Reproduction and Development in Annelida 9780429584480, 0429584482

Table of contents :
Content: Cover --
Title --
Copyrights --
Preface to the Series --
Preface --
Acknowledgements --
Contents --
Chapter 1. Introduction --
1.1 Annelidan Science --
1.2 Species and Structural Diversity --
1.3 Geographic Distribution --
1.4 Gutless Oligochaetes --
1.5 Hydrothermal Vents and Cold Seeps --
1.6 Energy Budget --
1.6.1 Osmotrophism --
1.6.2 Anaerobiosis --
1.7 Life Span and Generation Time --
1.8 Gametogenesis --
1.9 Reproductive Modes and Dispersal --
1.10 Fertilization Site and Success --
1.11 Annelidan Larvae --
1.12 Defense and Parental Care --
1.13 Metamorphosis and Settlement --
Chapter 2. Sexual Reproduction --
Introduction --
2.1 Reproductive Systems --
2.2 Gonochorism --
2.2.1 Sex Ratio --
2.2.2 Ovary Somatic Index --
2.4 Parthenogenesis --
2.4.1 Parthenogenic Types --
2.4.2 Parthenogenic Levels --
2.4.3 Ploidy Levels --
2.3 Hermaphroditism --
2.5 Fecundity --
2.5.1 Sexuality --
2.5.2 Oogenic Anlage --
2.5.3 Egg Size --
2.5.4 Body Size --
2.6 Poecilogony and Dispersal --
2.7 Mating Systems --
2.7.1 Simultaneous Hermaphrodites --
2.7.2 Sequential Hermaphrodites --
2.7.3 Labile Gonochorics --
Chapter 3. Regeneration --
Introduction --
3.1 Regeneration and Reproduction --
3.2 Incidence and Prevalence --
3.3 Regenerative Process --
3.3.1 Wound-healing --
3.3.2 Cell Migration --
3.3.3 Blastema and Differentiation --
3.3.4 Segmentation and Reorganization --
3.4 Anterior vs Posterior --
Chapter 4. Asexual Reproduction --
Introduction --
4.1 Obligate Cloners? --
4.2 Incidence and Prevalence --
4.3 Observations and Characteristics --
4.4 Architomy --
4.5 Naidu's Monograph --
4.6 Paratomy --
4.7 Restoration of Sexual Reproduction --
4.8 Clonal Stem Cells --
Chapter 5. Epitoky --
Introduction --
5.1 Types and Characteristics --
5.2 Epigamy --
5.3 Schizogamy --
5.4 Vertical Migration --
5.5 Swarming Phenomenon --
5.6 Pheromones and Spawning. Chapter 6. Sex Determination --
Introduction --
6.1 Karyotype and Heterogamety --
6.2 Gametic Compatibility --
6.3 Sex Ratio and Variations --
Chapter 7. Sex Differentiation --
Introduction --
7.1 Endocrine Regulation --
7.1.1 Nereidids --
7.1.2 Nephtyids --
7.1.3 Phyllodocids --
7.1.4 Arenicolids --
7.2 Differentiation and Lability --
7.2.1 Dorvilleids --
7.2.2 Syllids --
7.3 Pollutants and Reproduction --
7.4 Vectors and Borers --
Chapter 8. Vermiculture --
Introduction --
8.1 Characteristics and Features --
8.2 Candidate Species --
8.2.1 Growth --
8.2.2 Reproduction --
8.2.3 Polyploids and Parthenogens --
Chapter 9. Summary and New Findings --
Chapter 10 . References --
Author Index --
Species Index --
Subject Index --
Author's Biography.

Citation preview

Series on Reproduction and Development in Aquatic Invertebrates Volume 4

Reproduction and Development in Annelida

T. J. Pandian Valli Nivas, 9 Old Natham Road Madurai-625014, TN, India

p, p,

A SCIENCE PUBLISHERS BOOK A SCIENCE PUBLISHERS BOOK

Cover page: Representative examples of annelid species. For more details, see Figure 1.1

CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2019 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed on acid-free paper Version Date:  International Standard Book Number-13:  (Hardback)

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Preface to the Series

Invertebrates surpass vertebrates not only in species number but also in diversity of sexuality, modes of reproduction and development. Yet, we know much less of them than we know of vertebrates. During the 1950s, the multivolume series by L.E. Hyman accumulated bits and pieces of information on reproduction and development of aquatic invertebrates. Through a few volumes published during the 1960s, A.C. Giese and A.S. Pearse provided a shape to the subject of Aquatic Invertebrate Reproduction. Approaching from the angle of structure and function in their multi-volume series on Reproductive Biology of Invertebrates during the 1990s, K.G. Adiyodi and R.G. Adiyodi elevated the subject to a visible and recognizable status. Reproduction is central to all biological events. The life cycle of most aquatic invertebrates involves one or more larval stage(s). Hence, an account on reproduction without considering development shall remain incomplete. With the passage of time, publications are pouring through a large number of newly established journals on invertebrate reproduction and development. The time is ripe to update the subject. This treatise series proposes to (i) update and comprehensively elucidate the subject in the context of cytogenetics and molecular biology, (ii) view modes of reproduction in relation to Embryonic Stem Cells (ESCs) and Primordial Germ Cells (PGCs) and (iii) consider cysts and vectors as biological resources. Hence, the first chapter on Reproduction and Development of Crustacea opens with a survey of sexuality and modes of reproduction in aquatic invertebrates and bridges the gaps between zoological and stem cell research. With capacity for no or slow motility, the aquatic invertebrates have opted for hermaphroditism or parthenogenesis/polyembryony. In many of them, asexual reproduction is interspersed within sexual reproductive cycle. Acoelomates and eucoelomates have retained ESCs and reproduce asexually also. However, pseudocoelomates and haemocoelomates seem not to have retained ESCs and are unable to reproduce asexually. This series provides possible explanation for the exceptional pseudocoelomates and haemocoelomates that reproduce asexually. For posterity, this series intends to bring out six volumes. August, 2015 Madurai-625 014

T. J. Pandian

Preface

Annelids are known for the unique spectacular epitoky, regeneration and clonal reproduction. These features have attracted attention more from academic interest. An objective of this book is to elevate annelids from academic interest to economic importance. Many books are authored or edited on annelids but are limited to a taxonomic group or a specific theme like phylogeny. This book is a concise, informative elucidation of reproduction and development in annelids covering from Aeolosoma viride to Zeppelina monostyla. The book is structured in nine chapters. In view of their importance, two chapters are devoted to regeneration and clonal reproduction, a chapter on epitoky and another on vermiculture. Some 81, 12 and 7% annelids are marine, freshwater and terrestrial inhabitants. Comprising 17,000 species, annelids are found mostly in aquatic habitats but a few in terrestrial habitats. Vertically distributed between 4,900 m depth and 2,000 m altitude, they are also found in unusual habitats like hydrothermal vents, subterranean aquatic system and migrating between the nutrient-rich anoxic and oxic zones in the sediments. For the first time, information on gutless oligochaetes and polychaetes, osmotrophism and anaerobiosis in some annelids is highlighted. In the absence of exoskeleton to escape predation, 42–47% polychaetes brood their eggs. The second chapter deals with sexual reproduction. Polychaetes are gonochores. But clitellates are hermaphrodites characterized by internal fertilization and laying cocoon enclosing a few eggs. Updating has revealed the incidence of hermaphroditism in 207 polychaete species from 27 families. Hence, only 74% of annelids are gonochores. A directory is generated and lists 75 parthenogenic annelid species, of which 56 are earthworms. The first estimate has revealed the incidence of external fertilization in 54% polychaetes. The existence of poecilogony with triple morphs and simultaneous sex change between mating partners are projected. Devoted to regeneration and clonal reproduction, the third and fourth chapters have brought to light a whole range of new findings. In oligochaetes, inadequate reserves in the chloragogue temporally separate the incidence of regeneration and reproduction. However, the sedentary polychaetes, which do not possess the chloragogue or its equivalent, undertake them together,

