-aquasperm (Franzén 1956)

introsperm (Kuper and Westheide 1997a)

ent-aquasperm (Franzén 1956, 1974)

sperm

schizogamy (Potts 1913)

schizogamy (Herpin 1925)

schizogamy (Okada 1937) schizogamy (Okada 1937; Potts 1913)

epitoky

(Cognetti-Varriale 1971)

dorsal incubation (Cazaux 1972; Potts 1913

parental care

(Cognetti-Varriale 1971) swarming (Herpin 1925)

copulation (Kuper and Westheide 1997b)

mating

"#& Reproductive Biology and Phylogeny of Annelida

Trypanosyllis asterobia Trypanosyllis coeliaca

Trypanosyllis aeolis

Syllis vivipara

Syllis variegata

Syllis sp.

Syllis ramosa

Syllis pulchra

Syllis prolifera

Syllis krohnii

Syllis hyalina

ect-aquasperm (Jamieson and Rouse 1989)

extraovarian ect-aquaperm (Eckelbarger 1984; (Heacox and Heacox and Schroeder 1981a) Schroeder 1981b)

planktotrophic; all larval stages and juveniles (Cazaux 1969, 1984)

lecithotrophic; all larval stages (Heacox 1980)

(Malaquin 1893)

schizogamy (Potts 1913) schizogamy (Okada 1937) schizogamy (Potts 1913)

schizogamy (Cazaux 1984)

schizogamy (Okada 1937; Potts 1913)

schizogamy (Potts 1913) schizogamy (Potts 1913) schizogamy (Okada 1937) schizogamy (Heacox 1980)

viviparous (Goodrich 1900)

none (Heacox 1980)

Table 10.1 contd

swarming (Heacox 1980)

Phyllodocida

"#'

Tomopteris helgolandica

Trypanosyllis crosslandi Trypanosyllis zebra

species

lecithotrophic; all larval stages (Plate and Husemann 1994; Åkesson 1962)

development

sperm

extraovarian ent-aquasperm?; (Eckelbarger 1984; biflagellate Åkesson 1962) (Franzén 1982b)

oogenesis

2

Now referred to Podarkeopsis, although Blake’s description actually seems to refer to a member of Gyptis. Kato and Pleijel (2002) suggested that the reported egg mass may belong to a parasitic copepod. 3 Previously referred to Autolytus (Nygren 2004). 4 Previously referred to Grubea, Brania, Grubeosyllis and Pseudobrania (San Martin 2003). 5 Previously referred to Grubea, Brania and Pseudobrania (San Martin 2003).

1

Tomopteridae

major taxon

Table 10.1 contd

schizogamy (Potts 1913) schizogamy (Okada 1937)

epitoky

parental care

mating

"$ Reproductive Biology and Phylogeny of Annelida

Phyllodocida

"$

Studied phyllodocids include the benthic Eulalia viridis (Olive 1975) and the pelagic alciopins Alciopa reynaudii and Rhynchonerella angelini Eckelbarger and Rice (1988). Discrete ovaries are absent in all three taxa. In Eulalia viridis oogonia and oocytes occur in clusters. The oocytes remain together till vitellogenesis takes place, and there are no follicle cells. At this stage the eggs obtain their characteristic emerald green colour. The mature oocytes are disc-shaped and measure 110–120 µm in diameter. Egg development in Alciopa reynaudii and Rhynchonerella angelini is very similar to E. viridis, also having extraovarian oogenesis with oogonia released into the coelom as clusters, although they differ in the presence of follicle cells. The oogonia are connected by intercellular bridges. Solitary oocytes are then released from the clusters where the vitellogenesis takes place. Oogenesis in Syllis pulchra (as Typosyllis) was described by Heacox and Schroeder (1981b), and this appears to be the only available syllid study. The early development takes place in peritoneal ovaries, whereafter the eggs are released as small oocytes. These are associated to sibling cells that may function as nurse cells. Final maturation occurs in the swimming stolon. As pointed out in Chapter 2, it appears difficult to discern clear phylogenetic patterns from our current knowledge on ovarian morphology and oogenesis in annelids, and this is no less true for Phyllodocida.

10.4 SPERM ULTRASTRUCTURE Scaleworm sperm studied in TEM to date include Alentia by Franzén and Rice (1988), Enipo kinbergi by Franzén (1956), Harmothoe imbricata, H. impar and H. longisetis by Bentley and Serries (1992), Lepidonotus by Rouse (1988), Psammolyce (Fig. 10.1E) and Taylorpholoe by (Rouse 1999), and Sigalion by Jamieson and Rouse (1989). We here also present the unusual sperm of the pholoid Laubierpholoe swedmarki (Fig. 10.1A–C) where females brood larvae inside the elytrae (Fig. 10.8) and so fertilization is almost certainly internal. With the exception of L. swedmarki, H. imbricata and Taylorpholoe, all of these have typical ect-aquasperm without obviously distinguishing features, although one difference is that Lepidonotus have a nuclear cone (Fig. 10.1D), which appears to be absent from both Alentia and Sigalion (unknown for Harmothoe). Harmothoe imbricata, which reproduce by pairing and also broods the fertilized eggs under the elytra (e.g. Daly 1972), instead has mature sperm with an elongated head region and a pointed acrosome, and a ring of up to 14 mitochondria surrounding the centrally positioned axoneme (Daly 1974; Bentley and Serries 1992). Taylorpholoe sp. also brood larvae under the elytrae (Pettibone 1992) and some form of sperm transfer and internal fertilization must also occur (see section 10.5). Among pisionids, sperm ultrastructure of Pisione remota was investigated by Westheide (1988). The sperm is aflagellate, and only part of the spermatogenesis takes place within the male, to be continued within the

"$

Reproductive Biology and Phylogeny of Annelida

female spermathecae after the copulation. The mature sperm within the male is rod-shaped with an elongated nucleus and vacuole-like structure, and with a number of mitochondria situated at both ends of the cell. Within the spermathecae the cells go through a series of changes, involving phagocytosis of the vacuole, enlargement of the nucleus, and development of pseudopodia which join to receptacle cells. Westheide (1988) suggested that these latter modifications are related to long-term storage of sperm within the females. There are three ultrastructural sperm studies on hesionids. In the smallsized capricornia the males have well-developed penes on segment 9, and the females a pair of ventral oviducts on segment 11. The spermatids develop in large clusters before splitting up at a late spermatid stage. The mature sperm has a cap-like acrosome, a cylindrical nucleus, and three mitochondria surrounding the base of the nucleus and the two centrioles (Fig. 10.2A; see also Pleijel and Rouse 2000). The likewise small-sized Lizardia hirschi has elongated sperm with a conical acrosome, and extended nucleus and three mitochondria (Pleijel and Rouse 2005). Both these taxa reproduce by copulation, as seen by the presence of penes in the males and spermathecae in the females, and fertilization is likely to be internal. Other examined hesionids include members of Gyptis, Micropodarke, Ophiodromus, Podarkeopsis and Syllidia, all provided with ect-aquasperm (pers. obs.). Eckelbarger et al. (2001) described the sperm of the cold-seep hesionid Sirsoe methanicola (as Hesiospina methanicola); it is a broadcast spawner with ectaquasperm with an elongated conical tip containing a cup-shaped acrosome, a rounded nucleus, six to eight mitochondria, and two centrioles

Fig. 10.2 A. Transmission electron micrograph of transverse section through mature endosperm of capricornia from One Tree Island, Great Barrier Reef. B. Transmission electron micrograph of transverse section through mature ext-aquasperm of Nephtys sp. from Papua New Guinea. Abbreviations: a, acrosome; d, distal centriole; m, mitochondrion; n, nucleus. Original.

Phyllodocida

"$!

associated to the 9+2 axoneme. Early stages develop as syncytial masses of sperm morulae connected to blood vessels in testes. They are connected by cytoplasmatic bridges and have a synchronized development. The spermatids leave the testes and enter the coelom, where they occur as tetrads. A number of other ultrastructural studies has been conducted on spermiogenesis, sperm and spermatophores of myzostomids, though all have been restricted to Myzostoma (e.g., Afzelius 1984; Mattei and Marchand 1988). Spermiogenesis occurs within spermiocysts, with all spermatids in a given spermiocyst at the same stage of development. Spermiogenesis is complicated compared to 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 assumed that the electron dense spheres are chromatin (e.g., Afzelius 1983; Eeckhaut and Jangoux 1991) and this is 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. The sperm usually swims (in seawater at least) with the flagellum foremost (Afzelius 1983). This form of locomotion is not particularly effective but 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). To our knowledge, there exists but one single illustration of nephtyid sperm in the literature (Rouse 1999); sperm from the same species are shown in Fig. 10.2B. As might be expected from what is known about nephtyid reproduction (e.g. Bentley et al. 1984) it has a commonly shaped ect-aquasperm with a

"$" Reproductive Biology and Phylogeny of Annelida spherical nucleus, a hemispherical acrosome, a single midpiece, and four mitochondria surrounding the two centrioles. All nereidids examined to date have ect-aquasperm, with the exception of Platynereis massiliensis, which has, either introsperm or, possibly, entaquasperm (Pfannenstiel et al. 1987; Jamieson and Rouse 1989). Nereidid spermatogenesis follows a common pattern with spermatogonia and spermatocytes occurring in large clusters which are joined by cytoplamatic bridges (Bertout 1976; Kubo and Sawada 1977; Sawada 1984). The spermatogonia are first associated with the peritoneum, but subsequently appear in the coelomic fluid. The clusters break up and give rise to spermatid tetrads. One putative apomorphy for Nereididae (or a subclade of Nereididae; see below) is the presence of an acrosomal rod in mature sperm, which extends from the acrosomal vesicle into the nucleus. It has been shown in a series of taxa belonging to Hediste (Bertout 1976; Sato 1999), Neanthes (Takashima and Takashima 1963; Bass and Brafield 1972; Sato and Osanai 1986), Nereis (Fallon and Austin 1967; Defretin and Wissocq 1974), Perinereis (Kubo and Sawada 1977), and Platynereis (Pfannenstiel et al. 1987), but is absent from Tylorrhynchus (Sato and Osanai 1983). Possibly acrosomal rods are visible also in Retzius’ (1909) sperm illustrations of Nereis pelagica and Hediste diversicolor. The acrosomal rod has a perforatorial role in the fertilization, and serves to bring the inner acrosomal membrane in contact with the egg envelope (Sato and Osanai 1986). The absence of acrosomal rods in Tylorrhynchus does not necessarily imply a homoplastic distribution, since this group may be one of the basal-most nereidid lineages (Santos et al in press). Similar structures have been reported also from a few other polychaetes, including a sabellariid (Pasteels 1965) and an opheliid (Ochii et al. 1977), although these occurrences may warrant further investigation. To date, there have been no studies of pilargid sperm. We here present an electronmicrograph of Sigambra sp. from Japan, showing what appears to be a typical ect-aquasperm but nothing is known about reproduction in this group (Fig. 10.3A–C). Syllids exhibit a large amount of specialisation in their reproduction, and, as expected, sperm morphology also shows a lot of variation. Bührmann et al. (1996a; 1996b) studied the reproductive system and the ultrastructure of the sperm of the eusyllin Petitia amphophthalma. Although most eusyllins reproduce by epigamy (e.g., Garwood 1991; Nygren 1999), the interstitial Petitia ampophthalma lacks epitokous modifications. The mature sperm, which is stored in a seminal vesicle extending over a number segments, is filiform with a very elongated cap-shaped acrosome, and a likewise elongated nucleus which is posteriorly excavated where the basal part of the axoneme is positioned. There are two long mitochondria surrounding the basal part of the axoneme. It is unusual in having four rather than two centrioles in the primary and secondary spermatocytes, although the distribution of this feature in other taxa requires further study. The exogonin syllid Sphaerosyllis hermaphrodita also has unusual sperm that

Phyllodocida

"$#

Fig. 10.3 A–C. Transmission electron micrographs of the pilargid Synelmis sp. from Seto, Japan. A. longitudinal section. B. Detail of acrosome. C. Detail of midpiece. Abbreviations: a, acrosome; d, distal centriole; f, nuclear fossa; m, mitochondrion; n, nucleus; p, proximal centriole. Original.

is filiform and with a helically coiled acrosome (Kuper and Westheide 1997a). Among autolytins, Franzén (1982a) studied an Autolytus, member of the ‘prolifer-group’ (that group is now referred to Myrianida), which reproduce by gemmiparity (e.g., Nygren and Sundberg 2003; Nygren 2004). The anterior region of the mature sperm is oval elongated, the acrosome is laterally displaced to the anterior side of the nucleus, and there is no posterior excavation in the nucleus. The posterior part of the nucleus is surrounded by one or a few mitochondria. There are two centrioles, situated perpendicular to each other, and from the one associated to the axoneme there are three (?) striated rootlets that project anteriorly along the sides of the nucleus. Light microscopy studies of several Autolytus species were also carried out by Gidholm (1965). Syllis pulchra, similar to many autolytins, reproduces by means of stolons, but differs in that there seems to be no obvious mating behaviour between the stolons (Heacox and Schroeder 1981a). The mature sperm has a cylindrical head region, with an external incision separating the acrosome region from the nucleus. The acrosome is rounded with a distinct subacrosomal space, and the nucleus is barrel-shaped with five to six mitochondria at the base; the axoneme has the usual 9+2 pattern of microtubuli. The epigamous exogonin Salvatoria clavata (previously Brania) has a highly characteristic acrosome, shaped like a beaker with the opening directed posteriorly, and an elongated, cylindrical nucleus (Franzén 1974). The middle piece contains two centrioles and a single mitochondrion, orientated perpendicular to each

"$$ Reproductive Biology and Phylogeny of Annelida other. In the aberrant ectoparasitic Calamyzas amphictenicola (Åke Franzén, pers. com. in Jamieson and Rouse 1989), which possibly is hermaphroditic, the sperm is spherical, there is a disc-shaped nucleus dislocated to one pole, and a varying number of mitochondria, but a flagellum is lacking. However, the syllid affinity of Calamyzas remains questionable. Westheide (1984b) carried out TEM studies on sperm and spermatogenesis of the interstitial Hesionides arenaria. Males have a pair of penes, situated on either side of the median antenna on the prostomium, and transfer spermatophores from these to the females by hypodermic injection (Westheide 1982, 1984b). Mature sperm is long and thread-like with pointed ends. The acrosome has the shape of an extended cone, and the nucleus that of a very elongated cap. The basal body of the axoneme is situated inside the nuclear cap, just below its anterior end. Centrioles were not been observed. The axoneme has the standard 9+2 microtubular structure, and extends posteriorly through the entire length of the sperm. In the middlepiece the axonome is provided with nine double supporting walls and a mitochondrial complex that form a circular sheath. Three species of the likewise interstitial but hermaphroditic Microphthalmus (not necessarily closely related to Hesionides), M. carolinens, M. listensis and M. nahantensis, are also reported to have filiform sperm (Westheide 1984a; Westheide and Rieger 1987). In benthic phyllodocids, only one member of Eulalia is known in some detail (Rouse 1988). It has a typical ect-aquasperm morphology, with a caplike acrosome, elliptical nucleus, four mitochondria, two centrioles, and an axonem with a 9+2 pattern of the microtubuli (Fig. 10.4A). The spermatids develop in clusters (Fig. 10.4B). It is unusual in that the mitochondria are ensheathed by a common membrane. Alciopini is a group of holopelagic polychaetes which is likely to have a sister group situated somewhere within the phyllodocids (e.g. Pleijel 2001a), probably near Eulalia, Eumida and Pterocirrus. Alciopin sperm varies from more typical ectaquasperm shape to elongated ones. Rice (1987) provided information on external morphology for a series of Alciopini sperms, and Rice and Eckelbarger (1989) studied ultrastructure in Vanadis formosa and Krohnia lepidota. Both the latter are similar to each other with minor differences only, with the mature sperm having a caplike acrosome, an elongated nucleus, a short middlepiece with 5–7 mitochondria in the former and 8–10 in the latter, and two centrioles. The nucleus of V. formosa is unusual in having an irregular posterior part, and it contrasts to the grand majority of Phyllodocida in being hermaphrodite (Rice and Eckelbarger 1989). Rice and Eckelbarger (1989) suggested that an elongated shape may be associated with sperm storage, known to occur in several alciopids (Eckelbarger and Rice 1988; Rice 1987), including V. formosa and, possibly, K. lepidota. However, Franzén and Rice (1988), in their review of polychaete sperms, provided TEM illustrations of Naiades cantranii and Torrea candida, both of which have sperm storage, but also have more typical ect-aquasperm morphologies.