vi  Reproduction and Development in Annelida

of course, at the cost of reduced reproduction. The number of species capable of anterior, posterior and anterior cum posterior regeneration is 149, 206 and 143, respectively. When these numbers are considered as fractions of 13,012 polychaete and 3,175 oligochaete species, the percentage values (1.57, 1.80, 1.42) indicate that the potency is 1.5–2.0 times more prevalent in oligochaetes than the respective ones (0.88, 1.22, 0.85) in polychaetes. The earlier loss of the regenerative potency in ‘older’ anterior segments than in ‘young’ posterior segments located adjacent to the generative pygidium may be a reason for the observed less prevalence of anterior regeneration. Clonal reproduction can sustain a species for > 30–60 years. When stressed or induced, sexual reproduction is manifested, as Primordial Germ Cells (PGCs) are transmitted up to 1,000–3,000 clonal generations. Abundant food supply and low density trigger clonal reproduction but not intense predation; for example 290 tubiculous sabellids species, only 17 are cloners. Of 100 and odd annelid families, the incidence of clonal reproduction occurs only in 12 polychaete families and 5 oligochaete families. It ranges from 2% of spionids to 54% of naidids. Further, it occurs in 79 polychaete species but to as many as 111 oligochaete species. Clonal reproduction is considered to have been derived from regeneration. However, this view is not correct, for (i) in as many as 111 oligochaete species, cloning does obligatorily require the presence of neoblasts and (ii) even with anterior cum posterior regenerative potency, 34 out of 63 polychaete species, do not reproduce clonally. In most polychaetes, the stem cells responsible for cloning are located at the posterior end and also mid-body in a few. The epitokes are divided into semelparous epigamics and iteroparous schizogamics. For the first time, a directory is documented listing epigamy in 61 species from 12 families and schizogamy in 45 syllid species. Again for the first time, the assembled information on vertical distance traveled by 28 epitokous species reveals that the larger glycerids, nereidids and eunicids use muscular energy to climb up < 50 m but the smaller phyllodocids and ctenodrilids may engage reduced buoyancy to climb the vertical distance of up to 4,000 m. The sixth chapter deals with sex determination by genes harbored on chromosomes. Karyotyping and breeding experiments have found heterogametism in six polychaete species only. A directory is assembled for the chromosome numbers in annelids. By selective fertilization of large eggs by X-carrying sperm, Dinophilus gyrociliatus have nullified the chromosomal mechanism of sex determination. In Capitella capitata, expression of W gene(s) is stable but that in Z chromosome is labile resulting in generation of phenotypic ZZ hermaphrodites and females. Our understanding of endocrine sexualization in syllids and regulation of reproductive cycles in others is based on temperate polychaetes alone. A dozen neuroendocrines/hormones secreted mostly by the ‘brain’ regulate the reproductive cycle.

Preface  vii

The ninth chapter on vermiculture (i) emphasizes the need for information on growth and reproduction in cultivable species, (ii) considers parthenogens and cloners, as they do not have adequate genetic diversity and cloners increases the number but may not the biomass and (iii) recognizes ‘layers’ as distinct from ‘brooders’. There is an urgent need for research input to harvest tubificids and naidids at appropriate intervals, as it may reduce the input of nitrogen fertilizers in ricefields. The fastest growing earthworms, nereidids, tubifex and pot worms are recommended as cultivable species. For the first time, the fast growing Branchiura sowerbyi fed on waste paper immersed in water is identified as potential candidate species for vermiculture. This book is a comprehensive synthesis of 737 publications carefully selected from widely scattered information from 237 journals and other literature sources. The holistic approach and incisive analysis have led to several new findings and ideas related to reproduction and development of annelids. Hopefully, the book serves as a launch-pad to further advance our knowledge on annelids. July, 2018 Madurai-625 014

T. J. Pandian

Acknowledgements

It is with pleasure that I thank Drs. P. Murugesan and E. Vivekanandan for critically reviewing parts of the manuscript of this book and for offering valuable suggestions. In fact, I must confess that I am only a visitor to the theme of this book. However, a couple of joint publications with my student the late Dr. M. Peter Marian, my earlier book in this series and editorial service on annelidan energetics (Pandian, 1987, Animal Energetics, Academic Press) have emboldened me to author this book. I wish to thank my former students and associates Drs. Premkumar David (USA), G. Kumaresan, V. Mohan, P. Murugesan, S. Sudhakar and Prof. W. Westheide (Germany) for by providing copies of some publications. The manuscript of this book was prepared by Mr. T.S. Balaji, M.Sc. and I wish to thank him profusely for his competence, patience and co-operation. Firstly, I wish to thank many authors/publishers, whose published figures are simplified/modified/compiled/redrawn for an easier understanding. To reproduce original figures from published domain, I gratefully appreciate the permission issued by Dr. S.M. Mandaville. For permissions issued to reproduce original figures from dissertations/protocols of his students, I remain thankful to Dr. S. Sudhakar, who has also provided me his consent to reproduce unpublished figures. Dr. P. Murugesan has kindly provided me a hand drawn figure. For advancing our knowledge in this area by their rich contributions, I thank all my fellow scientists, whose publications are cited in this book. July, 2018 Madurai-625 014

T. J. Pandian

Contents

Preface to the Series iii Preface v Acknowledgements ix 1. Introduction 1.1 Annelidan Science 1.2 Species and Structural Diversity 1.3 Geographic Distribution 1.4 Gutless Oligochaetes 1.5 Hydrothermal Vents and Cold Seeps 1.6 Energy Budget 1.6.1 Osmotrophism 1.6.2 Anaerobiosis 1.7 Life Span and Generation Time 1.8 Gametogenesis 1.9 Reproductive Modes and Dispersal 1.10 Fertilization Site and Success 1.11 Annelidan Larvae 1.12 Defense and Parental Care 1.13 Metamorphosis and Settlement

1 1 5 8 14 16 19 21 25 27 31 35 42 46 48 51

2. Sexual Reproduction Introduction 2.1 Reproductive Systems 2.2 Gonochorism 2.2.1 Sex Ratio 2.2.2 Ovary Somatic Index 2.3 Hermaphroditism 2.4 Parthenogenesis 2.4.1 Parthenogenic Types 2.4.2 Parthenogenic Levels 2.4.3 Ploidy Levels

53 53 53 57 57 59 61 65 66 67 68

xii  Reproduction and Development in Annelida

2.5 Fecundity 2.5.1 Sexuality 2.5.2 Oogenic Anlage 2.5.3 Egg Size 2.5.4 Body Size 2.6 Poecilogony and Dispersal 2.7 Mating Systems 2.7.1 Simultaneous Hermaphrodites 2.7.2 Sequential Hermaphrodites 2.7.3 Labile Gonochorics

70 71 73 73 76 84 87 88 89 90

3. Regeneration Introduction 3.1 Regeneration and Reproduction 3.2 Incidence and Prevalence 3.3 Regenerative Process 3.3.1 Wound-healing 3.3.2 Cell Migration 3.3.3 Blastema and Differentiation 3.3.4 Segmentation and Reorganization 3.4 Anterior vs Posterior

92 92 93 98 105 106 106 111 112 119

4. Asexual Reproduction Introduction 4.1 Obligate Cloners? 4.2 Incidence and Prevalence 4.3 Observations and Characteristics 4.4 Architomy 4.5 Naidu’s Monograph 4.6 Paratomy 4.7 Restoration of Sexual Reproduction 4.8 Clonal Stem Cells

125 125 125 127 130 135 141 144 148 149

5. Epitoky Introduction 5.1 Types and Characteristics 5.2 Epigamy 5.3 Schizogamy 5.4 Vertical Migration 5.5 Swarming Phenomenon 5.6 Pheromones and Spawning

153 153 154 155 160 162 165 167

Contents  xiii

6. Sex Determination Introduction 6.1 Karyotype and Heterogamety 6.2 Gametic Compatibility 6.3 Sex Ratio and Variations

170 170 170 173 175

7. Sex Differentiation Introduction 7.1 Endocrine Regulation 7.1.1 Nereidids 7.1.2 Nephtyids 7.1.3 Phyllodocids 7.1.4 Arenicolids 7.2 Differentiation and Lability 7.2.1 Dorvilleids 7.2.2 Syllids 7.3 Pollutants and Reproduction 7.4 Vectors and Borers

179 179 180 180 182 183 183 184 184 186 190 194

8. Vermiculture Introduction 8.1 Characteristics and Features 8.2 Candidate Species 8.2.1 Growth 8.2.2 Reproduction 8.2.3 Polyploids and Parthenogens

196 196 196 202 202 211 215

9. Summary and New Findings

218

10 . References

224

Author Index 257 Species Index 264 Subject Index 275 Author’s Biography

277

1 Introduction

1.1  Annelidan Science* Annelids are bilaterally symmetrical, triploblastic, schizocoelomatic, metamerically segmented vermiform invertebrates. Their segmented body has a well-developed ladder-like central nervous system with a bi-lobed cereberal ganglion and sense organs, a closed blood-vascular system, coelom, an excretory system and a fairly well developed endocrine system. Among aquatic fauna, they are a fascinating taxon displaying (i) epitoky, a spectacular unique phenomenon involving transformation from benthic to meroplanktonic reproductive morphism, (ii) osmotrophism displayed by the gutless tubificids acquiring cent percent nutrients across the body surface from ambient sea water, (iii) partial (tubificids) and complete (vestimentiferans) chaemoautotrophism by engaging symbiotic microbes to draw energy, (iv) poecilogony, another unique feature shared only by some gastropods (Pandian, 2017), (v) metamerism, the most distinguishing feature of annelids and (vi) regeneration, which may be followed by bidirectional (a genet divided into two ramets) and multidirectional (a genet divided into multiple ramets) asexual reproduction. They are classified into Archiannelida, Polychaeta, Oligochaeta and Hirudinea (Table 1.1). In polychaetes, a pair of bilaterally located parapodia facilitates burrowing and locomotion. It is, however, reduced to setae in oligochaetes and totally missing in archiannelids and hirudineans. The marine interstitial archiannelids are phylogenetically enigmatic annelids and include five families. The exclusively marine polychaetes include burrowing, crawling, digging, drifting (Tomopteris spp), and tubiculous forms (Fig. 1.1) commonly known as sea mice (Aphrodita), scale- (Polynoe), paddle- (Notophyllum), pile/rag/clam- (Nereis virens),

* Names of most annelids species are listed following Worms—World Register of Marine Species; however, some are named, according to author’s citation.

2  Reproduction and Development in Annelida

Table 1.1 Systematic resume of Annelida (compiled from Barnes [1974] and others; terrestrial and amphibious taxa are indicated by bold and italic letters; representative family names alone are indicated) Phylum:  Annelida (15,000 [Wildlife J Junior, 2017], 16,763 [Chapman, 2009], 22,000 [Aguado et al., 2014]) Class: Archiannelida, Polygordius, Nerilla (60 species [Westheide, 1984]) Class: Polychaeta (8,000 [IASzoology.com], 10,000 species [Minelli, 1993], 13,000 [Australian Museum, 2015]) Sub class:  Erranta Families: Aphroditidae, Polynoidae, Siboglinidae, Phyllodocidae, Amphinomidae, Nereididae, Pisionidae, Alciopidae (pelagic), Tomopteridae (pelagic), Hesionidae, Syllidae, Nephtyidae, Glyceridae, Eunicidae, Lysaretidae, Arabellidae, Lumbrineridae, Histriobdellidae, (Ectoparasites), Ichthyotomidae (Ichthyoparasites), Myzostomidae Sub class:  Sedentaria Families: Orbiniidae, Spionidae, Siboglinidae, Magelonidae, Chaetopteridae, Cirratulidae, Flabelligeridae, Opheliidae, Capitellidae, Echiuridae, Cossuridae, Arenicolidae, Maldanidae, Oweniidae, Sabellariidae, Pectinariidae, Ampharetidae, Terebellidae, Sabellidae, Serpulidae, Fabriciidae Class:  Oligocheta (1,700 species [Martin et al., 2008]) Orders:  Lumbriculida (145 species, Ferraguti et al., 1999 [Lumbriculus]), Moniligastridae Order: Haplotaxida Sub order:  Haplotaxina Sub order:  Tubificina Families: Enchytraeidae (670 species, Schmelz and Collado, 2015), Tubificidae (1,000 species, Martin et al., 2008), Naididae (175 species, Ferraguti et al., 1999), Phreodrilidae, Opistocystidae, Dorydrilidae Sub order:  Lumbricina (33–670 species and subspecies in ~ 48 genera [Wikipedia]) Families: Allurioididae, Glossoscolecidae, Lumbricidae, Megascolecidae (Pheretima), Eudrilidae Class:  Hirudinea (700–1,000 species [Govedich and Bain, 2005]) Order:  Acanthobdelida, Rhynchobdellida Families: Glossiphoniidae (Placobdella), Piscicolidae (Pontobdella) Order:  Gnathobdellida Families: Hirudinidae, Haemadipsidae (50+10 species [Won et al., 2014]) (Hirudo, Haemopis) Order:  Pharyngobdellida: Primarily aquatic with some semi-terrestrial species

nuclear- (Namalycastis), fire- (Amphinome), blood- (Glycera), tube- (Onuphis), lug- (Arenicola), bamboo- (Maldane), trumpet- (Pectinaria), spagehetti(Terebella), fan- (Sabella) and feather duster- (Hydroides) worms. The drifting holoplanktonic polychaetes are included in two families Alciopidae and

Introduction  3

Figure 1.1

Fig 1.1 A. Nereis, B. Arenicola, C. Chaetopterus, D.Riftia (Public domain, Wikipedia), E. Polygordius (freehand A. Nereis, B. Arenicola, C. Chaetopterus, D. Riftia (Public domain, Wikipedia), drawing from Fraipont), F. Tomopteris kils (redrawn), G. Ampharete, H.Amphitrite, I. Hydroides, (H and I freehand E. from Polygordius (freehand drawing K. from Fraipont), F. Tomopteris kils (redrawn), drawings R.P.Dales, 1963) J. Earthworm, Amphichaeta (permission by [email protected]), L.Tubifex, G. Ampharete, Amphitrite, I. Hydroides (H andO. I freehand drawings fromfrom K.H.Mann, (copyrightH. free projects.ncsu.edu), N. A. hembrichi, Piscicola (freehand drawing M. Aeolosoma 1962), Dales, P. Human leech J.(A, B, C, F, G, J and O are all copyright free imagesby from dreamstime.co 1963), Earthworm, K. Amphichaeta (permission limnes@chebucto.

ns.ca), L. Tubifex, M. Aeolosoma (copyright free projects.ncsu.edu), N. A. hembrichi, O. Piscicola (freehand drawing from Mann, 1962), P. Human leech (A, B, C, F, G, J and O are all copyright free images from dreamstime.com).