Phyllodocida

"

%$Fig. 10.4 Transmission electron micrographs of Eulalia sp. from Queensland, Australia. A. Longitudinal section through mature sperm. B. Spermatids developing in clusters, surrounded by mature sperm. Abbreviations: m, mitochondrion; n, nucleus. G. Rouse.

The holopelagic tomopterids not only hold an enigmatic position within Phyllodocida, but Tomopteris helgolandica also has highly unusual sperm, as seen from Franzén (1982b). The sperm is characterized by the presence of two flagella, by lacking a distinct acrosome, and by an axoneme microtubular pattern of 9+0, rather than the common 9+2. However, there is no further information on the reproductive modes of tomopterids.

10.5 MATING AND FERTILIZATION Aphroditiformia. There is remarkable little information on mating in scaleworms. All known members are gonochoric, with the exception of Macellicephala violacea that was reported to be hermaphroditic by Wirén (1907). For taxa that lack external genital organ and have ect-aquasperm an external fertilization is expected. However, a number of deep-sea polynoids are provided with ventral segmental papillae on a limited number of segments—whether or not these are involved in reproduction deserves further investigation. The best studied scaleworm member is the polytelic Harmothoe imbricata (Daly 1972; Daly et al. 1972), which form pairs with the male positioned on the dorsum of the female. The female discharges the eggs through a limited number of swollen segmental organs, and then males spawn, probably by the prolonged nephridial papillae that occurs only in males. The sperm are ent-aquasperm. Chrysopetalidae. For chrysopetalids we have no actual information on breeding, although one species, Dysponetus bipapillatus, may have external paired genital organs on one segment (Dahlgren 1996).

"$& Reproductive Biology and Phylogeny of Annelida Glyceridae. There are many reports of swarming in glycerids (see Table 10.1). Epitoky involves changes in body musculature, atrophy of the gut and modifications of the parapodia. For Glycera dibranchiata Simpson (1962b) provides detailed information, see also Böggemann (2002) for references on this and other glycerid taxa. The animals swim head first, with lateral undulating of the body, similar to nereidids and different from the atokous swimming in glycerids. Shedding of gametes takes place through the posterior end, probably through ruptures, and the animals die afterwards. Hesionidae. There is no direct evidence for mating procedures in hesionids, although from sperm ultrastructure and from the lack of genital organs in most taxa one might expect broadcast spawning or pseudocopulation. The majority are gonochoric, although at least some members of Hesione are hermaphrodites (Bergmann 1903, pers. obs.). Several of the recently described smaller taxa, however, differ in having external genital organs. Sinohesione genitaliphora, capricornia and Lizardia hirschi all are provided with penes, in the two former on median segments and in the last on the pygidium or the last segment (Westheide et al. 1994; Pleijel and Rouse 2000, 2005). Both latter taxa also have introsperm, indicating copulation. Myzostomida. In myzostomes there are five pairs of protonephridia that are separate from the gonoducts (Pietsch and Westheide 1987). The pairs of nephridia lie anterior to each pair of parapodia. A pair of ciliated ducts that usually connects the uterus to the intestine has commonly been referred to as metanephridia. They are thought to serve an excretory function in removing waste gametes but may not actually represent metanephridia (Grygier 2000). Myzostomida are usually protandric hermaphrodites and pass through a functional male stage to be simultaneous hermaphrodites at maturity (Grygier 2000) though some taxa pass from being males to females (Wheeler 1896). The male system is paired with the testes, usually diffuse, lying ventral to the gut. Seminal vesicles are connected to the testes via vasa deferentia and exit on each side of the body next to the third parapodia. A pair of protrusible penes is present in ectocommensitic myzostomes, but absent in all endoparasitic forms. Spermatophores are placed on the surface of recipient worms and the spermiocysts pass into the body of the worm before migrating to the uterus (Grygier 2000). The female system has one or two ovaries dorsal to the gut that lead into uterine diverticula that fuse into a median uterus that exits, via an oviduct, near the anus. A pair of ciliated ducts (referred to above as possible metanephridia) also leads from the diverticula to the intestine. Fertilization is internal and probably occurs in the uterus. Nephtyidae. In nephtyids the segmental organs are closed to the exterior and cannot be used for discharge of gametes (Goodrich 1945). For two studied polytelic broadcast spawners, Nephtys caeca and N. hombergi, Bentley et al. (1984) showed that both male and female gametes are emitted via a posterior opening between the coelom and the gut, and subsequently

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through the anus. Bentley et al. (1984) suggested that this may be a general feature in nephtyids. While spawning the animals remain in the sediment with the posteriormost part projecting above the surface. Nereididae. As far as known, most nereidids are gonochoric and discharge the gametes by irreversible rupturing of the body wall. There is, however, a lot of variation present in the group. Members of Micronereis undergo some changes in association with maturity, including enlargement of the eyes, development of accessory cirri on the parapodia, and development in males of copulatory hooks on the third chaetiger that are used to hold the females during the coupling (Paxton 1983 and references within). Many nereidids, especially members of Nereis, Neanthes and Platynereis, are well-known for their epigamous epitoky and swarming behaviour where they discharge eggs and sperm during a ‘nuptial dance’. This has been described in detail, for e.g. Platynereis dumerilii (see Fischer and Dorresteijn 2004 and references within). The maturing males and females metamorphose into a ‘heteonereis’ stages (Fig. 10.5A), including a profound reorganization that includes enlargement of eyes, reorganization of musculature and blood vessels, and from segment 16 (males) or 22 (females) the parapodia are modified and the chaetae are exchanged for paddle-like ones. The animals are mature during one single day when they swim at night-time searching for partners. The mating or the ‘nuptial dance’ takes place by high-speed swimming in circles while they emit sperm and eggs in close vicinity of each other. The pheromones involved in the onset of gamete discharge were identified by Hardege (1999). Fertilisation takes place in the water, and the spent adults die soon after the mating. Other nereidids do not have these drastic metamorphoses, and, for example, in Hediste diversicolor, the only external visible difference is that both males and females change from brownish to a green colour. Mature females have been observed swimming at night, and laboratory observations indicate that they may be mating with their tails knot to each other (Herpin 1925). The actual fertilization was described in several ultrastructural studies by Sato and Onsai (1983, 1986, 1990) on the Japanese palolo, Tylorrhynchus heterochaetus. Other nereidiforms. Mating in the interstitial Nereidiformia Microphthalmus and Hesionides (previously referred to Hesionidae; see Pleijel and Dahlgren 1998) exhibits some variation. All Microphthalmus are hermaphroditic with the anterior segments being differentiated as males and the posterior segments as females. Mating in M. aberrans is by hypodermic injection with spermatophores (Westheide 1967), whereas in M. similis, M. szelkowii and M. arenarius there is true copulation (Westheide 1967, 1973, 1984a). The male organs in M. carolinensis were described in detail in Westheide (1978). Members of Hesionides are gonochoric, and mature males are provided with paired penes flanking the median antenna on the prostomium. Mating in H. arenaria takes place by hypodermic injection with spermatophores (Westheide 1967).

"% Reproductive Biology and Phylogeny of Annelida

Colour Figure

Fig. 10.5 A. Heteronereis of nereidid from the Swedish west-coast. B. Myrianida pachycera from Sydney, Australia with stolons. G. Rouse.

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Phyllodocidae. There are few direct observations on phyllodocid reproduction. Sach (1975) reported breeding Phyllodoce mucosa, with large number of animals appearing on intertidal flats, forming reproductive groups with a single female and many males. A mucous bag is secreted by the female, and eggs and sperm are deposited within this. Known benthic phyllodocids lack external reproductive organs, and the few sperm studies all indicate ect-aquasperm (Olive 1975; Rouse 1988). Reproduction in these taxa at least may be limited to pseudocopulation as in P. mucosa, or to broadcast spawning. In many alciopins, in contrast, the females have receptacles and sperm storage (Rice 1987; Eckelbarger and Rice 1988; Rice and Eckelbarger 1989), indicating that copulation likely takes place (although sperm storage is known to occur in some other polychaetes without copulation, such as in sabellids (Rouse 1996). Pisionidae. All known Pisionidae are gonochoric. Mating, achieved by copulation, is remarkable, as seen from Stecher’s (1968) study on Pisione remota. The male organs are external paired structures that arise from the ventral part of the parapodia on a large number of median segments, and the female organs consist of paired genital papillae and spermatheace on a number of segments. The actual copulation therefore involves multiple coupling between a large number of segmental genital organs. For further details on reproduction in Pisione are provided by Schroeder and Hermans (1975) and Yamanishi (1991, 1998), and by Hartmann-Schröder (1970) on Pisionidens. Syllidae. Most syllids are gonochoric and have epitoky, and both epigamy and schizogamy occur. Schizogamy in syllids occurs into two forms; normal scissiparity where existing segments are transformed into stolons, and gemmiparity, unique to syllids, where stolons are produced from newly produced segments, specialised for the purpose (see e.g. Garwood 1991; Franke 1999; Nygren 1999 and references within). Myrianida pachycera, a shizogamous autolytin is illustrated in Fig. 10.5B. The stolons in schizogamous taxa may look similar for both sexes, or males and females may be different. Epigamy is present in many Exogoninae and Eusyllinae. As an example, Daly (1975) described the reproduction in Odontosyllis polycera, where epigamous modifications involve eye enlargement, luminescence, external opening of the segmental organs, development of compound long-bladed notochaetae, and shift from crawling to swimming in the open water where the mating takes place. The animals survive the swarming and returns to a benthic live. Details on various epigamous modifications in Odontosyllis spp. can be found in Galloway and Welch (1911), Goodrich (1933), Verger-Bocquet (1983), and Wolken and Florida (1984). Both epi- and schizogamy exist in Autolytinae (Nygren 2004). Myrianida prolifera and M. edwarsi (previously Autolytus; see Nygren 2004) provides well-known examples of gemmiparous schizogamy (e.g. Dales 1951; Gidholm 1965; see Nygren 2004 for later references). Male and female stolons have different morphologies and the males swim at much higher speed than the females. A male autolytin stolon is shown in Fig. 10.10B. The

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Reproductive Biology and Phylogeny of Annelida

‘nuptial dance’ starts when the male has localised the female and swims rapidly in close circles around her, with the dorsum towards her and antennae and cirri touching her. The initiation of the rapid swimming in the male is likely to be induced by pheromones produced by the female. The male emits sperm in slimy threads while swimming around her, and then leaves. The female, enveloped in semen, lays the eggs in a gelatinous sac about an hour later. Probably the female survives some time in the plankton while brooding the larvae.

10.6 DEVELOPMENT Phyllodocida. There are detailed morphological and anatomical developmental studies for a number of Phyllodocida (more molecular, ‘evodevo’ studies are dealt with in Chapter 5), although they tend to be concentrated to a few taxa, and there are many larger taxa for which we lack information. Examples are given below, a more complete list is provided in Table 10.1. Chrysopetalidae. For chrysopetalid larvae Cazaux (1968) provides a description of different stages of Chrysopetalum debile and Blake (1975) of Palaenotus bellis, with both appearing very similar. The early trochophore (seen in P. bellis only) has a prominent apical tuft that is absent in the later trochophores, when a pair of bundles with characteristic, stout barbed larval chaetae appear (see Fig. 10.11C). A meniscotroch is present in C. debile, and although not shown by Blake this appears to be the case also for P. bellis (B. Pernet, pers. com. in Rouse 2000). (Note that there are several definitions of ‘meniscotroch’—we apply the term in the larger sense; see Chapter 6.) Compound neurochaetae appear when there are about four segments, and paleae start to develop in the metatrochophore when it has about six chaetigers (Fig. 10.6A). Both larvae are planktotrophic. At these stages there are no obvious signs of enlarged, cephalised anterior dorsal and ventral cirri. Glyceridae. Cazaux (1967) and Blake (1975) also described glycerid larvae, Glycera convoluta by the former author and G. tenuis and Hemipodia simplex (as Hemipodus borealis; see Böggemann 2002) by the latter. The most complete one is on G. convoluta and the following relates to this description. The early trochophore has a distinct apical tuft that soon is reduced. A meniscotroch is present in the later trochophore. There are no specialized larval chaetae. In a metatrocophore with seven to eight chaetigers the antennae and palps start to develop, and soon thereafter the proboscis and jaws are visible and the typical glyceriform head is appearing (see also Fig. 10.6B). At this stage the two first chaetigers are uniramous, and the following ones biramous, as is the case in the adults. The larvae are planktotrophic. A metatrochophore with about six segments is illustrated in Fig. 10.6B Hesionidae. For the hesionid Ophiodromus obscurus there is actually a full developmental cleavage series, from the zygote to the trochophore

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Fig. 10.6 A. Scanning electron micrograph of pelagic chrysopetalid larva from Carrie Bow Cay, Belize. B. Scanning electron micrograph of pelagic glycerid larva from Carrie Bow Cay, Belize. G. Rouse.

(Treadwell 1901; as Podarke obscura). Following on from there, Blake (1975) and Haaland and Schram (1983) provide descriptions of the larval development of O. pugettensis and O. flexuosus, respectively. A noticeable

"%" Reproductive Biology and Phylogeny of Annelida difference between the two is that the former is lecithotrophic and the latter planktotrophic. The early trochopore of O. pugettensis, about 100 µm long, has a meniscotroch but no (observed) apical tuft, and remarkably early appearing pygidial cirri. There are no special larval chaetae. A 5-chaetiger nectochaete has developed median and paired antennae, segment 1 is provided with a pair of dorsal and a pair of ventral cirri, and segment 2 has elongated dorsal cirri, neurochaetae, but not yet elongated ventral cirri. Segment 3 has unusually small dorsal cirri, and this does not appear to be the case in other hesionid larvae. The larvae at this stage are feeding, presumably as encounter predators. This agrees with the description by Haaland and Schram (1983) of O. flexuosus, and differs from the adults in that the prototroch is still present, in that the median antenna has a much more dorsal position, in that dorsal and ventral cirri segment 3 have not yet become enlarged, in that neurochaetae are still present on segments 2 and 3, and in that no notochaetae are developed. Furthermore, the larvae described by Haaland and Schram has 10 distal papillae on the proboscis, whereas in the adults these are reduced. The development of a number of other hesionid larvae has also been described (see Table 10.1). Myzostomida. Larval development has been described for several Myzostoma taxa (see Kato 1952; Eeckhaut and Jangoux 1993) and it would appear that all fertilized eggs are spawned into the surrounding water, though Grygier (2000) reports brooding in some Endomyzostoma. The fertilized eggs of Myzostoma are very small at 50 µm in diameter or less and give rise to a planktonic, non-feeding trochophore-like larva that is ready to settle on a host after 5–8 days. Larvae develop two bunches of very long barbed chaetae, similar to those seen in the larvae of Chrysopetalidae. After settling, the larvae pass through further development in a cyst before becoming free-living. Larvae of endoparasitic and permanent cyst-dwellers have yet to be documented. Nephtyidae. For nephtyids there are no detailed descriptions of the whole larval development. An oral brush was described in the trochophore of Nephtys hombergi by Wilson (1936), but this was questioned by Phillips and Pernet (1996) as this feature is known only from some scaleworm groups. Thorson (1946) illustrates metatrochophores and nectochaetes of N. ciliata, and Lacalli (1980) some stages of N. caeca and Aglaophamus neotenus, plus trochophore and metatrochophores of N. incisa. As far as can be seen from these studies a meniscotroch is absent, as is an oral brush. Nereididae. There is a wealth of information on nereidids, and we here only provide a single example from Hediste diversicolor, described by both Dales (1950; as Nereis diversicolor) and Cazaux (1969, as Neanthes diversicolor). Dales also provided the early cleavages. Further taxa and references are given in Table 10.1. Like all known nereidids it has a lecithotrophic development (although Blake (1975) reported that Platynereis bicanaliculata may feed from the 5-chaetiger nectochaete stage). The trochophores have a prototroch and a telotroch but limited swimming ability, which is the case also for later stages. In the metatrochophores, three