Tomopteridae and are being expanded to Lopadorhynchidae, Pontodoridae, Iospilidae, Poeobiidae and Typhloscolecidae (Bonifazi et al., 2016). The fire worms (Amphinomidae) are known for their brittle poisonous setae and some blood worms can be venomous (Aguado et al., 2014). Though the swarming palolo worms Eunice viridis are collected and consumed by Samoan Islanders (Caspers, 1984), annelids do not directly

4  Reproduction and Development in Annelida

contribute to the global fisheries. However, they serve as key bait in recreational fisheries. More than 65 million anglers from Europe, USA, Canada and Australia catch annually 2 million ton (mt) high value fishes (Pandian, 2015). Between 1966 and 1980, 27–38 million pileworms Nereis virens were harvested annually (Creaser and Clifford, 1982). The estimated market value of the bait worms for Europe alone is ~ US$ 250 million (Olive, 1999). van der Have et al. (2015) have listed some 25 polychaete (pile-, lug-, blood-, sand- and nuclear-worms) species used as bait by anglers in Europe, Korea and China. The demand for these bait worms is so high that N. virens is cultured in the UK and Netherlands; the bloodworm Diopatra bilobata and Korean lug worm Perinereis spp are also cultured in Vietnam, Korea, China and Japan (for export). Among freshwater oligochaetes, Tubifex and the like serve as live feed for domestic ornamental fishes. Many publications (e.g. Marian and Pandian, 1984, 1985, Marian et al., 1989) and privately circulated books (e.g. Pandian and Marian, 1985a) are available describing the procedures for rearing and harvesting the worm are available. Being ecosystem engineers, services rendered by the earthworms needs no emphasis. These worms accelerate degradation of organic matter and molecules produced by plants and other organisms, and render nutrients, especially nitrogen reusable by plants. Total production of mineral nitrogen by the worms ranges between 30 and 50 kg/hectare (ha)/year (y). By altering porosity, these worms contribute to soil structure and thereby water absorption and retention; for example, water infiltration rate through soil can be increased by the worms from 15 to 27 mm/hour (h), resulting in reduced runoff. In soil formation, they breakdown the primary minerals and incorporate them with organic matter. Their aquatic counterparts, the polychaetes and tubificids serve also as ecosystem engineers to turbulate (see Hutchings, 1998) and make the organic matter from sediments and deposits available for benthic productivity. For example, Tubifex benedii, Amphichaeta sannio, Paranais litoralis (naidid) and Manayunkia aestuarina (polychaete) jointly contribute 50–90% of the total invertebrate production in the Forth estuary, Scotland (see Giere, 2006). Inhabiting 1.6 km long, 3 m width and 30 cm depth of intertidal zone of the Pacific coast, USA, the small (25 mm long) opheliid Euzonus mucronata annually turbulates 14,600 ton (t) sediments. Hence, the indirect contribution by the annelids to the global benthic fisheries must be of high order. However, not much information is yet available on quantitative contribution by the polychaetes to the trophodynamics of many ecosystems (cf Pandian, 2016). Time memorial, leeches, especially Hirudo medicinalis have been used in hirudotherapy. H. medicinalis secretes hirudin, a 65 amino acid peptide that inhibits thrombin-catalyzed conversion of fibrinogen to fibrin and prevents host blood from clotting. By inflicting the deepest bite and the most-prolonged post-bite extravasation, a leech can engorge maximally with 50–100 ml human blood (Govedich and Bain, 2005).

Introduction  5

Figure 1.2 A. Described polychaete species during the period from 1750–2020 (from Polychaeta Statistics, modified). A1 in window shows the same relationship for hirudineans (modified from Sket and Trontelj, 2008). B. Number of publications relevant to polychaetes during the period from 1800–2010 (from Faulwetter et al., 2014, modified). B1 in window shows the same relationship for earthworms (modified from Sturzenbaum et al., 2009).

For polychaetes, relatively more information is available on the number of species and publications for the past two centuries. Syllidae, Nereididae, Spionidae and Serpulidae are speciose families each comprising > 500 species (Faulwetter et al., 2014). Described polychaete species number has remained < 2,000 until the 19th century and is expected to increase to ~ 13,000 species by 2020 (Fig. 1.2A). This trend also holds true to hirudineans (Fig. 2.1A1) and perhaps to oligochaetes. Publications relevant to polychaetes, which have remained less than a dozen per decade until 1960, have increased rapidly to ~ 300/decade during 2000–2010 (Fig. 1.2B). This type of spurt may hold true for earthworms (Fig. 1.2B1) and other clitellates also. Hence, this book provides only a ‘snap-shot’ of annelid reproduction and development rather than an in depth or exhaustive description of each item listed in the ‘Contents’.

1.2  Species and Structural Diversity A vast majority of annelids are gonochores and reproduce sexually. Not surprisingly, this feature is reflected in their species diversity. For example, any macrofaunal sample from the Australian soft sediments is reported to hold from 24% (Bass Strait) to 36% (Port Phillip Bay) polychaete species (see Hutchings, 1998). Indicating the sustained contribution to annelidan taxonomy, the described species number has progressively increased from 8,700 (Barnes, 1974) to 14,000 in polychaetes, 4,000 + in oligochaetes and 800 in hirudineans (Rouse and Pleijel, 2006), 16,763 (Chapman, 2009, see also Westheide and Purschke, 2013) and to 22,000 (Aguado et al., 2014). More recently, Bleidorn et al. (2015) have considered that annelids comprise >

6  Reproduction and Development in Annelida

17,000 species (Table 1.2, see also p 5). This number may further increase, as the described polychaete species number alone, which was around 10,000, is estimated to shoot to 25,000 (Hutchings, 1998). Similarly, the number of hirudinean species has also increased from 500 (Barnes, 1974) to 700–1,000 (Govedich and Bain, 2005). Martin et al. (2008) reported that of 1,700 valid species of aquatic oligochaetes, of which the most speciose Tubificidea holds over 1,100 species. However, it must be stated that the annelidan taxonomy remains fluid but dynamic. Erected new species by mis/wrong identification is being continuously corrected. For example, a check-list of polychaete species from the Black Sea reveals that 51 species reported between 1868 and Table 1.2 Distribution and number of annelid species in marine, freshwater and terrestrial habitats Taxon

Habitat (no.)

Total (no.)

Marine

Freshwater

Terrestrial

60

–

–

60

Polychaeta

> 13,000

~ 12†

2**

13,002

Hirudinea

102

482

92

684

+

+

Archiannelida

Oligochaeta Moniligastrida

Australian Museum (2015), Erseus (1994)** Sket and Trontelj (2008) Martin et al. (2008)

145 + 24*

Haplotaxina

++

169

Feragutti et al. (1999)

16

Giere (2006)

~ 670

670

Schmelz and Collado (2012)

+

16†

Enchytraeidae Tubificidae

Westheide (1985)

+

Lumbriculida Tubificina

Reference

600

1,100 + 13*

+

1,713

Naididae

–

175

–

175

Ferraguti et al. (1999)

Martin et al. (2008)

Phrocodriliae

+

+

432

Rhoden (2015)

Ophistocystidae Dorydrilidae

++

Lumbricina

+

Alluroidae

+

++

Lumbricidae

432

Megascolecidae

+

Eudrilidae

+ 13,776

1,939

1,202

16,911

Bleidorn et al. (2015)

* subterranean species recognized by Chatelliers et al. (2009), + freshwater, ++ mostly freshwater, – absent, † riverine polychaetes

Introduction  7

Figure 1.3 A. An errant Nereis showing head/peristomium, trunk and tail regions. B. Ventral view of Nereis head. (modified and drawn from Snodgrass). C. Anterior portion of Hesionides. Note the ventrally directed parapodia adapted for crawling. D. Ventral view of parapodium of Nereis. E. Surface ciliation in filter feeding Phyllodoce (modified and drawn from Segrove). Arrows indicate direction of water current (A. all are free hand drawings; A and D, courtesy Dr. P. Murugesan).