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chaetigers are formed simultaneously, something with putative implications for the heteronomy theory by Iwanoff (1928) (see Chapter 5). At this stage the antennae and the first pair of dorsal situated cirri are beginning to form, whereas a ventrally situated pair of cirri forms later. However, the actual origin of these first cirri is not clear, and they are of importance for accurate homology statements between nereidids and other related groups. Dales refers to them as prostomial, and others (e.g. GilpinBrown 1959) have also argued that they are presegmental in nature. In contrast Pleijel and Dahlgren (1998), based on similarities with other, nonnereidid taxa, considered them as segmental. The issue remains to be settled. Illustrations of 4-chaetiger larvae are provided in Fig. 10.7A, B. Scale worms. A number of scaleworm groups, i.e. Pholoidae, Polynoidae and Sigalionidae, are characterised by having an oral brush in the planktotrophic larvae, although its absence in other scaleworms needs confirmation as these have not been studied in any detail. Cazaux (1968) gave a description of the larvae of Pholoe synophthalmica which is lecithotrophic, in spite of the presence of a prominent oral brush. It also has a paired band of apical cilia (akrotroch sensu Cazaux) and a meniscotroch. The 4-chaetiger metatrochophore starts to develop a median antenna and a pair of cirri from behind the prototroch (presumably the ‘tentacular’ cirri). At the 5-chaetiger nectochaete stage the palps and antennae are present, dorsal protuberances presumably are developing elytrae and dorsal tubercles, jaws are present, and notopodia and notochaetae start to develop. It is notable that Cazaux reported that this nectochaete was 27 days old, as the assumption is often made that lecithotrophic larvae have short planktonic stages. Pholoids, however, shows a lot of reproductive variation, and the taxa Laubierpholoe and Taylorpholoe instead are viviparous and brood the larvae in the elytrae (Laubierpholoe), or first in the body cavity and then in egg sacs under the elytrae (Taylorpholoe) (Pettibone 1992). Figure 10.8A, B shows a brooding female of Laubierpholoe swedmarki and a detached elytra with a larva in it. The trochophore of Sthenelais boa is similar to P. synophthalmica, although this instead likely has a planktotrophic development. It differs in lacking the paired band of apical cilia. There are several studies on larval development in the polynoid Harmothoe imbricata (see Table 10.1), the following is based on Korn (1958) and Cazaux (1968). It is planktotrophic and has a trochophore very similar to that of P. synophthalmica, including the bands of apical cilia. In the metatrochophore the parapodia with aciculae and noto- and neurochaetae pertaining to eight segments are developing, apparently at the same time or nearly so. A later metatrochophore has one additional segment, the dorsal and ventral cirri of segment have become elongated, elytrae are present on segments 2, 4, 5 and 7, as in the adults, and large pygidial cirri. A 7-chaetiger polynoid larva is shown in Fig. 10.8C. Phyllodocidae. For phyllodocids, we have no information on the development of the holopelagic alciopins (but see Buchholz 1869 for the description of a late larva, check no others), but a number of descriptions

"%$ Reproductive Biology and Phylogeny of Annelida

Fig. 10.7 A. Scanning electron micrograph of pelagic nereidid larva from Carrie Bow Cay, Belize. Dorsal view. B. Scanning electron micrograph of pelagic nereidid larva from Carrie Bow Cay, Belize. Ventral view, same scale as A. G. Rouse.

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Fig. 10.8 Scanning electron micrographs of scaleworms. A. Laubierpholoe swedmarki, adult female with juveniles brooded in the elytra. B. Detached elytra with juvenile. C. Pelagic polynoid larva. G. Rouse.

"%& Reproductive Biology and Phylogeny of Annelida

Colour Figure

Fig. 10.9 A. Micrograph of the syllid Sphaerosyllis with attached embryos from the Swedish west coast. B. Micrograph of median segments of mature female of Sphaerodum flavum from the Swedish west coast. G. Rouse.

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Fig. 10.10 Scanning electron micrographs of syllids. A. Female of Exogone with attached embryos from Belize. B. Male autolytin stolon, ‘polybostrichus’, from Belize. G. Rouse.

"& Reproductive Biology and Phylogeny of Annelida of benthic ones, especially from Phyllodoce (including Anaitides), and we here selected the description of P. mucosa by Cazaux (1969) since it is one of the most complete ones (see also Fig. 10.11A, B, D). The mucous bags are attached with a long string in sand or directly on sea grass (also described by Sach 1975). The larva is planktotrophic and has a pelagic life up to two months, and the early trochophore is provided with an apical tuft, a meniscotroch, a prototroch with short and long cilia, and a neurotroch running till the posterior telotroch. A metatrochophore one month old shows developing nuchal organs and 10 emerging parapodia, and the prolonged dorsal and ventral cirri of segments 1–3 are emerging one week later. After six weeks the proboscis is present, as are dorsal cirri, neuropodial lobes, neurochaetae and ventral cirri. The posterior incision on the prostomium and the nuchal papilla are not distinguished until after two months, whereafter the larva is ready to settle. Figure 10.11A, B, D shows a larva of Phyllodoce sp. and Tzetlin (1998) also described giant larvae of this taxon. Lecithotrophic development certainly occurs in other phyllodocids, as in Eteone barbata described by Thorson (1946), and the observation of many (especially smaller-sized) mature phyllodocid females with large eggs, such as Eumida arctica (about 150 µm; pers. obs.). Sphaerodoridae. There is not much information available on the development in sphaerodorids. Christie (1984) described the gametogenesis in Sphaerodorum flavum (as S. gracilis), and based on the egg size (200–250 µm in diameter) he suggested that development may be lecithotrophic and non-pelagic. Mileikovsky (1967) gives some information on juveniles of Sphaerodoropsis minuta and S. philippi (both as Sphaerodorum). The situation is not better with pilargids, where only very late pelagic larvae of Ancistrosyllis groenlandica with about 12 segments are known (Bhaud 1973). Syllidae. For syllids, in contrast, there are a large number of studies available; we here describe only two taxa with different modes of development. Syllis variegata has a pelagic larvae, planktotrophic although lasting only for about two days (Cazaux 1984). The first segments are added successively and a 3-segment stage has a well-developed median antenna, emerging paired antennae, a muscular proventricle, dorsal cirri on segments 1 and 3 (absent on segment 2), neuropodia with chaetae, and large pygidial cirri. The larvae of Epigamia alexandri were described by Qian and Chia (1989; as Autolytus alexandri). The trochophores were obtained from the brood pouches of the female stolons, and have an apical tuft, an akrotroch, a prototroch, a neurotroch and, laterally, a metatroch and a telotroch (note that the paddle cilia likely represent a fixation artefact, e.g. Short and Tamm 1991). Although an akrotroch has been described in several syllid larvae (Okada 1930; Dales 1951; Qian and Chia 1989), these all appear in Autolytinae, and appears to be absent from studied syllids outside this group. However, SEM studies shows that many adult Phyllodocida have a transverse dorsal ciliated band across the prostomium that could represent an akrotroch; the matter warrants further investigation. A late metatrochophore with five segments shows budding median and

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Fig. 10.11 A. Micrograph of metatrochophore of Phyllodoce sp., same scale as B. B. Micrograph of nectochaete of Phyllodoce sp. C. Micrograph of chrysopetalid nectochaete, same scale as B. D. Scanning electron micrograph of nectochaete of Phyllodoce sp., same scale as B. All from Carrie Bow Cay, Belize. G. Rouse.

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Reproductive Biology and Phylogeny of Annelida

lateral antennae, dorsal cirri and neuropodia with a few compound chaetae, but no ventral cirri (as in the adults). Brooding is common in syllids, and in Figs. 10.9A and 10.10A we show examples of brooding exogonins. Pisionidae. Åkesson (1961) demonstrated how segment 1 in the development of Pisione remota comes to surround the head region, and that the larvae prior to this metamorphosis share a number features with scaleworms. A recent study based on morphological and molecular data indeed corroborates that pisionids are scaleworms, and that they may be nested within sigalionids (Wiklund et al. in press). Interestingly, there is another pisionid, Pisionidens indica, which in the adult stage does not show the metamorphosed head region of Pisione (Aiyar and Alikunhi 1940), and this taxon certainly deserves further study.

10.7 EVOLUTION OF REPRODUCTIVE MECHANISMS The only studies that explicitly couple evolution of reproductive features with phylogeny within Phyllodocida are those by Nygren on syllids (1999) and, especially, on autolytins (1999, 2003, 2004). It is shown that epigamy is plesiomorphic in syllids, and that the occurrence of schizogamy in Syllinae and Autolytinae are non-homologous features. Both epigamy and schizogamy occur within Autolytinae but the results were ambiguous as to which of these is ancestral in that group. In the absence of other wellsupported phylogenies for the relationships within the Phyllodocida, discussions relating to the evolution of reproductive features would at present be highly speculative, and we postpone these to the future.

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

10.9 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 Naturhistorisk Forening 145: 11–21. Aiyar, R. G. and Alikunhi, K. H. 1940. On a new pisionid from the sandy beach, Madras. Records of the Indian Museum 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 zoological. 43: 135–199. Åkesson, B. 1967. On the nervous system of the Lopadorynchus larva (Polychaeta). Arkiv för zoologi 20: 55–78. Arwidsson, I. 1899. Studien über die Familien Glyceridae und Goniadidae. Bergens Museums Aarbog l898: 1–69.

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11

CHAPTER

Cirratuliformia Magdalena N. Halt1, Greg W. Rouse1, Mary E. Petersen2 and Fredrik Pleijel3

11.1 INTRODUCTION Fauchald and Rouse (1997) recognized the following family ranked taxa in their review of polychaetes: Acrocirridae, Cirratulidae, Flabelligeridae, Ctenodrilidae, Fauveliopsidae, Poeobiidae and Sternaspidae. However, among these there is no evidence of monophyly for Acrocirridae, Cirratulidae, Ctenodrilidae or Fauveliopsidae, though this was not surprising given the taxonomic history of the group (see below). Following their cladistic analyses, Rouse and Fauchald (1997) included these seven families in the taxon Cirratuliformia. These taxa, along with Flota (recently placed in its own family, Flotidae, Buzhinskaya 1996), were also referred to as Cirratuliformia in Rouse and Pleijel (2001). Cirratuliform polychaetes are characterized by the presence of a gular membrane (diaphragm), a heart body and no chaetae on the first segment (Rouse and Fauchald 1997; Rouse and Pleijel 2001). Most cirratuliforms also possess paired or multiple grooved palps. The present level of knowledge on reproduction, gametogenesis and development in this group is sadly limited. For instance in the fascinating pelagic taxa Poeobius and Flota, little is known about the general biology and life-history and no detailed reproductive studies have yet been performed. The reproductive biology of Cirratulidae has most recently been reviewed by Petersen (1999); in this an attempt was made to include as many published (and the author’s own unpublished) observations regarding reproduction as possible, no matter

1

South Australian Museum North Terrace, Adelaide. SA 5000, Australia and School of Earth and Environmental Sciences, University of Adelaide, SA 5005 Australia 2 Zoological Museum, University of Copenhagen, Universitetsparken 15, DK-2100 Copenhagen, Denmark; present address: Darling Marine Center, University of Maine, 193 Clark’s Cove Road, Walpole, ME 04573, USA 3 Tjärnö Marine Biological Laboratory, Department of Marine Ecology, 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

12 8

s1 s2

Described Sperm species

13

Epitoky

Fauveliopsis Laubieriopsis

Fauveliopsidae Hartman, 1971

Aphelochaeta Aphropharynx Caulleriella Chaetozone Cirratulus Cirriformia Ctenodrilus Dodecaceria Monticellina Protocirrineris Raphidrilus Raricirrus Tharyx Timarete

16 4

19 1 18 >26 33 8 2-3 12-15 10 13 2 3 8 13

s? s

s, l4 s s, l1 l s s6 s? s -

-

1 1 1 1 1 1 1 1? 1 1 -

Cirratulidae Ryckholdt, 1851 (including Ctenodrilinae Kennel, 1882)

Acrocirrus Macrochaeta

Acrocirridae Banse, 1969

Taxa

-

1 1 1 1? 1? 1 1 -

-

Herm

-

1 1 1 1 1 -

-

Vivip

-

1 -

-

Parth

pl? lc?7

d, lc d, lc ? d, lc pc d d, lc d -

-

Devel

-

1 1 1 ? -

-

BC

-

1, p 1?, a 1?, a 1, a 1, p 1, a 1, p 1, a 1, a 1, a 1, a

-

Asex

-

1 ? 1 -

-

Jelly

Table 11.1 Cirratuliform groups with known reproductive features (from Völker (1997), Petersen (1999; pers.obs), Rouse (1999a) and Rouse and Pleijel (2003) except where indicated): 1 presence, - no info, and ? unconfirmed reports. Sperm: l = long-headed, s = short-headed; Herm = hermaphroditism; Vivip = viviparity; Parth = parthenogenesis; Devel = development mode (lc = lecithotrophic, pl = planktotrophic, d = direct, in = indirect, pc = poecilogony); BC = brood care; Asex = asexual reproduction (a = architomy, p = paratomy); Jelly = eggs in jelly or mucus mass.

"'& Reproductive Biology and Phylogeny of Annelida

12

Sternaspis Otto, 1821

-

s9 s9 s s1 s11 s12 -

-

-

d, lc

d, lc d in, lc -

1 -

-

-

2

Sawada (1984) This study; Kudenov (1976) 3 Caullery and Mesnil (1898) 4 Also Dales (1951) 5 Jamieson and Rouse (1989) 6 Monticelli (1910); Sokolow (1911) 7 Blake and Petersen (2000); Petersen (2000b) 8 Indications of number of flabelligerid species within genera have been taken from Fauchald (1977). For all Flabelligeridae Rouse and Pleijel (2003) report a total of about 130 nominal species. 9 Spies (1977) 10 The placement of Poeobius within Flabelligeridae is based on Rouse and Pleijel (2003) and Burnette et al. (2005). 11 Robbins (1965) 12 Rietsch (1882)

1

18 1 43 7 1

Flabelligera Flabelliderma Pherusa Diplocirrus Poeobius10

Flabelligeridae de Saint-Joseph, 18948

Cirratuliformia

"''

# Reproductive Biology and Phylogeny of Annelida how small (Table 11.1, Petersen 1999). As these include many observations on live material, including color of eggs and sperm, they should also be of aid in helping to identify many of the older taxa whose identities are still being questioned. As will be reiterated in this chapter (see Rouse and Pleijel 2001, 2003) the traditional concept of Cirratulidae is ‘highly’ paraphyletic, and the whole group of Cirratuliformia requires revision. Although at present we do not know the actual extent of the paraphyly of Cirratulidae, it is clear that Ctenodrilidae are nested within that group. For this reason we refer to the latter as “Ctenodrilinae”, and we use the name “Cirratulidae” to include the group as traditionally delineated, together with Ctenodrilinae. For the traditional delineation of the group (i.e. excluding Ctenodrilinae), we only apply the vernacular name “cirratulins”. Given the previous detailed review by Petersen (1999) and the little new knowledge since then cirratulins will be treated in less detail here in order to include information on other cirratuliforms, especially Ctenodrilinae, a small group currently with only four genera, all of whose reproductive biology has been studied in some detail.

11.2 PHYLOGENY AND SYSTEMATICS 11.2.1 Taxonomic Review The taxonomic history of Cirratulidae and hence Cirratuliformia is complex, and the group was for many years a ‘dumping ground’ for various taxa of uncertain affinities, e.g., Cossuridae and Heterospio (Day 1967). The inclusion of such groups clearly made Cirratulidae a polyphyletic assemblage. These taxa are now placed elsewhere and at present Cirratulidae is likely to be paraphyletic, owing to the exclusion of such taxa as Ctenodrilidae and Acrocirridae. The first acrocirrid, described by Sars (1835), was Macrochaeta clavicornis. Macrochaeta, and the subsequently erected Acrocirrus, were placed in Cirratulidae by Grube (1873). This was accepted by later workers, until Banse (1969) erected Acrocirridae. Banse (1969) argued that Acrocirridae shared more features with Flabelligeridae (such as the structure of the epidermal papillae and the compound hooks) than with Cirratulidae. The similarity of Acrocirridae like Acrocirrus to Cirratulidae such as Dodecaceria (paired palps, anterior branchiae and anterior pair of segmental organs) suggests the possibility that the sister group for Acrocirridae could be among Cirratulidae (Rouse and Pleijel 2001). The fact that Flabelligeridae (and taxa like Fauveliopsidae and Poeobius) share, among other features, epidermal papillae with Acrocirridae means that they may also be nested in Cirratulidae. The retractile head found in Acrocirridae such as Chauvinelia and Flabelligella is a feature also seen in Fauveliopsidae, Flabelligeridae and Poeobius. The relationships among all these taxa need to be assessed in detail. The systematic placement of Ctenodrilinae was assessed by Mesnil and Caullery (1897), who argued that these animals showed several features in

Cirratuliformia

#

common with Cirratulidae. Subsequently, they were often considered as part of Cirratulidae (Day 1967; Fauvel 1927), though Dales (1963) considered them a separate taxon within Cirratulida, and Fauchald (1977) and Hartmann-Schröder (1996) as belonging to an isolated taxon Ctenodrilida, together with Parergodrilidae. A review of Ctenodrilinae by Petersen and George (1991) resulted in the movement of Raricirrus from Cirratulidae to Ctenodrilinae. They also considered Ctenodrilus, Raphidrilus and Aphropharynx as valid parts of Ctenodrilinae. Zeppelina, which has had a long history as part of Ctenodrilinae, was shown by George and Petersen (1991) to be comprised various taxa belonging in Cirratulidae. It seems quite likely that the Ctenodrilidae may not be monophyletic and may therefore cease to exit as a separate family. Fauveliopsidae was erected by Hartman (1971) to accommodate a number of taxa previously included in Flabelligeridae, though it is now regarded as only containing Fauveliopsis and Laubieriopsis (Petersen 2000). Blake and Petersen (2000) provide a detailed review of the taxonomic history of the group. The presence of the retractable head, the papillose body and certain chaetal forms suggests that the recognition of Fauveliopsidae may make Flabelligeridae paraphyletic. Fauchald and Rouse (1997) indicated that the monophyly of Flabelligeridae as currently formulated was supported by the presence of gills on an expanded peristomial membrane. This is probably an erroneous statement and Rouse and Pleijel (2001) instead suggested that the branchiae of Flabelligeridae are more likely to be segmental structures and thus homologous with the anterior branchiae of Acrocirridae and Cirratulidae such as Dodecaceria. Rouse and Fauchald (1997) found Flabelligeridae to be the sister group to Acrocirridae or as part of clade (with Fauveliopsidae, Poeobiidae and Sternaspidae) that was the sister group to Acrocirridae. This clade was then the sister group to Cirratulidae. This relationship was also postulated by Mesnil (1899). In the review by Fauchald (1977), Flabelligeridae contained 126 nominal species placed into 14 genera, with groups like Brada, Flabelligera and Pherusa containing the majority of taxa. There has been no major revision of the group since that time and there have only been a few recently described taxa. Poeobius meseres, a pelagic polychaete, was described by Heath (1930) and for which he also erected the family Poeobiidae. Heath regarded Poeobius as a ‘connecting link’ between polychaetes and Echiura. This view has largely been rejected in the sense that Heath argued, and since Hartman (1955) and the detailed study by Robbins (1965), Poeobiidae has been regarded as closely related to Flabelligeridae (e.g., Fauchald 1977). The presence of a retractable head, a pair of grooved palps, an anterior ring of branchiae, epidermal papillae, and a gelatinous mucus-sheath all suggest that Poeobiidae may be nested within Flabelligeridae. Rouse and Pleijel (2003) showed, via morphological evidence from a cladistic analysis, evidence for Poeobius to be nested within Flabelligeridae.

#

Reproductive Biology and Phylogeny of Annelida

Flota flabelligera, another pelagic polychaete, was described by Hartman (1967) and placed in Flabelligeridae, though she subsequently transferred it to Fauveliopsidae. Buzhinskaja (1977) described Flota vitjasi as part of Fauveliopsidae but later, she (Buzhinskaja 1996) erected a new family taxon for it, Flotidae. The presence of a gelatinous sheath, epidermal papillae and barred chaetae suggests a close relationship between Flota and Flabelligeridae. Rouse and Pleijel (2001) showed Flota to be nested within Flabelligeridae and sister to Poeobius. Although molecular analysis is still lacking, recent related data on Poeobius (Burnette et al. 2005) support the placement of this group within Flabelligeridae. During the 20th century, Sternaspis (containing around 15 nominal species) has generally been considered as an “aberrant” polychaete group, and so in recent classifications they were considered a separate family and order (Dales 1962; Fauchald 1977). Mesnil (1899) did review the anatomy of Sternaspis in relation to Flabelligeridae, but concluded that the points of similarity such as the retractable head, the pair of anterior segmental organs and the epidermal papillae represented convergences. Dahl (1955) argued for a close relationship between Flabelligeridae and Sternaspis and raised the possibility that taxa such Echiura and Sipuncula could also be closely related to these groups. In most of their cladistic “complete analyses”, Rouse and Fauchald (1997) found Sternaspis to form a polytomy with Flabelligeridae and Fauveliopsidae. This group was part of a larger clade that included Poeobiidae, Acrocirridae and Cirratulidae.

11.2.2 Recent Systematic Revisions Two recent phylogenetic analyses involving cirratutliforms have appeared; Rouse and Pleijel (2003), using morphology and Bleidorn et al. (2003), using 18S rDNA. Rouse and Pleijel (2003) performed a morphological analysis on the cirratuliform assemblage outlined above, resulting in the strict consensus tree is shown in Fig. 11.1. The traditional delineations of taxa into their family groups are indicated across the appropriate branches. In the grouping of Cirratulidae, Ctenodrilidae, Fauveliopsidae and Flabelligeridae, as traditionally delineated, no monophyly could be demonstrated. Poeobiidae, Flota and Sternaspis were only included as single terminals and so their monophyly was not assessed here. Of the taxa included with multiple terminals, only Acrocirridae was found to be monophyletic in all shortest trees. In one of these 28 shortest trees (Fig. 11.1) it can be seen that Cirratulidae is made paraphyletic by recognition of Acrocirridae, Ctenodrilidae, Fauveliopsidae, Flabelligeridae, Flotidae, Poeobiidae and Sternaspidae. Furthermore, Flabelligeridae is paraphyletic because of Poeobiidae and Flotidae nested inside, and Fauveliopsidae by recognition of Sternaspidae. Bleidorn et al. (2003) had a smaller sampling of cirratuliforms but did show that Ctenodrilus was nested inside Cirratulidae. Given the support from two different analyses for this we here place Ctenodrilidae within Cirratulidae as Ctenodrilinae.

Cirratuliformia

#!

Fig. 11.1. A. Strict consensus tree of 28 shortest trees found in the cladistic analysis of Cirratuliformia. From Rouse, G. W. and Pleijel, F. (2003). Hydrobiologia 496: 175-189, Fig. 3. B. One of 28 shortest trees in the Cirratuliformia analysis with the outgroup removed, showing various characters used in the analysis; p. comp. chaetae - pseudocompound chaetae. * transformation for chlorocruorin is indicated at that point on the assumption that several taxa scored with ‘?’ have this blood pigment. ** barred chaetae are subsequently lost in Poeobius. **** grooved palps have been lost twice on this tree topology. **** branchiae are lost in Ctenodrilus. From Rouse, G. W. and Pleijel, F. (2003). Hydrobiologia 496: 175-189, Fig. 4.

#" Reproductive Biology and Phylogeny of Annelida

11.3 ANATOMY WITH REFERENCE TO THE REPRODUCTIVE SYSTEM 11.3.1 Gonads and Gonoducts An overview of the current classification and the number of recognized species is given in Table 11.1 though genera are omitted where there are no data on reproduction. A large number of additional species and more inclusive taxa are waiting to be described, and many species will undoubtedly be moved to other genera. With the exception of some cirratulids, which can be hermaphroditic, all taxa considered in this section are gonochoric and exhibit no sexual dimorphism (Marion 1874; Mesnil 1899; Okuda 1934; Banse 1969; Spies 1977; Riser 1987; Petersen 1999; Fiege and Buetfering 2000; Petersen 2000a). The reproductive system of cirratulins is basically simple, usually with gametes developing freely in the coelom (Olive 1973). In general, there are no discrete ovaries although Gibbs (1971) reported distinct ovaries in Tharyx marioni. The gametes are usually either released into the free water masses during spawning by epitokes or deposited in variously shaped jelly masses by iteroparous species e.g., Cirratulus cirratus. Gametes leave the body through a pair of coelomoducts on the posterior part of the body (Caullery and Mesnil 1898; Olive 1970, for Chaetozone setosa see Fig. 11.3E) or through genital pores (for Raricirrus variabilis; Dean, 1995). From the available anatomical descriptions of Acrocirrus and Macrochaeta (Marion 1874; Mesnil 1899; Okuda 1934; Banse 1969; no information is available on Flabelligella and Chauvinelia) it appears that the gonads start in the first or second abdominal segment and continue for 1725 segments (although this may be difficult to determine as the body appears to be poorly differentiated). In one Acrocirrus individual, Okuda (1934) recorded 25 pairs of gonads in as many segments. Gametes are reported to occur after segment 10 (Okuda 1934) but no gonoducts have been documented. Free-floating gametes have also been observed in the abdominal coelomic cavity (Okuda 1934). The so far unstudied anterior segmental organs and the accompanying looped nephridia which have been reported (Marion 1874; Marion 1875; Mesnil 1899; Okuda 1934; Banse 1969) appear to more closely resemble excretory than reproductive organs. No further information is available on the reproduction or larval development of this group. From the limited studies available of Fauveliopsis internal anatomy, females appear to bear single or paired gonads on the posterior wall of the septum generally between the 5th and the 10th chaetigers (Núñez et al. 1997; Riser 1987). An elongated, fluid-filled ovisac (formed from the septal walls) extends posteriorly from the ovaries to chaetiger 8-10 in Fauveliopsis arenicola (Riser 1987) and carries the developing eggs. In Laubieriopsis cabiochi genital papillae (Fig. 11.3D) also occur in this region (Petersen 2000b). In males, testis extend from chaetiger 5-8 in Laubieriopsis brevis and from 5-12 in L. arenicola with the vas deferens leading from the sacs to the

Cirratuliformia

##

nephridial papillae (Riser 1987). The male pores occur anteriorly to chaetiger 7. The bulk of information available about flabelligerid reproduction originated from two seminal papers by Spies (1977) on Flabelligera and Amor (1994) on Pherusa. Robbins (1965) described most of the anatomy known for Poeobius. In all, the sexes are separate and discrete gonads are restricted to the middle segments. There appear not to be any morphological differences between the sexes with the exception of the colouring of the gametes visible through the body wall. The body of Flabelligera commensalis is divided by a sac-like septum which encloses 8 pairs of gonads and the digestive system (Spies 1977). The total number of gonads appear to vary between 6-8 in mature individuals, perhaps due to the slow development of anterior gametes in young worms (Spies 1977). A sub-branch of the main segmental circulation system supplies blood to the gonads. In Flabelliderma commensalis gametes mature within the walls of blood vessels surrounding the gonads (Spies 1977). The ovaries are elongate and are enveloped by an arrangement of blood vessels on their surface. Similarly, the disc-shaped testis that line chaetigers 2-9 are heavily vascularized and are enclosed in a thin epithelial membrane, presumably to allow for the migration of maturing spermatids. Reproductive anatomy of Sternaspis closely resembles that of some Flabelligeridae. All known Sternaspis species have separate sexes and exhibit no sexual dimorphism (Fiege and Buetfering, 2000). Paired gonads are discrete sacs situated behind segment 6, each with a duct leading posteriorly to genital papillae (Rietsch 1882). Gametes are ejected via pores in the genital papillae. Ovaries are composed of four irregular lobes (Rietsch 1882; Fig. 11.3B). The gonads are highly vascular with blood vessels penetrating into and enclosing the organs (Goodrich 1945).

11.3.2 Accessory Appendages Many mature cirratuliforms have various arrangements of reproductive papillae, often containing a gonopore. In fauveliopsids both sexes can have one or two pairs of ‘genital papillae’ which occur on the same segment (Riser 1987; Petersen 2000a). Riser (1987) noted that the male gonoduct exits close to or through the papillae but this remains unconfirmed. In Poeobius gametes are released through a pair of specialized lateral nephridia close to the gonads (Robbins 1965). In contrast, the nephridiopores on the branchial membrane in Pherusa serve to discharge both waste and gametes (Goodrich 1945; Amor 1994). All Sternaspis species have laterally-placed genital papillae used to eject gametes (Fiege and Buetfering 2000; see Fig. 11.2A). The morphology of the genital papillae appears to be species-specific: in Sternaspis scutata the papillae are long, broad and extended whilst in S. fossor they are slender and contracted (Petersen 2000a). In S. scutata the papillae appear to originate in the groove between segments 6 and 7 (Rouse and Pleijel 2001; in contrast to Fiege and Buetfering 2000).

#$ Reproductive Biology and Phylogeny of Annelida

Colour Figure

Fig. 11.2. Some representatives of Cirratuliformia. A. Sternaspis scutata from Banyuls sur Mer, France. B. Aphelochaeta sp. female with eggs, from Iceland. C. Paratomy in ctenodrilus serratus from Lizard Island, Australia. D. Fauveliopsis sp. female from Banyuls sur Mer, France. E. Macrochaeta australiensis from Australia. F. Dodecaceria concharum, male epitoke from Sweden. G. Asexual reproduction in Dodecaceria sp. from Australia. All images by G. Rouse.

Cirratuliformia

#%

Few species of ctenodrilins, e.g. some Ctenodrilus and Raricirrus, have stout neurochaetal spines and blades restricted to a few segments (see Fig. 11.4C; Dean 1995; Petersen and George 1991; Rouse and Pleijel 2003). These additional spines have been noted in smaller individuals (see Table 11.3) and seem to be absent in larger worms. Smaller C. serratus also appear to have both oocytes and sperm and have more gametes in general (see Table 11.3). Since many ctenodrilins are protandric hermaphrodites, it is feasible that such spines may serve a reproductive function. There is also a sole record of a reproductive stylet, leading into the seminal vesicle, in Raricirrus variabilis (Dean 1995).

11.4 ENDOCRINOLOGY OF REPRODUCTION Little is known of the endocrinology in the Cirratuliformia, and the limited data available concern members of the Cirratulidae. This group does not appear to be governed by environmental stimuli (Olive 1970). Olive (1970) treated the reproduction of a Northumberland population of Cirratulus cirratus, and later (1973; 1975) he examined the germinal epithelium activity in cirratulids in relation to periodicity of reproduction. He found that the developing oocytes in the coelom exert a local negative feedback to the ovaries in the segment (Olive 1973). He postulated that the vitellogenic gametes release a hormone-like regulator chemical that reduces germ cell proliferation in the germinal epithelia. The chemical nature of this mechanism is unknown. The removal of the cerebral ganglia has no direct effect on oocytes growth but appears to partially inhibit spawning (Olive 1973). Since reproduction in C. cirratus is asynchronous, and females shed all their coelomic gametes when they spawn, the gonadotrophic effect of the ganglion results in a continuous proliferation of oocytes in readiness for the next spawning event. Mature oocytes in the coelom then inhibit further gamete production by the ovary. In addition, a growth hormone is believed to govern both somatic and gonad cell proliferation (Clark and Olive 1973; Olive 1973). Growth tends to suppress epitoky, and is in itself suppressed by the presence of vitellogenic oocytes. Environmental control of reproduction was reviewed by Olive (1984).

11.5

OOGENESIS

Ovary structure and oogenesis in Cirratulus cirratus and the regulation of ovary function in this species were treated by Olive (1971, 1973). As described above, the proliferation of oocytes is largely governed by feedback mechanisms. The eggs undergo vitellogenesis in the coelom in readiness for the next spawning event (Olive 1973). Cirratulid eggs exhibit a great variation in size and color, e.g., for Raphidrilus nemasoma, greencolored fertilized eggs measured 125-145 µm (Qian and Chia 1989) and some species of Aphelochaeta and Cirratulus eggs measure 400-450 µm in

#& Reproductive Biology and Phylogeny of Annelida diameter (Petersen 1999). Clark (1973) and Schroeder and Hermans (1975) provide a further review of Cirratulidae egg morphology. Acrocirrid oocytes have been reported in specimens collected over Northern Hemisphere summer. In Macrochaeta and Acrocirrus polygonal eggs with a sculptured outer membrane measured 120-130 µm in diameter, and had large germinal vesicles (Banse 1969; Kudenov 1976). They are believed to undergo extra-ovarian development. In Poeobius, mature oocytes measure approximately 160 µm in diameter and are yellow in color. The egg color facilitates sex recognition in this group. In the flabelligerid Pherusa sp., green-tinted eggs measure about 140 µm in diameter (Amor 1994). In contrast to other polychaetes (with the exception of Polynoidae, Rouse 1999a) gametes complete their development within the gonads, and are only released in readiness for spawning (Spies 1977). Riser (1987) noted developing oocytes and germinal vesicles in the ovaries of fauveliospids. In the ovisac, the nuclei appeared to be in prophase and no germinal vesicles were present. Oocytes began vitellogenesis in the ovary and continued proliferation in the ovisac surrounded by a cyanophilus membrane which thickened with maturity. No nephridial pore was found at the end of the ovisac which envelops the developing oocytes. In Laubieriopsis cabiochi two rows of 10 oocytes, 185-200 µm in diameter, lined the ovisac (Petersen 2000a). The variation in egg sizes found by Riser (1987) has lead to suggestions of direct or planktotrophic development (Blake and Petersen 2000), however no larval studies are available (Petersen 2000a). The oogenesis of Sternaspis is well documented by several early researchers (Child 1900; Rietsch 1882; Vejdovsky 1882). The yellow oocytes are around 150 µm in diameter, are slightly tapered at one end and are surrounded by a thick membrane envelope (Strathmann 1987). In Sternaspis scutata a vascular loop supplies blood to the developing oocytes.