2011 has been synonymized with other species (Sahin and Cinar, 2012). In fact, many reviewers refer only generic names, as species-level revisions are an ongoing research. The annelidan body is distinctly divided into a series of similar segments. These segments are arranged in a linear series along the antero-posterior axis. The prostomium and pygidium, which are not true segments, are located at the anterior and posterior ends of the body. The formation of new segments occurs just anterior to the pygidium. The oldest segments are therefore anterior and the youngest are posterior (Fig. 1.3). Considering polychaetes as representative taxon, the number of segments ranges from a few to as many as 300 in the orbiniid Nainereis dendritica, the body length from 2 mm in the cirratulid Monticellina serratiseta to 3 m in an Eunice sp and the width from 0.4 mm in the cirratulid Caulleriella lafolla to 6 mm in the orbiniid Phylo nudus. Obviously, thinning of the body facilitates burrowing to deeper depths. Table 1.3 and Fig. 1.3 briefly summarize the remarkable structural diversity in prostomium as well as structures like the parapodia associated with each segment of the body. Sensory organs attached to the prostomium and bilaterally located parapodia are greatly modified to suit the burrowing, crawling and tubiculous mode of life and to collect food, as well.

8  Reproduction and Development in Annelida

Table 1.3 Diversity of segmental structures in annelids Polychaeta Crawlers Well developed preoral prostomium bears antennae, palps, eyes, nuchal organ. Fleshy biramous parapodia consisting of an upper notopodium and lower neuropodium terminate by an invaginated series of setae. Cirri form processes arise from the bases of these podia. The segment is supported by one or more chitinous rods (Fig. 1.3A, D) Drifters Structures like the crawlers. Transparent. Enormously large eyes in alciopids and membranous parapodial pockets in tomopterids (Fig. 1.1). Setae are absent Burrowers Reduced prostomium. Antennae, palps and eyes absent but carry food collecting structures (e.g. tentacles of Amphitrite, Fig. 1.1A). Reduced uniramous parapodia represented by transverse ridges, which may be modified into hooks Borers Spionid Polydora burrows into the shell aided by a viscous fluid, which dissolves the exposed calcite crystals of the shell (Pandian, 2017) Diggers Heads of the pectinariid worms bear rows of large conspicuous seta used for digging Tubiculous worms Carnivores: Segmental structures do not greatly differ from those of crawlers. Inhabit in vertically or horizontally straight hyaline/membranous tubes Others: Prostomium and sensory structures reduced or absent. Specialized food collecting structures present. Sabellids build membranous sand grain tubes. Serpulids secrete calcareous tubes. Oweniids carry their tubes Oligochaeta No parapodia. Setae can be long or short, straight or curved, heavy or needle-like and blunt, pointed, forked, pinnate or plumose. The setal shaft is S-shaped with middle swelling or nodule. Longer setae are characteristics of aquatic species. Setal number is fixed at 8 in aquatic species and Lumbricus. In mature oligochaetes, certain anterior adjacent segments thickened and swollen by glands that secrete mucus for copulation and for formation of cocoon. Their glandular area is called clitellum, which forms a girdle around the body Hirudinea No head appendages but eyes are present. No parapodia and setae except in Acanthobdella. Dorso-ventrally flattened body with suckers at the anterior and posterior ends. Segment number is fixed at 34. Septa absent

1.3  Geographic Distribution Geographic range is the horizontal and vertical areas, in which populations of a species are distributed. It is a species specific trait with evolutionary consequences inclusive of determination of life span and exposure to different environmental and biological factors. In freshwaters, the oligochaetes Tubifex

Introduction  9

tubifex is cosmopolitan in distribution; so is the polychaete Nereis virens in the intertidal zone of marine environment. The archiannelids and polychaetes are exclusively marine, although a couple of polychaetes Peregodrilus heideri and Hrabeiella periglandulata are almost terrestrial inhabiting moist soil (see Rota et al., 2001), and a dozen of them inhabit rivers and estuaries. On the other hand, oligochaetes and hirudineans are found in marine, freshwater and terrestrial habitats. Experts have provided counts on the number of species for groups within, oligochaetes but a total of which does not tally with that reported for the class (Table 1.2). As a result, the sum of subtotals does not tally with recently reported 22,000 species of annelids (Aguado et al., 2014). For example, the estimate of 5,000 clitellate (Oligochaetea + Hirudinea) species by Martin et al. (2008) does agree with the subtotal estimates reported by different experts for the suborders and families. Despite this constraint, and considering 17,000 species number for annelids (Bleidorn et al., 2015), ~ 11.5 and 7% annelids are distributed in freshwater and terrestrial habitats, respectively; the remaining 81% are marine inhabitants. Reports on geographic distribution of a specific taxon in some of these habitats are available; for example, polychaetes in the Black Sea (Sahin and Cinar, 2012), naidids in African freshwaters (Grimm, 1987) and terrestrial enchytraeids in Latin America (Schmelz et al., 2013). A report on distribution within the major geographical zones is also available but only for freshwater hirudineans (Table 1.4). In general, the geographic range is increasingly limited in the following descending order: planktotrophy < lecithotrophy < brooding/viviparity < exclusive asexual reproduction. Within Cirratulidae, the range is extended from the Washington coast of North America to Puget Sound, Washington (DC) for Chaetozone acuta, which broadcasts small (50–60 µm) planktotrophic egg; but it is limited between California and Mexico in the west Pacific coast for C. corona spawning 75 µm egg, to arctics alone for C. setosa releasing 120–160 µm egg, and to the east and west coasts of North America around the equator for Aphelochaeta monilaris spawning 275–300 µm egg (see Blake, 1996). Table 1.4 Number of freshwater hirudinean genera and species present in major geographical zones (compiled from Sket and Trontelj, 2008) Zone

Genera (no.)

Species (no.)

Palaearchtic

45

187

Nearchtic

24

79

Neotropric

27

107

Afrotropic

22

50

Indotropic

27

64

Australisian

15

34

10  Reproduction and Development in Annelida

Polychaetes dominate the benthic macrofauna (Gremare et al., 1998). From their long- (20 years) term study on polychaetes at two sites in the western English Channel, Quiroz-Martinez et al. (2012) have analyzed the dynamics of abundance as function of species diversity. With increasing abundance up to 8,000 individuals, species diversity is also increased from ~ 40 to ~ 60 in a site but ~ 25 to ~ 35 species in another site. Understandably, vertical expansion especially into abysmal depth encounters more environmental challenges than horizontal distribution. The maximum depth, at which the soft-bodied annelids have been collected, is in the range of ~ 4,000 m (Table 1.5). This depth is comparable with the external shell-less molluscs like that aplacophorans (3,200–4,000 m) and cephalopods (2,430–4,850 m) (Pandian, 2017). However, the asteroid and ophiuroid echinoderms with internal skeleton penetrate to 6,035 m (Pandian, 2018). Strikingly, the shelled molluscs are capable of expanding up to 6,370 m (bivalves) and 9,050 m (prosobranchs) (Pandian, 2016). The presence of hard shell(s) enables these shelled molluscs to expand up to the greatest depths. The abysmal depths, from which annelids have been collected, ranges from 1,500 m for the orbiniid Phylo nudus to 4,016 m for the cirratulid Chaetozone gracilis for polychaetes and 700 m for the naidid Nais abissalis to 1,680 m for four taxa of Tubicinae in Lake Baikal and to 4,900 m depth in the Indian Ocean for the tubificid Abyssidrilus stilus (Table 1.5). Describing the pattern and scale of biogeographic variability of abyss in the North Atlantic Ocean, Smith et al. (2006) have reported that the diversity of polychaete species declines perceptibly below 3,000 m depth. Bathymetric distribution of polychaetes seems to be determined by the availability of dissolved oxygen. For example, the Black Sea becomes anoxic and is contaminated with hydrogen sulfide below 150–200 m depth. The number of polychaetes species present in the sea progressively decreases from ~ 100 between 0 and 10 m to 0 below 150 m depth (Sahin and Cinar, 2012). Obviously, the polychaetes in the Black Sea are unable to switch over to anaerobiosis and colonize the anoxic depths. However, Hartman (1966, 1967) has indicated that Antarctic collections have included 23 polychaete species belonging to 14 genera from the depths of 4,930–4,963 in the South Sandwich trench. In freshwater, the naidids occur up to 50 m depth but rarely Nais abissalis has been collected from 700 m depth in Lake Baikal. Typically, they feed on phytoplankton (Brinkhurst and Gelder, 1991). With increasing depth, they do not find adequate phytoplankton below 100 m depth (Bondarenko et al., 1996). However, the other oligochaetes, feeding on detritus from sediments, represented by enchytraeids (1,600 m), haplotaxids, lumbriculids and tubificids have been collected from the depth of 1,680 m in Lake Baikal but with perceptible decline from ~ 1,700 no./m2 at ~ 50 m depth to < 10 no./m2 at 1,680 m depth (Martin et al., 1999). In contrast to the Black Sea, both water column and sediments of Lake Baikal is well oxygenated at all depths (Martin et al., 1999). Considering, hydrostatic pressure, light, temperature and food

Introduction  11

Table 1.5 Vertical distribution of annelids (from Erseus, 1994*, Blake, 1996, Dean, 1995†, Martin et al., 1999**) Species

Depth (m)

Location

Polychaeta Orbinidae (14–15 species) Califa calida Phylo nudus

470–2,000 400–1,500

Off California California

Paraonidae (85 species) Arcidea monicae

200–300 590–1,745 80–1,480 10–2,000 44–2,400 100–3,000

A. wassi A. ramosa A. simplex

Mediterranean California California California Western Pacific, Japan California

Apisthobranchidae (5 species) Apisthobranchus

3,000 Spionidae (90 species)

Spiophanes anoculata S. kroeyeri

463–2,400 3,500

California Australia-Antartic

Poecilochaetidae Poecilochaetus johnsoni

90–189

California-Mexico

Chaetopteridae (30 species) Phyllochaetopterus limicolus

119–3,000

California

Cirratulidae (46 species) Chaetozone spinosa

280 2,623–2,955 4,016 1,260–2,400 255–3,000

C. gracilis Tharyx kirkegaardi

Japan California Catalina Island California Atlantic

Cossuridae (16–17 species) Cossura pygodactylata C. candida C. modica C. brunnea C. rostrata

1–2,720 11–2,400 985–2,955 1,600–2,200 6–3,348

Western France Mexico, Baja California Oregon-California North Carolina-Mexico Oregon-Western Mexico

Oligochaeta Nais abissalis (Naichidae) Propappus glandulosus (Enchytraeidae) Haplotaxis sp (Haplotaxidae) Stylodrilus asiaticus (Lumbriculidae) Balkaiodrilus maievici (Tubificidae) Abyssidrilus stilus (Tubificidae)

700 1,600 1,680 1,680 1,680 4,900

Lake Baikal** Lake Baikal** Lake Baikal** Lake Baikal** Lake Baikal** Indian Ocean*

Ctenodrilidae Raricirrus variabilis

4,000

Virgin Islands†

12  Reproduction and Development in Annelida

availability, Martin et al. (1999) have found that other than oxygen level, food availability is the most dominant factor that determines the bathymetric distribution of the haplotaxids. Incidentally, the cocoons of Tubifex tubifex and Potamothrix hammoniensis deposited at the sediment surface are all eaten by the fish Abramis brama but > 99% of them survive, when deposited at 20 mm depth (Newrkla and Mutayoba, 1987). There are adequate indications that in the deep anoxic sediments, tubificids may become anaerobic (e.g. Narita, 2006). With abundance of food at 4,900 m depth, A. stilus may have switched to anaerobiosis. Interestingly, the highest altitude, at which the enchytraeid Buchholzia appendiculata has been collected, is ~ 2,000 m above sea level (asl) in the montane regions of South America (Schmelz et al., 2013). Amazingly, the oligochaetes have a range of vertical distribution of 6,900 m, i.e. from the depth of 4,900 m to an altitude of 2,000 m. In the absence of moisture/water, the oligochaetan earthworms are unable to penetrate below 80 cm depth (e.g. Glossodrilus, Jimenez and Decaens, 2000). Soil moisture potentials, measured in –kPa unit, reduce the optimum for growth and reproduction at –2 kPa to almost 0 at –50 kPa (Johnston et al., 2014). In the presence of water at the peculiar subterranean habitats, the stigobiont oligochaetes are known to flourish (Chatelliers et al., 2009). With their elongated, segmented and flexible body shape, these stigobiont oligochaetes are pre-adapted to the subterranean habitats and do not exhibit troglomorphic features like the absence of body pigments and eyes, elongated appendages and increased sensory structures. The stigobiont oligochaetes belonging to 42 genera in 17 families are reported from the subterranean aquatic habitats. Of them, the number of species belonging to the lumbriculid Trichodrilus and tubificid Rhyacodrilus accounts for 23 and 11%, respectively (see Chatteliers et al., 2009). In his informative taxonomic description of Californian polychaetes, Blake (1996) has provided useful data on body length and depth, at which orbiniids, paranoids, spionids, cirratulids and others have been collected. When data on body length are plotted against depth, different trends become apparent (Fig. 1.4), clearly indicating that body size may not be a factor for polychaetes to penetrate into greater depths. Unlike oligochaetes, not all polychaetes switch over to anaerobiosis at hypoxic/anoxic depths (e.g. Black Sea, Sahin and Cinar, 2012). Hence, it is likely that the oxygen levels of water and sediments as well as respiratory structures may prove to be important factors in bathymetric distribution of polychaetes. Endemism: Depending on limited powers of motility and larval dispersal being at the mercy of waves and currents, many annelids are endemic. Not surprisingly, of 155 oligochaete species, 114 are endemic to the truly ancient and long-lived Lake Baikal (Martin et al., 1998). Data on the distribution of stigobiont oligochaetes in the subterranean habitat suggest pronounced endemism. More than 60% species are known only from the type locality (see Chatelliers et al., 2009). Likewise, hydrothermal vent-inhabiting

Introduction  13

Figure 1.4 Vertical distribution of polychaetes belonging to four selected sedentarian families (drawn from data reported by Blake, 1996).

vestimentiferan polychaetes are also endemic. Though polychaetes are continuously occupying a wide vertical depth range, for example Cossura candida from 11 to 2,400 m depth off California, Mexican waters (Table 1.5), many of them are reported as endemic. From 19 expeditions carried out during the last 124 years for polychaete taxonomic research around the southernmost tip of the South American continental shelf, Martin et al. (2005) have recorded 431 species belonging to 108 genera and 41 families. Subregions on the Pacific and Atlantic sides are characterized by the presence of 10% endemic species. Investigation on the geographic distribution of the endemic 178 polychaete species between 18º and 56º S of South America has revealed the marked peak endemic hotspot between 36º and 41º S, corresponding to a peak in species richness (Moreno et al., 2006). Symbiosis: Many annelids symbiotically inhabit as (e.g. Dipolydora commensalis on hermit crab within the shell, Lindsay and Woodin, 1993) or as endosymbiont in another polychaete, e.g. Veneriserva pygoclava meridionalis in aphroditid host Laetmonica product (Micaletto et al., 2002) and within the canals of aquiferous systems of sponges. For example, 33 syllid species constituting > 9% of all the known symbiotic polychaetes are reported to inhabit the canal system of sponges. As many as 600 Haplosyllis spongicola

14  Reproduction and Development in Annelida

happily inhabit within a small (16.5 cm3) Ciona sp but without disturbance to the host. H. spongicola can inhabit in 36 species of host sponges (Lopez et al., 2001). The earthworm Eudrilus eugeniae symbiotically engage Bacillus endophyticus to draw riboflavin, an essential nutrient for regeneration (Samuel et al., 2012, Subramanian et al., 2017).