11.6 SPERMATOGENESIS Studies of spermatogenesis in cirratulids are still few. The vast majority of species have short-headed sperm, associated with external fertilization, but a few species, including a northern European Aphelochaeta with brood protection, have long-headed sperm that bear further study (see Fig. 11.5F; Dales 1951; Petersen 1999). In Ctenodrilus serratus sperm head is spherical with a cluster of small mitochondria at the base and a short flagellum (Rouse 1999a). The acrosome lies close to the plasma membrane and is separated from the nucleus by cytoplasm. This species was found by Rouse (1999; Rouse and Pleijel 2003) to be viviparous which implies that the sperm type is most likely introsperm. This sperm shape is illustrated from another Ctenodrilus from Belize in Fig 11.5C, E (Rouse, pers. obs.). Sperm with the same function may well vary in shape as in another viviparous species, Raphidrilus nemastoma (Monticelli 1910; Sokolow 1911), the sperm had a more elongated headpiece. In cirratulids, early sperm development occurs

Cirratuliformia

#'

within the gonads, and the developing spermatids are then released into the coelomic cavity to completely mature (Gibbs 1971). In Dodecaceria caulleryi, Gibson (1981) observed morulae dense with developing sperm in the coelom. In acrocirrids, the sperm has a spherical headpiece and a flattened acrosome (Kudenov 1976), presumably related to external fertilization. Sawada (1984) recorded four mitochondria, a globe shaped nucleus, and well-developed cytophores in Acrocirrus validus. New description of the sperm of Macrochaeta clavicornis is provided here (see Fig 11.5A). In agreement with Kudenov (1976) the sperm was found to be round to ovoid in shape with a 9+2 flagellum. Akin to the observations of Sawada (1984) for A. validus, there is a spherical nucleus with four mitochondria. An ellipsoidal acrosome lies at the apex of the nucleus. It has a large subacrosomal space with a narrow basal opening that expands to a broad space. As a consequence, the anterior layer of the acrosome is thin with kidney-shaped lateral extensions. This is in contrast to the description of a ‘flattened’ acrosome for Macrochaeta australiensis by Kudenov (1976) although this is difficult to compare due to a lack of illustrations in his paper. A proximal centriole is located centrally between the mitochondria. The distal centriole is linked to the proximal centriole and to the plasma membrane by an anchoring apparatus. There is a pronounced nuclear fossa. Interestingly, each mitochondrion contains an oval-shaped inclusion, the function of which is unknown. Acrocirrid spermatogenesis was partially described by Kudenov (1976) and in more detail by Sawada (1984). Kudenov (1976) recorded all stages of sperm development, including sperm platelets, morulae and mature spermatozoa in the coelom of male Macrochaeta australiensis from southeastern Australia. The spermatogonia leave the gonads during the early stages of spermatogenesis, and continue to proliferate in the morulae. Gametogenic stages undergo proliferation until mature sperm detach and float freely in the coelomic fluid (Sawada 1984). In the flabelligerid taxon Flabelligera the sperm has a round head and a midpiece with four to five mitochondria arranged around the base of the flagellum (Spies 1977, Rouse (1999). In Diplocirrus glaucus (new description provided here), there is a flattened broad acrosome that covers the anterior surface of the head (see Fig. 11.5D). The small subacrosomal space is centrally restricted. A pair of centrioles is linked together and to the plasma membrane by an anchoring apparatus for the axenome (Rouse 1999a). From morphology Rouse (1999) tentatively classified flabelligerid sperm as ectaquasperm implying its function in external fertilization. The sperm of Poeobius appears to bear many similarities to that of flabelligerids. They have a round head and a spherical acrosome with three to five mitochondria and a long flagellum, possibly indicating a similar externally fertilizing mechanism (Robbins 1965). In contrast to most other polychaetes, in flabelligerids the whole process of spermatogenesis is enclosed within the testis and the adjoining

# Reproductive Biology and Phylogeny of Annelida blood vessels (Spies 1977). As an example in the flabelligerid Flabelliderma commensalis, the primary spermatocytes undergo proliferation and migrate toward the centre of the testis, where they develop their large nuclei. This development mode is characteristics of most flabelligerids and Poeobius known to date. In Laubieriopsis brevis (Fauveliopsidae) sperm at various developmental stages was found in reproductive sacs, presumably testis, extending between chaetigers 5 and 8 (Riser 1987). There is no public record of the fauveliopsid sperm type and morphology. Round-shaped spermatozoa has been recorded for Sternaspis (Rietsch 1882).

11.7 MATING AND FERTILIZATION Most of what we know about fertilization in this group is largely inferred from the shape of the spermatozoa and from the morphology of the reproductive system. In fauveliopsids, Riser (1987) noted that no sperm was found inside the body cavity of Fauveliopsis arenicola, possibly indicating external fertilization. This has also been suggested for Sternaspis (Gravier 1923). Riser (1987, p. 23) also noted ‘a single clear spot which looks suspiciously like a male pronucleus in each oocyte undergoing vitellogenesis’. This could indicate an internal fertilization mechanism or perhaps hermaphrodism, more unlikely as only one type of gonad was found. Robbins (1965) noted that Poeobius meseres was reproductively mature during the winter months of the Northern Hemisphere. From the shape of the spermatozoa, this group appears to have external fertilization. This suggestion is supported by Robbins’ (1965) finding of mature gametes in the nephridia. Free-swimming sperm were found in the anterior coelom of the flabelligerid Piromis roberti indicating spawning (Spies 1977). Another flabelligerid, Pherusa sp., was found to spontaneously spawn in seawater leading to successful fertilization (Amor 1994). In cirratulids, external fertilization is most common, followed by asexual reproduction and epitoky (Petersen 1999). Various species of Dodecaceria illustrate this reproductive variability (Fig 11.2G): D. caulleryi first reproduces asexually and then metamorphoses into epitokes (Gibson 1981); D. concharum reproduces asexually by fragmentation and sexually by epitokes; most D. ater are parthenogenic females but female, and less commonly male, epitokes have also been recorded (see Caullery and Mesnil 1898: pl. 1, fig. C2; Spies 1977) and D. pulchra from South Africa reproduces asexually and is parthenogenic, but no epitokes are known for the species. Other species of Dodecaceria have also been shown to be parthenogenic (see Table 11.1 for summary). It is not yet clear whether D. saxicola (Grube 1855) from the Mediterranean, for which a male epitoke with ripe sperm and red eyespots but without any epitokous capillaries is illustrated in Petersen (1999, fig. 1C-D), is identical to the northern European D. ater. Their great similarity may confirm the presence of male

Cirratuliformia

#

epitokes in D. ater. Asexual reproduction by architomy also occurs in many Dodecaceria, and some Caulleriella, Cirratulus, Timarete and Protocirrinereis (Petersen and George 1991; Petersen 1999). In ctenodrilins this mode of reproduction occurs in Raricirrus and Ctenodrilus (Monticelli 1910; George and Petersen 1991; Petersen 1999). Metamorphosis into an epitoke is documented by Gibson (1981) on two species of Dodecaceria. The first stage involves the development of swimming chaetae and eyes. In some species of Cirratulus, Dodecaceria and Raricirrus, the parents develop fine natatory capillaries, the nuchal organs may become more prominent, and eyespots may appear (Fig. 11.2F). Mature male epitokes have paired ciliated sperm ducts and females have specialized, but ‘inconspicuous’ (Gibson 1981), segmental pores. Atokous and epitokous stages of the parthenogenetic Dodecaceria saxicola (Grube 1855) are shown by Petersen (1999: Fig. 1A-D). Epitokes die following spawning. Gibson (1997) found that at epitoky the cerebral ganglia enlarge and suggested that this was “presumably due to epitoky and not gametogenesis since it [the enlargement] is not evident in Dodecaceria pulchra which does not have an epitoke.” In sexually developing ctenodrilins, females are often viviparous as is the case in Ctenodrilus serratus (Rouse 1999b; Rouse and Pleijel 2003). Fertilization is internal and eggs develop in the coelom until they are a few chaetigers in length when they ‘hatch’ (see Fig. 11.3A for Raphidrilus nemasoma). When fertilization occurs outside the body, the fertilized eggs often have some form of brood protection (Table 11.2, Table 11.3 and Petersen 1999). Most often it consists of a jelly mass but brooding in a burrow and covering with shed tentacles have also been recorded. In the hermaphroditic Aphelochaeta sp. (Petersen 1999), the young are brooded in a specially constructed burrows (see Fig 11.3C).

11.8 DEVELOPMENT Most cirratulids have lecithotrophic development with some (e.g. Cirriformia tentaculata) having a brief planktonic phase (Petersen 1999). Early development usually takes place within gelatinous masses deposited in the sediment, as in the case of Cirriformia spirabrancha (Blake 1975). Most available records are based on species with direct development and some type of brooding. The larvae begin feeding when they emerge from the egg mass, even though most have a well-developed gut at early trochophore stage (Qian and Chia 1989). Fertilized eggs are often laid in jelly masses which protect them from infection by protozoans (Qian and Chia 1989). In Qian and Chia’s (1989) study of the development of Raphidrilus nemasoma, on average 1000 vivid green eggs (125-145 µm in diameter) were attached to the wall of the jelly mass by mucus strings. Their development is asynchronous, with embryos to late trochophore stages found in the jelly mass. Eggs reach the blastula stage approximately 48 hours after fertilization, and develop into young

7

3

10 x 0.3

17 x 0.7

7.59 x 0.6

Raphidrilus nemasoma

Raphidrilus sp.4

Raricirrus maculatus

Raricirrus beryli

Raricirrus variabilis8 none seen

A

≥35 35

A

–

A

30

>19

35

P

P

1

Asex

short

–

short

short?

–

long

3

–

Sperm Head

yes

yes

yes

6

yes?

no

no

3

–

–; ca.100

–; 323.4 ± 1.57 (n = 29)

–

Eggs: Color, Size, µm

no

no

no

–; to 140 in epitoke

cream, yellow, peach; to 90

7

–

red, 2 pairs green, in jelly; in larvae; 125-145, direct adults? development

in Form B

–

not in atoke

Epitoky Eyes

?

yes, SV, stylet, GP

no

no?

no

protandric

3

–

Herm

yes

no

no

no

no

yes

3

–

Vivip

no

none seen

5 reproductive spines in M

–

spines in M

giant chaetae in M

3

–

Sex Dimorph

2

Fixed division between segments 6-7, 7-8 and 8-9 (Wilfert 1974) Wilfert (1973) 3 Völker (1997), unpublished Ph.D. dissertation, included with her permission 4 Sensu Qian and Chia (1989), as Raphidrilus nemasoma, not this, see Petersen and George (1991: 206), also P.-Y. Qian, pers. comm. to M.E. Petersen 5 Petersen and George (1991:Fig. 1C,D) 6 Petersen in Dean (1995: 177) 7 Color in formalin 8 Dean (1995)

1

2.8 × 0.12-0.9 (strain C)

F=13 M= 18

3

2

Ctenodrilus serratus 3

15

1.5 × 0.15

Aphropharynx heterochaeta

Max No Segs

Max size: L × W, mm

Species

Table 11.2 Reproductive features of species of Ctenodrilinae; Asex = asexual reproduction (A = architomy, P = paratomy), Eyes = presence of eyespots, F = females, GP = genital pore, Herm = hermaphroditism, L = length, M = males, Max = maximum, Sex Dimorph = sexual dimorphism, Segs = segments, Stylet = reproductive stylet, SV = seminal vesicle, Vivip = viviparity, W = width, ? = unconfirmed report, – = unknown. Information compiled by M. E. Petersen, from original descriptions unless otherwise noted.

# Reproductive Biology and Phylogeny of Annelida

Cirratuliformia

#!

trochophores within three days. At this stage larvae have a band of paddleshaped cilia that form the neurotroch. This feature, most probably used to increase the surface area and strengthen attachment to the mucosal walls, is lost prior the leaving the egg mass. After around 11 days the 3-4 chaetiger larvae emerge from the egg mass and metamorphose over the following two weeks. Cirratulids have a wide range of reproductive and development modes ranging from gonochorism to protandric hermaphrodism (Table 11.2; Monticelli 1910; Sokolow 1911; Petersen 1999). A number of species, e.g. Ctenodrilus serratus (Monticelli 1907), Raricirrus nemasoma (Fig. 11.3A;

Fig. 11.3. Reproductive anatomy. A. Raphidrilus nemasoma with various stages of young in the body. Adapted from Monticelli, F.S. (1910). Archivio Zoologico Unione Zoologica Italiana Napoli 4: 401-436, Table 12, Fig. 9. B. Sternaspis scutata ovary. Adapted from Rietsch, M. (1882). Annales des Sciences Naturelles (Zoologie et Paléontologie) (série 8) 13: 1-84, Fig.53. C. Aphelochaeta sp. brooding eggs in a burrow. From Petersen, M. E. (1999). Hydrobiologia 402: 107-128, Fig. 4. D. Laubieriopsis cabiochi with oocytes maturing in the coelom. Adapted from Petersen, M. E. (2000b). Bulletin of Marine Science 67: 491-515, Fig. 2G. E. Chaetozone setosa internal anatomy showing ovaries and gonoducts. Adapted from Meyer, E. (1888). Mittheilungen aus der Zoologischen Station zu Neapel 8: 462-662, Plate 23.5.

#" Reproductive Biology and Phylogeny of Annelida Monticelli 1910) and Raricirrus variabilis (Dean 1995) exhibit viviparity. Rouse (pers. obs Table 11.3) observed multiple larval developmental stages in the coelom of C. serratus and noted the presence of seven chaetiger juveniles. The same species have been recorded to reproduce by paratomy (Peters 1923). Similarly, Monticelli (1910) described the life-history of Ctenodrilus serratus which gives birth to live young (emergent larva is shown in Fig.11.4B) as a protandric hermaphrodite and reproduces by achitomy at other times. Viviparity also occurs in four to five other taxa of Cirratulidae and paratomy is also common in representatives of Cirratulus, Protocirrinereis, Timarete and Dodecaceria (George and Petersen 1991; Petersen 1999). As yet undescribed juvenile of a species of Dodecaceria from Bondi Beach, Australia (collected by G.W. Rouse) is pictured separating from the parent stock in Fig. 11.2G. Sternaspis larvae have been reared under laboratory conditions, and appear to be lecithotrophic and settle in less than two days (Rietsch 1882). Strathmann (1987) also reports benthic, lecithotrophic larvae in Sternaspis from California. In the flabelligerid Pherusa, larval development is lecithotrophic and larvae settle from the plankton after around a week. Pherusa exhibits indirect development, as demonstrated by Amor (1994), with a lecithotrophic pelagic stage, followed by a pelago-benthic stage and a brief larval benthic stage. Early in the pelagic development a dorsal depression signals the future location of the anus. The number of branchiae and eyespots increase during this time and adult chaetae appear. Both the

Fig. 11.4. Aspects of ctenodrilin reproduction. A. SEM of Ctenodrilus from Belize. B. Emergent larva; c - ciliated tufts. C. Chaetae on male adult; s - copulatory spines. D. LM image of a developing embryo in the coelom of female. All images by G. Rouse.

Cirratuliformia

##

Table 11.3 Morphological characteristics of Ctenodrilus sp. sampled off Carrie Bow Cay, Belize, by G.W. Rouse in 1992. Where available the mean is given with the total sample number in brackets. The data suggests this Ctenodrilus sp. is a protandric hermaphrodite.