1.4  Gutless Oligochaetes Since the discovery of a couple of gutless tubificids in coralline sands of Bermuda (Giere, 1979), over hundred species belonging to two phallodrilan genera Olavius and Inanidrilus (= Phallodrilus) with no digestive system and excretory organs have been described (Erseus, 2003). In I. leukodermatus, the integument surface of the long, slender worm is much annulated and highly folded into numerous tiny irregular ridges. These expansions increase the worm body surface nearly 10 times (Giere, 1981). Unlike in other annelids, the epidermal layer is unusually thick with extensions crossing the cuticle (microvilli) and ending in epicuticular projections. Consequently, a wide cuticle-epidermal interface is present. All the studied phallodrilan 30 species are reported to incorporate a fairly thick layer of extracellular bacteria beneath the cuticle. Only in two species Olavius algarvensis and O. ilvae from the Island Elba, Italy, the microbes are enclosed by the epidermal cells and thus attain an intra-cellular position (Giere and Erseus, 2002). In these gutless tubificids, the symbiotic microbes belonging to the following phytotypes are present: 1. Large, oval-shaped γ Proteobacteria, which are shown to be sulfide oxidizers (Dubilier et al., 1995). 2. Small rod shaped α Proteobacteria, which reduce sulfate into sulfide. This unique ‘cyclotrophism’ by these symbiotic microbes enables the gutless tubificids to thrive not only in sulfide/oxic interfaces but also in oxic layers (Giere et al., 1991). 3. Some phytotypes, including Spirochaeta also occur; however, their symbiotic function is not known (Giere, 2006). Yet, the quantum of symbiotic microbes harbored in relatively smaller area of these gutless tubificid oligochaetes is too small, in comparison to the massive trophosome filled with prokaryotic symbionts in the hydrothermal vent-inhabiting vestimentiferan siboglinid polychaetes. Hence, the overall contribution by the symbiotic microbes is ranked low (Giere et al., 1984). Interestingly, these gutless tubificids draw nutrition through (i) osmotrophism and (ii) symbiosis. With relatively larger body surface area to volume ratio and thin cuticle, the oligochaetes are well adapted to uptake nutrients from ambient water across the body wall. Absorption of many amino acids and glucose by osmotrophic fauna may proceed from extremely low concentration against concentration gradients of 4–6 orders magnitude; on accumulation, these organic substances are metabolically used (see Pandian,

Introduction  15

1975). Southward and Southward (1980) have estimated that the gutless worms can absorb glucose to meet 30% of the metabolic needs. The pore water within the sediments, from which the gutless tubificid I. leukodermatus have been collected, contains an extremely high concentration of glucose and fairly high levels of amino acids. I. leukodermatus is able to absorb mainly hexoses (but not pentoses) at the rate of ~ 150 µg glucose/h. Among amino acids, aspartate is preferably absorbed in comparison to glutamate (Giere et al., 1984). As in hydrothermal vent inhabiting vestimentiferans, the tubificids display substantial activity of ribulose-1,5-biphosphate-carboxylase, a marker enzyme of the Calvin-Bensen cycle, known to be present only in bacteria and plants. The high levels of ATP-sulfurylase and sulfide oxidases indicate that enzymatic sulfur metabolism is carried by the symbiotic bacteria. These enzyme studies suggest that the gutless tubificids are able to draw ATP through the symbiotic microbes and oxidize sulfide for the fixation of inorganic CO2 from ambient water. Hence, the gutless tubificids like I. leukodermatus can also thrive in sediments containing high concentrations of hydrogen sulfide. The preferred zone of I. leukodermatus lies between 5 and 7 cm depth, which correspond to +50 and –50 mV, i.e. in and around the redox discontinuity (RPD) layer (Fig. 1.5), where extremely high concentrations of sugars and amino acids are present in the pore water within the sediments. The worm keeps migrating between the upper oxic and lower anoxic depths. At the lower anoxic depth, it acquires and accumulates reduced sulfur but the necessary binding of oxygen occurs at the upper micro-oxic sediment layers (Giere et al., 1991). Strikingly, these gutless tubificids have successfully conquered and colonized an ecological niche, so far unoccupied by any other interstitial fauna (Giere et al., 1984). In these gutless tubificids, the location of the microbial symbionts and complicated mode of reproduction suggests vertical transmission. In

Figure 1.5 Vertical distribution of the gutless oligochaete Inanidrilus leukodermatus as functions of A. redox potential and B. worm density, and C. worm density as function of sediment redox potential (compiled, modified and redrawn from Giere et al., 1991).

16  Reproduction and Development in Annelida

fact, studies on I. leukodermatus from Bermuda have revealed the vertical transformation of oval sulfide oxidizing γ-Proteobacteria, infecting the sticky eggs laid singly on the sediment grains and not deposited in cocoons, as in other oligochaetes. However, the differences in the bacterial association among the populations of O. algarvensis indicate the horizontal transmission, i.e. environmental acquisition of the symbionts. Among the vent-inhabiting bivalves, vertical transovarian transmission occurs in the vesicomyid clams but horizontal environmental acquisition in mytilid mussels (see Pandian, 2017). It is likely the vertical transmission occurs in Inanidrilus species but horizontal one in species belonging to Olavius.

1.5  Hydrothermal Vents and Cold Seeps Deep sea hydrothermal vent habitats are characterized by temperatures higher than ambient for deep sea, lower oxygen concentration, higher levels of toxic sulfide compounds, iron, and other metals and gases (Chevaldonne et al., 1997). Mixing of the hot water arising from the volcanic spring with cold ambient water creates biologically habitable regimes (Nyholm et al., 2008). Typically, the hydrothermal vents are linear, discontinuous and shift along the ridges and thereby generate short-lived habitations. In the Pacific, the vents are distributed between 48º N and 22º S (Lutz, 1988, however see also Van Dover, 1994). The startling discovery of hydrothermal vents in 1977 heralded the description of dense benthic fauna in the vents. In these benthic communities, the sole base of the food chain is the chemoautotrophic production by microbial symbionts using hydrogen sulfide as geothermal energy source (see Cavanaugh et al., 1981). The chemoautotrophic symbionts harness geothermal energy through oxidation of reduced chemical substances, fix inorganic compounds, and use them for biosynthesis and growth, although it is not yet known how the vestimentiferans acquire nitrogen for protein synthesis. Indeed, hydrothermal vents are now known to support extensive but endemic biological communities (Nyholm et al., 2008) that are found world over at marine hot springs distributed along the midocean system (Hurtado et al., 2004). Expectedly, island-like patchy endemic populations are distributed intermittently along tectonically or volcanically active spreading vent segments, which are separated by tens to hundreds of kilometers (Hurtado et al., 2004). In these vents, the faunal assemblage includes a few crustacean species (e.g. galatheids: Munidopsis subsquamosa, M. lentigo; brachyuran Bythograea thermydron; caridean shrimp Alvinocaris lusca), the bivalve molluscs belonging to Vesticomyidae, and Bathymodiolinae, a single clade within the family Mytilidae (see Pandian, 2017) and ~ 32 polychaete species (Lutz, 1988). Perhaps, the vestimentiferan tubeworms are the first to be