Length of adults

Total no. segments

Total larvae

0-1 mm 1-2 2-3 3-4 4-5 5-6 6-7 7-8 8-9 9-10

9 10 12 13 14 14 15 14 -

7 [n=6] 10 [n=5] 9 [n=3] 12 -

[n=5] [n=8] [n=8] [n=6] [n=14] [n=3] [n=2] [n=2]

oocytes 2 3 -

Present in segments: sperm larvae blades

spines

2-10 2-9 2-9 2-14 2-9 -

1-3 1-2 1-11 -

2-11 2-14 2-12 ? -

4-8 3-8 3-11 -

Fig. 11.5. Cirratuliformia sperm: A-D. TEM of longitudinal section of sperm. A. Macrochaeta clavicornis. B. Flabelligera sp. from Japan. C. Ctenodrilus from Belize. D. Diplocirrus glaucus sperm with insert showing one spermatozoon. E. Light micrograph of sperm from male stage Ctenodrilus sp. from Belize. Note short flagellum F. Light micrograph of sperm from Aphelochaeta sp. from Iceland. Abbreviations: a – acrosome, f – flagellum, m – mitochondrion, mi – mitochondrial inclusions, n – nucleus. All images by G. Rouse.

#$ Reproductive Biology and Phylogeny of Annelida mouth and the anus are developed when the larva begins its benthic stage. During the late pelago-benthic stage larvae also starts to secrete an adhesive substance from a retractable organ on the posterior of the worm, presumably used in settlement and burrowing (Amor 1994). The development of Flabelliderma and Flabelligera was described by Gravier (1923) and Spies (1977). Larvae of Flabelliderma commensalis underwent rapid growth from 7-15 segments within 30 hours when cultured in vitro (Spies 1977). Early larvae (three to seven segments) bear three circular rows of cilia, the prototroch, metatroch and telotroch, which it uses to swim. There is a pair of eyespots on the head. The gut at this stage is not complete although its rudimentary form is visible through the gut wall. Toward the seven-chaetiger stage, larval chaetae begin to appear in the first four segments, and the segments bear lateral pigmentation. At the eight-chaetiger stage, the larva grows and develops longitudinal muscle bands that it can use to move with. The next two stages include the development of epidermal papillae, the broadening of the head and the growth of palps. The gut is developed and the larva starts to rely on muscle rather than ciliary movement for locomotion. At 11 chaetigers, four adult eyes are present, and the structures of the adult head are almost complete. Adult notochaetae begin to develop anteriorly, and a pair of nephridia grow in the anterior part of the coelom. By 13 chaetigers, the cephalic cage begins to form. Unfortunately no further information is available on the later stage larvae or on when the transformation into juveniles occurs.

11.9 ACKNOWLEDGEMENTS M.E. Petersen would like to extend her sincere thanks to Heinz-Dieter Franke for the loan of a copy of Ute Völker’s unpublished doctoral dissertation on Ctenodrilus serratus Schmidt, and to Ute Völker for giving her unrestricted permission to use information from it. M.N. Halt was supported by the Australian Postgraduate Award and the CSIRO Marine Research Postgraduate Scholorship.

11.10 LITERATURE CITED Amor, A. 1994. Gametes, fertilization and development of Pherusa sp., an endolithic worm (Polychaeta, Flabelligeridae). Mémoires du Muséum Nationale d’Histoire Naturelle 162: 612. Banse, K. 1969. Acrocirridae n. fam. (Polychaeta Sedentaria). Journal of Fisheries Research Board of Canada 26: 2595-2620. Blake, J. A. 1975. 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. 2000. 1. Family Flabelligeridae Saint Joseph, 1894. Pp. 1-24. In J. A. Blake, B. Hilbig and P. H. Scott (eds), Taxonomic Atlas of the Benthic Fauna of the Santa Maria Basin and western Santa Barbara Channel. Volume 7 — The Annelida Part 4. Polychaeta: Flabelligeridae to Sternaspidae. Santa Barbara Museum of Natural History, Santa Barbara.

Cirratuliformia

#%

Blake, J. A. and Petersen, M. E. 2000. 3. Family Fauveliopsidae Hartman, 1971. Pp. 31-45. In J. A. Blake, B. Hilbig and P. H. Scott (eds), Taxonomic Atlas of the Benthic Fauna of the Santa Maria Basin and western Santa Barbara Channel. Volume 7 — The Annelida, Part 4. Polychaeta: Flabelligeridae to Sternaspidae., Santa Barbara Museum of Natural History, Santa Barbara. Bleidorn, C., Vogt, L. and Bartolomaeus, T. 2003. New insights into polychaete phylogeny (Annelida) inferred from 18S rDNA sequences. Molecular Phylogeny and Evolution 29: 279-288. Burnette, A. B., Struck, T. H., Halanych, K. M. (2005). Holopelagic Poeobius meseres (“Poeobiidae”, Annelida) is derived from benthic flabelligerid worms. Biological Bulletin 208: 213-220. Buzhinskaya, G. N. 1977. Flota vitjasi sp. n. a new deepwater pelagic species of the family Fauveliopsidae (Polychaeta, Annelida). Issledovaniya fauny morei, Zoologicheskii Institut Akademii Nauk USSR 20: 9-12. Buzhinskaya, G. N. 1996. Unique morphological features of certain deep-water pelagic polychaetes. Doklady Biological Sciences 347: 139-141. Caullery, M. and Mesnil, F. 1898. Les formes épitokes et l’évolution des cirratuliens. Annales de l’Universiteté de Lyon 39: 1-200, plates 1-6. Child, C. M. 1900. The early development of Arenicola and Sternaspis. Wilhelm Roux’ Archiv für Entwicklungsmechanik der Organismen 9: 587-722. Clark, R. B. and Olive, P. J. W. 1973. Recent advances in polychaete endocrinology and reproductive biology. Oceanography and Marine Biology Annual Review 11: 175-222. Dahl, E. 1955. On the morphology and affinities of the annelid genus Sternaspis. Lunds Universitets Årsskrift N.F. 51: 1-22. Dales, R. P. 1951. Notes on the reproduction and early development of the cirratulid Tharyx marioni (St. Joseph). Journal of the Marine Biological Association of the United Kingdom 30: 113-117. Dales, R. P. 1962. The polychaete stomodeum and the interrelationships of the families of the Polychaeta. Proceedings of the Zoological Society of London 139: 289328. Dales, R. P. 1963. Annelids. Hutchinson University Library, London, 200 pp. Day, J. H. 1967. A monograph on the Polychaeta of Southern Africa. Part II. British Museum of Natural History Publications: 656: 459-878. Dean, H. K. 1995. A new species of Raricirrus (Polychaeta, Ctenodrilidae) from wood collected in the Tongue of the Ocean, Virgin Islands. Proceedings of the Biological Society of Washington 108: 169-179. Dehorne, A. 1933. La schizométamérie et les segmentes tétragemmes de Dodecaceria caulleryi, n. sp. Bulletin Biologique de la France et de la Belgique 67: 298-326, plates 10-12. 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. 1927. Polychètes sedentaires. Addenda aux Errantes, Archiannelides, Myzostomaires. Faune de France, Paris 16: 494 pp. Fiege, D. and Buetfering, B. 2000. SEM-investigations on the morphology and anatomy of Sternaspis scutata (Ranzani, 1817) (Annelida: Polychaeta). Bulletin of Marine Science 67: 662-663. George, J. D. and Hartmann-Schröder, G. 1985. Polychaetes: British Amphinomida, Spintherida and Eunicida. Keys and notes for the identification of the species. Synopses of the British Fauna (New Series) 32: i-vii + 1-221.

#& Reproductive Biology and Phylogeny of Annelida George, J. D. and Petersen, M. E. 1991. The validity of the genus Zeppelina (Vaillant) (Polychaeta: Ctenodrilidae). Ophelia, Supplement 5: 89-100. Gibbs, P. E. 1971. A comparative study of reproduction cycles in four polychaete species belonging to the family Cirratulidae. Journal of the Marine Biological Association of the United Kingdom 51: 745-769. Gibson, P. H. 1981. Gametogenesis in the cirratulid polychaetes Dodecaceria concharum and D. caulleryi. Journal of Zoology, London 193: 355-370. Gibson, P. H. 1997. The nuchal organs and the nervous system of the head of four species of the cirratulid Dodecaceria. Polychaete Research No. 17. BioLine Publications, ISSN 0961320X. Goodrich, E. S. 1945. The study of nephridia and genital ducts since 1895. Quarterly Journal of Microscopical Science 86: 113-392. Gravier, C. 1923. La ponte et l’incubation chez les Annélides Polychètes. Annales des Sciences Naturelles (Zoologie et Paléontologie) (serie 10) 6: 153-247. Grube, A. E. 1855. Beschreibung neuer oder wenig bekannter Anneliden. Archiv für Naturgeschichte 21: 81-136. Grube, A. E. 1873. Die familie der Cirratulidien. Jahres-Bericht der Schlesiche Gesellschaft fuer vaterlandische Cultur, Breslau 50: 59-66 Hartman, O. 1955. Endemism in the North Pacific Ocean, with emphasis on the distribution of marine annelids, and descriptions of new or little known species. Pp. 39-60. In A. H. Foundation (ed.), Essays in the Natural Sciences in Honor of Captain Allan Hancock on the occasion of his birthday July 26, 1955, University of Southern California Press, Los Angeles. Hartman, O. 1967. Polychaetous annelids collected by the USNS Eltanin and Staten Island cruises, chiefly from Antarctic seas. Allan Hancock Monographs in Marine Biology 2: 1-387. Hartman, O. 1971. Abyssal polychaetous annelids from the Mozambique Basin off southeast Africa, with a compendium of abyssal polychaetous anneids from world-wide areas. Journal of the Fisheries Research Board of Canada 28: 14071428. Hartmann-Schröder, G. 1996. Annelida, Borstenwürmer, Polychaeta. Gustav Fischer, Jena, 648 pp. Heath, H. 1930. A connecting link between the Annelida and the Echiuroidea (Gephyrea Armata). Journal of Morphology 49: 223-249 Kudenov, J. D. 1976. Polychaeta from southeastern Australia 1. Acrocirridae Banse, 1969, from Victoria and New South Wales. Records of the Australian Museum 30: 137-151. Marion, A-F. 1974. Sure les Annélides du Golfe de Marseille. Comptes Rendus de L’Académie des Sciences, Paris, France. 79: 398-401. Mesnil, F. 1899. La position systématique des Flabelligériens St. Joseph (Chlorémiens Quatrefages) et des Sternaspiens. Zoologischer Anzeiger 22: 81-85. Mesnil, F. and Caullery, M. 1897. Sur la position systématique du genre Ctenodrilus Clap.; ses affinités avec les Cirratuliens. Compte rendu Hebdomadaire des Séances de l’Academie des Sciences 125: 542-544. Meyer, E. 1887. Studien über den köperbau der Anneliden. I-III. Mittheilungen aus der Zoologischen Station zu Neapel 7: 592- 741. Meyer, E. 1888. Studien uber den Körperbau der Anneliden. IV. Die Körperform der Serpulacean und Hermellen. Mittheilungen aus der Zoologischen Station zu Neapel 8: 462-662. Monticelli, F. S. 1907. Sessualita e gestazione nello Ctenodrilus serratus O. Schm. Congresso della Naturalista Italiani, Milano, Italy. 1906: 524-526.

Cirratuliformia

#'

Monticelli, F. S. 1910. Raphidrilus nemasoma Montic. nuovo Ctenodrilide del Golfo di Napoli (Revisione de’ Ctenodrilidi). Archivio Zoologico Unione Zoologica Italiana Napoli 4: 401-436. Núñez, J., Ocaña, O. and Brito, M. C. 1997. Two new species (Polychaeta: Fauveliopsidae and Nerillidae) and other polychaetes from the marine lagoon cave of Jameos Del Agua, Lanzarote (Canary Islands). Bulletin of Marine Science 60: 252-260. Okuda, S. 1934. The polychaete genus, Acrocirrus, from Japanese waters. Journal of the Faculty of Science, Hokkaido University. Ser. VI, Zoology 2: 197-209. Olive, P. J. W. 1970. Reproduction of a Northumberland population of the polychaete Ciratulus cirratus. Marine Biology 5: 259-173. Olive, P. J. W. 1971. Ovary structure and oogenesis in Cirratulus cirratus (Polychaeta). Marine Biology 8: 243-259. Olive, P. J. W. 1973. The regulation of ovary function in Cirratulus cirratus (Polychaeta). General and Comparative Endocrinology 20: 1-15. Olive, P. J. W. 1975. Germinal epithelium activity in polychaetes in relation to periodicity of reproduction. Pubblicazioni della Stazione Zoologica di Napoli 39 (Supplemento): 267-281. Olive, P. J. W. 1984. Environmental control of reproduction in Polychaeta. Pp. 17-38. In A. Fischer and H.-D. Pfannenstiel (eds). Polychaete Reproduction. Progress in Comparative Reproductive Biology. Fortschritt Zoologi 29. Peters, N. 1923. Über das Verhältnis des natürlichen zur künstlichen Teilung bei Ctenodrilus serratus (O. Schmidt). Zoologische Jahrbücher, Abteilung für Allgemeine Zoologie und Physiologie der Tiere. 40: 293-350. Petersen, M. E. 1999. Reproduction and development of the Cirratulidae (Annelida: Polychaeta). Hydrobiologia 402: 107-128. Petersen, M. E. 2000a. 11. Family Sternaspidae Carus, 1863. Pp 311-336. In J. A. Blake, B. Hilbig and P. H. Scott (eds) Taxonomic Atlas of the Benthic Fauna of the Santa Maria Basin and western Santa Barbara Channel. Volume 7 — The Annelida, Part 4. Polychaeta: Flabelligeridae to Sternaspidae. Santa Barbara Museum of Natural History, Santa Barbara, Petersen, M. E. 2000b. A new genus of Fauveliopsidae (Annelidae: Polychaeta), with a review of its species and redescription of some described taxa. Bulletin of Marine Science 67: 491-515. Petersen, M. E. and George, D. J. 1991. A new species of Raricirrus from northern Europe, with notes on its biology and a discussion of the affinities of the genus (Polychaeta: Ctenodrilidae). Ophelia, Supplement 5: 185-208. 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. Rietsch, M. 1882. Études sur le Sternaspis scutata. Annales des Sciences Naturelles (Zoologie et Paléontologie)(série 8) 13: 1-84. Riser, N. W. 1987. A new interstitial polychaete (Family Fauveliopsidae) from the shallow subtidal of New Zealand with observations of related species. Bulletin of the Biological Society of Washington 7: 211-216. Robbins, D. E. 1965. The biology and morphology of the pelagic annelid Poeobius meseres Heath. Journal of Zoology, London 146: 197-212. Rouse, G. and Fauchald, K. 1997. Cladistics and polychaetes. Zoologica Scripta 26: 139-204. Rouse, G. W. 1999a. Polychaeta, including Pogonophora and Myzostomida. Pp. 81124. In B. G. M. Jamieson (ed.), Reproductive Biology of Invertebrates, Volume IXB.

#  Reproductive Biology and Phylogeny of Annelida Progress in male gamete ultrastructure and phylogeny. Oxford and IBH Publishing Co., New Delhi. Rouse, G. W. 1999b. Trochophore concepts: ciliary bands and the evolution of larvae in spiralian Metazoa. Biological Journal of the Linnean Society 66: 411-464. Rouse, G. W. and Fauchald, K. 1997. Cladistics and polychaetes. Zoologica Scripta 26: 139-204. Rouse, G. W. and Pleijel, F. 2001. Polychaetes. Oxford University Press, London, 354 pp. Rouse, G. and Pleijel, F. 2003. Problems in polychaete systematics. Pp. 175-189. In E. Sigvaldadóttir, A. S. Y. Mackie, G. V. Helgason, D. J. Reish, J. Svavarsson, S. A. Steingrímsson and G. Guðmundsson (eds) Advances in Polychaete Research. Hydrobiologia 496. Sars, M. 1835. Beskrivelser og iakttagelser over nogle mærkelige eller nye i havet ved den Bergenske kyst levende dyr af polypernes, acelephernes, radiaternes, annelidernes of molluskernes classer, med en kort oversigt over de hidtil af forfattaren sammesteds fundne arter of deres forekommen, Bergen, 81 pp. 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. Sawada, N. 1984. Electron microscopical studies of spermatogenesis in polychaetes. Pp. 99-114. In A. Fischer and H-D. Pfannensteil (eds), Polychaete Reproduction. Gustav Fischer Verlag, Stuttgard, New York. Sokolow, I. 1911. Über eine neue Ctenodrilusart und ihre Vermehrung. Zeitschrift für wissenschaftliche Zoologie 97: 547-603. 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. 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. Vejdovsky, F. 1882. Untersuchungen über die Anatomie, Physiologie und Entwicklung von Sternaspis. Denkschriften der Akademie der Wissenschaften, Wien. 43: 33-90. Völker, U. 1997. Experimentelle und morphologische Untersuchungen zur geschlechtlichen Fortpflanzung des Polychaeten Ctenodrilus serratus Schmidt. Doctoral dissertation in Biology, University of Hannover, Germany. 115 pp. + 3 unnumbered (Unpublished dissertation, used with permission of the author). Wilfert, M. 1973. Ein Beitrag zur Morphologie, Biologie und systematischen Stellung des Polychaeten Ctenodrilus serratus. Helgoländer wissenschaftliche Meeresuntersuchungen 25: 332-346. Wilfert, M. 1974. Aphropharynx heterochaeta nov. gen. nov. spec.. ein neuer Polychaet aus der Familie Ctenodrilidae Kennel 1882. Cahiers de Biologie Marine 15: 495504.

12

CHAPTER

Sabellida Greg W. Rouse1,2 , Elena Kupriyanova2 and Eijiroh Nishi3

12.1 PHYLOGENY AND SYSTEMATICS Broad-scale classifications have previously grouped Sabellidae and Serpulidae under names such a Serpulacea, Sabelliformia and Serpulimorpha. Dales (1962) proposed that Serpulidae and Sabellidae be grouped as the order Sabellida and this name has subsequently become the most commonly used (e.g., Fauchald 1977; Pettibone 1982), although the membership of and hierarchical groupings within, have varied. Fauchald (1977) included five major taxa in Sabellida: Caobangiidae, Sabellongidae, Sabellidae, Serpulidae and Spirorbidae. Fitzhugh (1989) revised this to contain Sabellariidae, Sabellidae and Serpulidae. 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. Sabellariidae, Sabellidae Serpulidae and Oweniidae (Fig 12.1) were included by Rouse and Fauchald (1997) in Sabellida. Also, in a dramatic shift compared with traditional systematics Rouse and Fauchald (1997) placed Siboglinidae (Fig 12.1F) formerly outside polychaetes as Pogonophora and Vestimentifera, as part of Sabellida. In further analyses by Rouse (1999c; 2000), Oweniidae did not group with the remaining Sabellida, and Chaetopteridae (included here in as part of Spionida) did. Clearly further study is required. In addition, similarities between Sabellariidae and Pectinariidae (in Terebellida) also deserve further study. Sabellariidae has previously been considered as part of Terebellida (e.g., Fauchald 1977). Rousset et al. (2004) most recently assessed the placement of Siboglinidae, and so the composition of Sabellida, using a combination of morphological and molecular data. They found that sabellids and serpulids formed a clade, though the serpulids may actually belong inside Sabellidae. They also showed Oweniidae as sister group to Siboglinidae but neither this clade, nor Sabellariidae, formed a clade with sabellids and 1

South Australian Museum Nth Terrace, Adelaide. S.A. 5000 Australia Earth and Environmental Sciences University of Adelaide SA. 5005 Australia 3 Manazuru Marine Laboratory, Yokohama National University Iwa, Manazuru, Kanagawa 2590202, Japan. 2

#

Reproductive Biology and Phylogeny of Annelida

serpulids. Thus, the formulation of Sabellida used in this volume may well be artificial. The support values for the Rousset et al. (2004) study were quite low in general and further data are clearly needed.

Oweniidae Oweniidae has traditionally been placed as a group of Sedentaria but has rarely been explicitly associated with another polychaete taxon. Liwanow and Porfirjewa (1967) did suggest that Oweniidae was closely related to Pogonophora (now Siboglinidae), a suggestion later supported by Rouse and Fauchald (1997). A number of other authors have suggested that Oweniidae represents a primitive group of polychaetes, or should be excluded from the group altogether (see Minichev and Bubko 1992). Meyer and Bartolomaeus (1996) proposed that Oweniidae was sister group to a clade comprised Terebellida and Sabellida. Classifications since Dales (1962) isolated the group within it own order, until Rouse and Fauchald (1997) placed it in Sabellida. Relationships within Oweniidae have yet to be fully explored. Currently there would appear to be 28-37 or more accepted nominal species (Rouse and Pleijel 2001).

Sabellariidae Sabellariidae have also often been placed with Pectinariidae and Fauchald (1977) placed both these taxa in Terebellida. Knight-Jones (1981) and Fitzhugh (1989) both argued that the presence of uncini in a notopodial position (chaetal inversion) placed Sabellariidae with Sabellidae and Serpulidae. In their various analyses, Rouse and Fauchald (1997) generally found Sabellariidae to be the sister group to the clade formed by Sabellidae and Serpulidae. This was supported by the presence of uncini in a notopodia of some segments (referred to as chaetal inversion) and this placement is retained here. Further investigation into the systematic placement of Sabellariidae is certainly needed. The many similarities they share with Pectinariidae, especially in terms of larval metamorphosis and head structure, suggest Fauchald (1977) may have been right. Kirtley (1994) accepted 112 nominal species of Sabellariidae and placed them into 12 generic groupings. He divided these into two groups, Sabellariinae with three parathoracic segments and Lygdaminae with four parathoracic segments. There has been no cladistic analysis of Sabellariidae, but ideas of the evolutionary relationships within the group can be found in Gravier (1909) and Dales (1952).

Sabellidae Fitzhugh (1989) assessed the relationships amongst the taxa placed in Sabellida by Fauchald (1977), namely Caobangiidae, Sabellongidae, Sabellidae, Serpulidae and Spirorbidae. Fitzhugh (1989) found that recognition of the monotypic families Caobangiidae and Sabellongidae made Sabellidae paraphyletic and so placed Caobangia and Sabellonga

Sabellida

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Colour Figure

Fig. 12.1 Representatives of Sabellida. A. Lateral view of anterior end of Owenia fusiformis (Oweniidae) (Banyuls, France). B. Ventral view of anterior end of Idanthyrsus pennatus (Sabellariidae) (Sydney, Australia). C. Lateral view of Amphiglena mediterranea (Sabellidae showing two early embryos (arrow) laid in its tube (Ischia, Italy). D. Chitinopoma serrula (Serpulinae, Serpulidae) specimens on a shell (Iceland). Arrow indicates brood chamber filled with larvae. E. Romanchella quadricostalis (Spirorbinae, Serpulidae) removed from tube (Tasmania, Australia) showing brood of embryos attached by stalk. F. Lateral view of Oasisia sp. (Vestimentifera, Siboglindae), removed from its tube (Eastern Pacific). All images G. Rouse.

# " Reproductive Biology and Phylogeny of Annelida within Sabellidae. The first significant subdivision of Sabellidae was made by Rioja (1923) who divided it into Fabriciinae, Myxicolinae and Sabellinae, largely based on chaetal characters. The site of gamete formation and discharge has been used previously to separate sabellid genera into two groups. For instance, Zenkevitsch (1925) divided the Sabellidae into two tribes: Thoracogoneata and Abdominogoneata. In thoracogoneates, gametes develop in thoracic segments, with sperm (but not eggs) discharged anteriorly through a median sperm duct. Zenkevitsch (1925) based the thoracogoneate condition on a detailed study of Manayunkia baicalensis (Nusbaum), noting also the presence of a dorsal sperm duct in Fabricia stellaris. He postulated, erroneously however, that such genera as Amphicorina would also have a median sperm duct since these taxa also have thoracic gametes. Zenkevitsch (1925) also included Caobangia in Thoracogoneata, but with no explanation for this placement. All remaining sabellid genera were placed in the Abdominogoneata. While recognizing Thoracogoneata and Abdominogoneata, at least in terms of gamete distribution, Banse (1972) did note that some species in genera (e.g., Euchone) assigned to the Abdominogoneata have thoracic gametes. The distribution of gametes in Sabellidae was reassessed in detail by Rouse and Fitzhugh (1994) (see section 13.2). Fitzhugh’s (1989) cladistic analysis of Sabellidae resulted in significant changes to the classification of the group suggesting that only two subfamilies Fabriciinae and Sabellinae, could be accepted as monophyletic. Fitzhugh (1989) transferred a number of taxa previously included in Fabriciinae to Sabellinae. These included Chone, Desdemona, Euchone, Jasmineira and Fabrisabella. Myxicola was also placed into Sabellinae. The monophyly of Fabriciinae is supported by a number of synapomorphies and the most recent review of the group brings the total number of nominal species to more than 70 and more than 400 in Sabellinae (see Rouse and Pleijel 2001).

Serpulidae Several authors have speculated on the phylogeny of Serpulidae, or groups within this taxon. Caullery and Mesnil (1897) discussed the evolution of Spirorbinae and argued that they were a derived group of Serpulidae, based on a reduction of the number of thoracic chaetigers. In contrast, KnightJones (1981) proposed that Sabellidae and Serpulidae evolved from Spirorbinae-like ancestors. The argument by Knight-Jones (1981) implies that the recognition of Sabellidae makes Serpulidae paraphyletic. Smith (1991) rejected Knight-Jones’ (1981) hypothesis and instead argued that Sabellidae is paraphyletic, with Serpulidae the sister group to Sabellinae. Uchida (1978) proposed a series of complicated hypotheses for the evolution of Serpulidae and these have been criticised by ten Hove (1984). Ten Hove (1984) found Serpulidae (sensu lato) to be monophyletic and suggested a phylogeny of the group based on the morphology of the

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radiolar crown. He suggested that Filograninae is paraphyletic with taxa such as Josephella as potentially the sister group to Serpulinae. He also proposed that the sister group for Spirorbinae should be sought amongst taxa such as Pomatoceros. The analysis of Kupriyanova (2003) also suggests that spirorbins and serpulins constitute monophyletic groups, whereas filogranins are paraphyletic. Presently Serpulidae can reasonably be divided into Filograninae (though probably paraphyletic), Serpulinae (probably also paraphyletic) and Spirorbinae, with around 500 nominal species, though proper revision is required.

Siboglinidae Pogonophora was the name given by Johansson (1939) to an enigmatic group, usually found in deep sea, first described by Caullery (1914). Elevated to the status of phylum by Beklemishev (1944), pogonophores lack a gut lumen and their endodermal tissue is inflated and filled with symbiotic bacteria (Southward 1993). They can reach great size, as in the socalled Vestimentifera from hydrothermal vents and methane/sulfide seeps. The history of the treatment of this group of polychaetes as one or even two phyla is reviewed in Rouse and Fauchald (1995). Jones (1985) argued that, in spite of the many similarities between the two perceived groups of pogonophores, the form of the various coelomic cavities in the Vestimentifera was so different from the other Pogonophora that the taxonomic status of phylum was warranted. This has been shown to be erroneous by a series of studies (e.g. Bartolomaeus 1995; Rouse and Fauchald 1995; McHugh 1997). Pogonophora have now been reclassified as members of Sabellida (Rouse and Fauchald 1997) (but see Rouss et al. 2004) and the name originally formulated for the group, Siboglinidae Caullery, 1914 is now often used.

12.2 ANATOMY WITH REFERENCE TO THE REPRODUCTIVE SYSTEM Segmental organs were described in Owenia fusiformis by Gilson (1895). One or two pairs of funnels lie at the posterior end of chaetiger 6 and send a pair of epidermal ducts forward to exit at the front of the same segment. McIntosh (1917) denied they were nephridia, but could find trace of any other possible equivalent organs. Gilson (1895) suggested that the ducts he found served as an exit from the body for the gametes, though Watson (1901) subsequently showed that sperm or eggs actually are spawned though a pair of pores near the anus. Sabellariidae, Sabellidae, Siboglinidae and Serpulidae show a single anterior pair of segmental organs. Haswell (1884) was the first to notice that in addition to the anterior pair of segmental organs, there was a pair of ciliated ducts to the exterior in many of the abdominal chaetigers of serpulids, through which gametes were expelled. It seems likely that these are coelomoducts only, though a nephridial component has not been disproved (see Bartolomaeus 1999).

# $ Reproductive Biology and Phylogeny of Annelida Rouse and Fitzhugh (1994) identified three states for gamete distribution in Sabellida with reference to Sabellariidae, Sabellidae and Serpulidae only. Gametes limited to only abdominal chaetigers is found in all sabellariids (Eckelbarger 1984) and serpulids (e.g., Segrove 1941; Potswald 1967), and in most sabellin genera. This situation would arguably apply to Oweniidae but not to Siboglinidae where gametes are found in the trunk region only (see below). The occurrence of gametes in both thoracic and abdominal segments is found in several sabellin genera, e.g., Amphicorina, Chone, Euchone, and Jasmineira. The distribution of oocytes, however, tends to be more restricted than that of spermatids and/or sperm. In some instances for example, oocytes are limited to the thorax, but sperm are present in both regions (e.g., some Amphicorina, Desdemona aniara). Other genera, such as Chone and Euchone have eggs or sperm in both thoracic and abdominal segments (see Rouse and Fitzhugh 1994 for discussion). The condition of gametes being limited to only thoracic segments is only found in the Fabriciinae and the occurrence of the male dorsal sperm duct is also restricted to taxa to this subfamily (Rouse 1995a, 1996a). True gonads are absent in some serpulids (e.g. Hydroides dianthus, Ficopomatus enigmaticus) in either sex and the germ cells are produced by a germinal epithelium associated with the blood vessels in the intersegmental septa (Clark and Olive 1973; Dixon 1981). Distinct gonads have been described in Salmacina/Filograna (Faulkner 1929) and in Pomatoceros triqueter (Jyssum 1957). In several spirorbins, Potswald (1967b) noted the gonad as a discrete organ composed of germ cells arranged in two rows running the length of the abdominal segments. Apart from the anterior pair of excretory organs, the only other known segmental organs in Siboglinidae are the pair of gonoducts that lie in the trunk (arguably segment 1, see Rouse 2001). The ducts exit at the anterior border of the trunk in males and more posteriorly in females. Gardiner and Jones (1993) state that in Vestimentifera the male pores are in the vestimentum and the female pores are on the anterior part of the trunk. In contrast, Ivanov (1994) argues that the gonopores of both sexes in Vestimentifera lie on the trunk, as in other Siboglinidae. In frenulate and vestimentiferan siboglinids, the male reproductive system consists of a pair of testes lying ventrally along the median mesentery of the trunk, occupying most of the coelom that is not taken up by the trophosome. Anteriorly, each testis opens into a sperm duct that then runs to a gonopore opening (Bakke 1983; Gardiner and Jones 1993). In females a similar arrangement exist with paired ovaries giving rise to oviducts that open on the trunk surface. The testes and ovaries arguably represent coelomic pouches filled with developing gametes. In most siboglinids studied to date the sexes are of equal size. The exception is Osedax where there are dwarf males (Rouse et al. 2004). In Osedax the females have a large ovisac that gives rise to a single oviduct that runs along the trunk and extends as a tube out among the four palps. The detailed morphology of the ovary has yet to

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Fig. 12.2 Sperm and spermatids. A. Spermatids developing in tetrads of Circeis armoricana (Spirorbinae, Serpulidae) (Iceland). B. Spermatids developing in large clusters of Janua pagenstecheri (Spirorbinae, Serpulidae) (Iceland). C. Filograna/Salmacina sp. (Serpulidae), an intratubular brooder with sperm storage (Belize). D. Galeolaria caespitosa (Serpulidae), a broadcast spawner (Sydney, Australia). E. Paraprotis dendrova (Serpulidae), an extratubular brooder (Japan). Original. Abbreviations: a, acrosome; ax axoneme; m, mitochondrion; n, nucleus. All images G. Rouse.

# & Reproductive Biology and Phylogeny of Annelida be elucidated. The dwarf males of Osedax are basically larvae filled with sperm and spermatids. The sperm accumulate in an anterior duct (Rouse et al. 2004).

12.3 REPRODUCTION Information on reproduction in Oweniidae is largely restricted to Owenia fusiformis. This taxon is gonochoric and the gametes are spawned through freely into the water. Fertilization is external with eggs around 70 µm in diameter and the larvae pass through a normal trochophore phase before becoming a planktotrophic ‘mitraria’ (Watson 1901; Wilson 1932; Emlet and Strathmann 1994). In Europe, Owenia spawns in May to June each year, with females shedding on an average of 70 000 eggs. Individuals live for up to four years (Ménard et al. 1989; Gentil et al. 1990; Dauvin and Gillet 1991). All Sabellariidae studied to date appear to be gonochoric, broadcast spawners and produce planktotrophic larvae (Wilson 1991). Gametes are located throughout the coelom of the abdominal region. Males are whitish in the lateral abdominal region and females are orange or purplish. Wilson (1929, 1968a, 1968b, 1970, 1970) published a series of papers on European sabellariids, mainly on their larvae. He noted that in Sabellaria alveolata development in the plankton could take from six weeks to eight months. Eckelbarger (1975, 1976, 1977, 1978) has studied American sabellariids and their reproduction, particularly with regards to larvae and population aspects. Sabellariids are often gregarious and the settlement cues for larvae, and other aspects of larval life have been reviewed by Pawlik and Mense (1994). Sabellidae are mainly gonochoric, but some Sabellinae show simultaneous or protandric hermaphroditism. Depending on the taxon, simultaneous hermaphrodites can have eggs and sperm in the same segments or in different segments. All Fabriciinae thus far studied are gonochoric, continuous brooders of non-planktonic lecithotrophic larvae and females are capable of storing sperm. Sabellinae display a range of sexual reproductive modes, from broadcast spawning to ovoviviparity. Rouse and Fitzhugh (1994) have summarized information on reproduction and development in sabellids, with newer information available in Fitzhugh and Rouse (1999). Serpulidae show a range of sexuality patterns as well as reproductive and developmental mechanisms, encompassing asexual and sexual reproduction, gamete broadcast-spawning and sperm-storage, planktonic and parental benthic development (brooding) and feeding and non-feeding larval development. The reproduction and life history of serpulids (including spirorbins) was recently a subject of a comprehensive review by Kupriyanova et al. (2001). The sexes were traditionally considered to be separate in Serpulinae. However, hermaphroditism appears to be significantly under-reported as studies often reveal sequential (protandric) hermaphroditism with a very short intermediate stage in common fouling

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serpulins (e.g., Hydroides elegans, Pomatoceros triqueter, reviewed in Kupriyanova et al. (2001)). Unlike serpulins, all known spirorbins are simultaneous hermaphrodites. Their anterior abdominal segments contain eggs and the posterior segments contain male gametes. Simultaneous hermaphroditism is less common in serpulins and it seems to be a result of slower protandrous transition in small species such as Rhodopsis pusilla and Filograna/Salmacina. In latter genera, male gametes are usually in anterior segments and female are in posterior ones. Protandrous Salmacina incrustans can produce male gametes after emergence of female gametes (Vannini 1950). With the exception of Siboglinum poseidoni, all siboglinid taxa studied to date appear to be gonochoric (Gardiner and Jones 1993; Southward 1993). Males of all Siboglinidae, except Osedax, Sclerolinum and Vestimentifera, produce masses of leaf or spindle-shaped spermatophores that are spawned into the water. In the latter two taxa, it would appear that sperm are spawned freely or in unenclosed bundles (spermatozeugmata) (see review in Rouse 2001). Reproduction in Osedax has yet to be studied in detail. Hilário et al. (2005) recently demonstrated sperm storage and internal fertilization in vestimentiferans.

12.4 EGGS Nothing is known about oogenesis in Oweniidae. Oogenesis has been studied in only a few sabellids (Giangrande et al. 2000). In all cases it would appear that oogenesis is extraovarian and autosynthesis of yolk occurs. Coelomocytes are present in the coelom that decline in number during oogenesis (Dales 1961) and these probably store reserves for vitellogenesis. In Sabellariidae there have been some very detailed studies of oogenesis by Eckelbarger, particularly on Phragmatopoma lapidosa (Eckelbarger 1979). In P. lapidosa, vitellogenesis occurs within the ovaries before the eggs are released into the coelom. Two kinds of yolk are formed, one via autosynthesis and the other via endocytosis from the genital blood vessel (heterosynthesis). Estimates of egg numbers of sabellids were made by Rouse and Fitzhugh (1994) and ranged from a minimum of one in Caobangia (an estimate based on the assumption that only one egg can be fertilised at a time and then lie in the larval duct, (see Jones 1974) or 2-4 in many fabriciins, to nearly 666 000 (mean for Eudistylia vancouveri). Egg number was based on an estimate of fecundity from one spawning event, not lifetime fecundity since such data are not available for any sabellid. Estimates are also based on the number of mature or nearly mature oocytes at a given time, not total volume of oocytes. These values are, of course, significantly different in continuous breeders as opposed to iteroparous species. The smallest mature egg size reported for a sabellid to date is that of Branchiomma luctuosum (Sordino and Gambi 1992), with a diameter of 120 µm. The largest eggs reported were 600 µm by 400 µm for Amphiglena marita

#! Reproductive Biology and Phylogeny of Annelida by Chlebovitsch (1959), and 500 µm in diameter for Potamilla torelli by Rouse and Fitzhugh (1994). As in sabellids, developing gametes of serpulids are released into the coelom. In both sexes there is a lack of synchrony in the way gametes are produced, with all but the pre-spawning mature and recently spawned individuals containing gametocytes in different stages of development (Kupriyanova et al. 2001). Cytological events and ultrastructural details of oogenesis have been studied in serpulins Pomatoceros (Jyssum 1957) and Hydroides norvegicus (Nordback 1956) and in several spirorbins (e.g., King et al. 1969; Potswald 1967b, 1969). Egg sizes show a wide range of variation (reviewed in Kupriyanova et al. 2001). Serpulins they range from 40-45 µm (Pomatoleios kraussi, Hydroides ezoensis) to 180-200 µm (Chitinopoma serrula). Egg size in spirorbins ranges from 80 µm in Neodexiospira foraminosa to 230 µm in Pileolaria militaris. Species with eggs < 80 µm in diameter generally have planktotrophic larvae, larvae of those with larger eggs are usually lecithotrophic. The details of oogenesis in Siboglinidae have yet to be fully studied. Oogenesis is intraovarian and continuous, but as noted above the ovary is in a sense a coelomic compartment in siboglinids. Oocytes are formed at the anterior end of the ovaries, released into the ovarian coelom and pushed posteriorly as more oocytes are formed. In frenulates, there are no nurse cells and oogenesis appears to be solitary and autosynthetic (Southward 1975). In vestimentiferans follicle cells are seen in Riftia pachyptila associated with vitellogenic oocytes, though a nutritive role has yet to be established (Gardiner and Jones 1993). Eggs sizes range dramatically in siboglinids, with vestimentiferans having smaller spherical eggs in the range of 105 µm when spawned (Carey et al. 1989) for Riftia pachyptila. The eggs of Osedax are slightly elliptical with dimensions of 140-150 µm by 115-125 µm (Rouse et al. 2004). In frenulates such as Polybrachia and Galathealinum the eggs can be 150-160 µm in diameter, while in Oligobrachia they can be 400 µm. The largest eggs known in siboglinids are the massive elliptical eggs of Siboglinum caulleryi that are 650 µm long and 130 µm across (Southward 1975).

12.5 SPERM Little is known about spermiogenesis or sperm in Oweniidae. Rouse (1988) provided a description of the ultrastructure of an Owenia sperm from Australia. In Sabellariidae, the ultrastructure of spermiogenesis in Phragmatopoma lapidosa has been studied by Eckelbarger (1984). Spermatids develop in tetrads and mature sperm have spherical heads typical of external fertilizers, but have very elongate acrosomes. Eckelbarger (1984) postulated that the complicated acrosome was a result of the elaborate egg envelope in sabellariids. Other less detailed studies on sabellariid sperm have also been performed (Pasteels 1965; Kopp 1985; Franzén and Rice 1988).

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Fig. 12.3 Fertilization and brooding. A. In vitro fertilization of Galeolaria caespitosa eggs. B. Embryos adhering to tube wall of Circeis spirillum (Spirorbinae, Serpulidae) (Iceland). C. SEM showing larvae of Paraprotis dendrova (Serpulidae) (Japan) attached to the parent via a modified radiole (arrow). Images: A. E. Kupriyanova, B. G. Rouse, C. E. Nishi.

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Sabellidae is the best studied of all polychaete groups in terms of sperm ultrastructure, with descriptions of sperm of more than 40 species published to date. Spermiogenesis can be in tetrads or clusters of spermatids attached to a cytophore. While all Fabriciinae show the pattern of spermiogenesis with hundreds of spermatids connected to a cytophore, the situation in Sabellinae is more variable (Rouse and Fitzhugh 1994; Rouse 1999a). Across most Sabellinae, including all plesiomorphic forms, spermiogenesis occurs in tetrads of synchronously developing spermatids. However, in some of the more apomorphic sabellins such as Potamilla, spermatids form clusters of hundreds, joined by a central cytophore, as in Fabriciinae (Rouse and Fitzhugh 1994). Also in the derived Amphiglena and Laonome clade, Rouse and Fitzhugh (1994) described them as having spermiogenesis with sperm developing simultaneously in clusters of < 100 cells. Mature sperm have spherical or cylindrical nuclei and the mitochondria may be spherical or elongate wrapping around the axoneme to form a complex midpiece (Rouse 1992a, 1995a, 1999a). A complex subacrosomal space is found in a number of sabellin species such as Pseudopotamilla reniformis, Sabella pavonina and S. spallanzanii that presumably have external fertilization (Kryvi and Graebner 1975; Chughtai 1986; Giangrande et al. 2000). The reasons for this have yet to be elucidated. Rouse (1995a) described the sperm ultrastructure of 20 fabriciin sabellids and assessed their utility for systematics. He found that they shared a number of apomorphic features such as 1. A thick glycocalyx over the plasma membrane. 2. A distinctive nuclear projection, with an anterior thickening of the nuclear membrane. 3. A thickened, spiralling ridge of nuclear membrane. 4. An extra-axonemal sheath. 5. A unique sheath of mitochondrial material in the midpiece. Within Fabriciinae species differ markedly in sperm structure. This variability involves acrosome morphology; the structure of the sperm nucleus and nuclear projection; the spiral of thickened nuclear membrane; the structure of the extra-axonemal sheath; the mitochondrial sheath. The variability of sperm structure amongst the basal sabellin taxa is considerable (see Rouse 1992a) and further work is needed to resolve relationships among them to understand how sperm shape has evolved in sabellids. Sperm shape varies remarkably within Serpulidae (Rouse 1999b, 2005) and differences in sperm morphology reflect different modes of fertilisation or sperm transfer (Fig 12.2). Sperms that are characterised by a spherical to conical head (ect-aquasperm) are typical for broadcast-spawning serpulids such as Pomatoleios kraussi, Spirobranchus corniculatus, Protula globifera, Placostegus tridentatus, Serpula vermicularis, Hydroides norvegicus, H. dianthus, H. ezoensis, H. fusicola, and H. elegans, Floriprotis sabiuraensis and Galeolaria caespitosa (reviewed by Kupriyanova et al. (2001). Spermatogenesis, which can result in spermatids developing in tetrads (Fig. 12.2A) or in large clusters (Fig 12.2B) has been studied in serpulins Chitinopoma serrula (Franzén 1982), Hydroides norvegicus, Placostegus tridentatus, Protula globifera, and Serpula vermicularis (Franzén 1956), Hydroides diramphus (Mona et al.

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1994) and in spirorbins Simplaria potswaldi, Paradexiospira (Spirorbides) vitrea, Circeis spirillum, Protolaeospira eximia (Potswald 1967a, 1967b) and Spirorbis spirorbis (Picard 1980). Sperm with an elongated head (Fig. 12.2C) and midpiece (ent-aquasperm) are known for brooding serpulins such as Salmacina dysteri, Chitinopoma serrula, and Rhodopsis pusilla and for spirorbins. The elongated shape of the nucleus and mitochondria are attributed to sperm storage prior to fertilization. The brooding serpulin Paraprotis dendrova is unusual because its sperm (Fig 12.2E), although slightly elongated, is very similar to that of broadcasting serpulins. However, this species broods larvae on a modified radiole and lacks spermathecae (Nishi and Yamasu 1992c). It is likely in this species eggs attached to the brooding radiole are fertilized by the sperm freely released into water column. With regard to Siboglinidae, 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 1993) and Ridgeia piscesae (see Southward and Coates 1989) and is essentially similar in all three taxa. Spermiogenesis in general for the group has been reviewed several times (Bakke 1983; Gardiner and Jones 1993; Southward 1993). Spermatids develop in large morulae attached to a central cytophore. 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. Microtubules are present around the nucleus until very late in development. In Siboglinum ekmani they are involved in nuclear elongation (Franzén 1973) and are possibly involved in the final positioning of the acrosome in R. piscesae (Southward and Coates 1989). The acrosome of S. ekmani is helical and lies at the apex of the nucleus (Franzén 1973). Gardiner and Jones (1985) described nearly mature sperm of Riftia pachyptila as having a 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). The 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).

12.6 MATING AND FERTILIZATION Both Oweniidae and Sabellariidae are only known to have broadcast spawning and external fertilization (Rouse and Pleijel 2001). Sabellidae and Serpulidae on the other hand, may be broadcast spawners, brooders that brood their larvae either within the tube, attached to the radiolar crown or in a jelly mass or ring at the mouth of the tube. All Siboglinidae appear to

#!" Reproductive Biology and Phylogeny of Annelida

Fig. 12.4 Larvae. A. Differential interference contrast micrograph of Spirobranchus giganteus trochophore showing complete gut (Belize). B. SEM of Filograna/Salmacina sp. (Serpulidae) larva removed from tube (Belize). Shortly after this the larva would have left the tube to settle. C. Larva of Janua pagenstecheri (Spirorbinae, Serpulidae) (Iceland) removed from opercular brood chamber. Note two shell glands packed with tube precursor that allow the metamorphosing worm to secrete a calcareous tube immediately on settling. Original. Abbreviations: cg, calcareous tube gland; g, gut. All images G. Rouse.

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#!#

have internal fertilization but either release the eggs into the water or brood them in the tube. Within Sabellidae, adult females of all Fabriciinae studied to date brood larvae within their tubes, and the presence of paired spermathecae in the radiolar crown or peristomium has been described in a number of species. These spermathecae range in complexity from simple pits to elaborate ducts with several elements (Rouse 1996c). However, all are blind-ending and fertilization presumably occurs when the sperm are ejected from the spermathecae into the female’s tube to meet the spawned eggs. Interestingly, spermathecae have not been found in a number of other fabriciins and in these cases sperm are simply stored in epidermal cells in the radiolar crown or buccal region (Rouse 1996c). The spermathecae of Caobangia abbotti (a sabellid of uncertain affinity according to Rouse and Fitzhugh 1994) open into the coelomic cavity (Jones 1974). This then allows for internal fertilization and development of larvae within the body of the adult followed by birth through a ‘larval duct’ (Jones 1974). Plesiomorphic sabellins, such as Amphicorina have a reproductive mode similar to that of Fabriciinae, though homologies of reproductive structures are still unclear (Rouse 1992b). Spermathecae have been described in only three sabellin genera: Amphicorina, Amphiglena and Terebrasabella (Rouse 1992b; Rouse and Gambi 1998b; Simon and Rouse 2005). The spermathecae in Amphicorina may be simple blind ducts or divided into two distinct regions (Rouse 1992b). The spermathecae of Amphiglena spp. are narrow convoluted ducts or simple sacs (Rouse and Gambi 1998b). Terebrasabella is unusual in having a single spermatheca. It is blind-ending and runs from below the radiolar crown along the ventral epidermis to at least the first chaetiger (Simon and Rouse 2005). Intratubular brooding is uncommon in the more apomorphic Sabellinae, whereas broadcast spawning (e.g., Eudistylia, Schizobranchia), or extratubular brooding (e.g., Perkinsiana) is the norm. Exceptions to this trend include taxa such as Amphiglena, Potamilla and Terebrasabella, which show intratubular brooding. Sexual reproduction in Sabellidae was reviewed by Rouse and Fitzhugh (1994). More recent studies include those on Amphiglena, Perkinsiana, Sabella and Terebrasabella (Rouse 1996c; Rouse and Gambi 1998a; Fitzhugh and Rouse 1999; Gambi and Patti 1999; Gambi et al. 2000; Giangrande et al. 2000; Simon and Rouse 2005). Many serpulids are broadcast-spawners releasing gametes into the water column where fertilisation and subsequent development occur. The gametes are expelled through segmental organs and are delivered by ciliary beating in the faecal groove to the tube opening. Fertilisation success and factors affecting it were not studied for serpulids until very recently. Fertilisation in Hydroides elegans is successful over a broad range of sperm concentration and usually > 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

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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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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.

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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.

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##$ 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.

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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.

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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,

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Fig. 13.1 contd

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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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Reproductive Biology and Phylogeny of Annelida

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

#'" Reproductive Biology and Phylogeny of Annelida

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

$

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

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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.

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$!& 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).

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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

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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.

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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

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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.

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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.

Problematic Annelid Groups

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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

$#" Reproductive Biology and Phylogeny of Annelida

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.

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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.

Problematic Annelid Groups

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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.

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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.

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%$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