Introduction  17

discovered. Though initially grouped with Pogonophora, phylogenetic and embryological evidences have convincingly shown that these gutless deep sea tubeworms are siboglinid polychaetes (McHugh, 1997, Marsh et al., 2001, Rouse et al., 2008). Within the family Siboglinidae, three subfamilies are included: Vestimentifera, Monilifera (a small group of gutless worms living on submerged wood) and Osedacinae (with Osedax as the only genus). The subfamiliy Vestimentifera is again subdivided into three infra-families: Lamellibrachiinae, Escarpiinae and Tevniinae (Karaseva et al., 2016). The tevniinids inhabit exclusively on the rocky substrates of hydrothermal vents of the Pacific Ocean. The lamellibrachiinids and escarpiinids occupy both soft and rocky substrates of cold seeps (see Kobayashi et al., 2015) and the periphery of hydrothermal vents (Karaseva et al., 2016). Descriptions for 19 valid vestimentiferan species and their bathymetric distribution (Fig. 1.6) are available. Notably, their distribution ranges from > 100 m to 3,500 m depth,

Figure 1.6 Bathymetric distribution of vestimentiferan tubeworms in hydrothermal vents (from Karaseva et al., 2016, modified).

18  Reproduction and Development in Annelida

limited between 600 m and 1,000 m depth for Seephiophila jonesi but widely scattered for Lamellibrachia spp. Like the moniliferans, the vestimentiferan tubeworms also lack a functional digestive system. They derive nutrition from chemoautotrophic microbial symbionts harbored in the trophosome located on their elongated trunk. Not surprisingly, > 50% of the total body length (1.5 m) of the giant tubeworm Riftia pachyptila is occupied by the most extensive trophosome and the gonad (Cavanaugh et al., 1981). Scanning and transmission microscopic studies have confirmed the presence of prokaryote in the trophosome. Direct count has shown the presence of > 3.7 × 109 microbes/g live trophosome tissue. In 21 of 31 specimens of Riftia examined, crystals (100 µm) of elemental sulfur have been found within the trophosomal tissue. The presence of sulfur crystals within the tissue suggests that the prokaryotic microbes are able to chemotrophically oxidize sulfide (Cavanaugh et al., 1981). Felbeck (1981) has reported that high levels of the enzymes thiosulfate transferase, APS reductase and ATP sulfurylase are involved in generation of ATP through sulfide oxidation. Further, the extended vascular system within the trophosome (Jones, 1981) and the relative insensitivity of the blood oxygencarrying capacity to changes in levels of temperature and CO2 suggests the presence of special adaptation in this worm to ensure sustained supply of O2 and CO2 to the trophosomal symbionts (Arp and Childress, 1981). Structures similar to the symbionts have also been described within the trophosome of other vestimentiferans like Lamellibrachia luymesi and L. barhami. Hence, it is likely that the vestimentiferan tubeworms draw energy through the same chemoautotrophic pathway. Nyholm et al. (2008) have reported the metabolite flux and transcriptomic data for Redgeia piscesae and indicated that the host sustains substrate availability, which potentially regulates the host’s transcriptome or symbiont’s cell cycle. The growth rate of R. piscesae ranges widely from 0 to 25 cm/y and is highly variable between individuals within the same aggregate and between different vent sites. Consequently, the minimum age of the aggregate also ranges from 10 to 30 years (Urcuyo et al., 2007). Considering the similarity of faunal assemblages in these geographically widely separated hydrothermal vents, and sessile/sedentary nature of the adult worms, dispersal can occur only through the larval stage. In this context, the following observations are relevant: (i) R. pachyptila produces neutrally buoyant small (~ 100 µm) but lecithotrophic eggs. Its larvae can persist in the water column for ~ 34 days (Marsh et al., 2001), during which they may disperse over 100 km. (ii) Adults of Alvinella prompejana can exit from their tubes and swim vigorously. They release larger (~ 200 µm) negatively buoyant lecithotrophic eggs. The eggs may drift with bottom currents and are characterized by intermittently arrested embryonic development, which extends the dispersal duration (Pradillon et al., 2001). (iii) Megaplumes generated by volcanic eruption rises as much as 1,000 m above the axial

Introduction  19

walls and can potentially transport the buoyant larvae and juveniles across vast distances (Mullineaux et al., 1995). However, the juveniles may have to withstand the non-vent conditions for a certain period (see Chevaldonne et al., 1997). Hurtado et al. (2004) have studied the dispersal on a few polychaete vent species across ~ 7,000 km of the East Pacific Rise (EPR) and Galapagos Rift (GAR). The dispersing vent larvae have to overcome the following filters/barriers: (i) East Microplate Region (EMR), and (ii) that separating EPR and GAR populations, (iii) equator separating northern and southern EPR populations and (iv) Rivera Fracture Zone (RFZ). These filters are shown to work on different time scale and to different degrees among the examined vent taxons. The equatorial region exhibits combinations of deep oceanic currents and topographical features that limit faunal exchange between EPR and GAR and across the equator along the EPR. In the vestimentiferan, the mode of development, dispersal and genetic exchange have been repeatedly discussed. Of ~ 19 vestimentiferans, spermiogenesis has been described only for five species: Riftia pachyptila, L. luymesi, L. barhami, Paraescarpia echinospica and Ridgeia piscesae (see Marotta et al., 2005). Sperms are released as bundles (e.g. Tevnia jerichonana) and spermatozeugma (e.g. R. pachyptila). The sperm bundles eventually disintegrate in sea water and the freed spermatozoa are capable of swimming. Sperm masses adhering to female’s body (R. piscesae) or in the spermatheca (T. jerichonana) have been reported. Direct sperm transfer from males to females has been observed in R. piscesae. However, brooding of embryos has never been observed in any vestimentiferan. Although apparent spawning has been observed in R. pachytptila (Van Dover, 1994) and L. luymesi (Hilario et al., 2005), it is not known whether the ‘spawn’ consists of unfertilized eggs/ zygotes or developing embryo. Presumably, the eggs are fertilized either in the ovisac just before spawning or externally, as eggs are released. According to Hurtado et al. (2004), the annelids in the Pacific hydrothermal vents adopt different strategies for long distance dispersal. For example, Alvinella prompejana spawns large negatively buoyant eggs (~ 200 µm) that may be drifted by demersal currents. R. pachytptila releases neutrally buoyant (~ 100 µm) with adequate resource to survive in the water column for ~ 38 days. Branchipolynoe symmytilida produces very heavy large eggs (~ 500 µm) and shows no genetic differentiation over long distance. In fact, low mitochondrial diversity is reported in many vent polychaete species (e.g. T. jerichonana) is indeed great.

1.6  Energy Budget Resource allocation for reproduction may ultimately depend on one or more of the energy transformation steps, namely, food acquisition, digestion and

20  Reproduction and Development in Annelida

absorption, assimilation, respiration, somatic growth and reproduction (Fig. 1.7). For example, blood ingestion shows a direct relation to reproductive output in Hirudo medicinalis (Davies and McLoughlin, 1996). In animals, energy budget is assessed by estimation of C = F + U + R + P, where C is the food energy consumed, F, U and R, the energy lost on feces, urine and metabolism, respectively and P, the energy gained due to growth (e.g. Pandian, 1987). In ecological energetics, one or more energetics components have been estimated at population level in many polychaetes (e.g. Dixon, 1976, Banse, 1979) and oligochaetes (e.g. Dash and Patra, 1977, Senapati and Dash, 1983). The physiological energetics is associated with relatively more precise estimates of these components (e.g. Ivleva, 1970, Goerke, 1971). Obviously, energetics becomes important for resource allocation for reproduction and development. For allocation to regeneration, the earthworm Eudrilus eugeniae engages symbiotic microbes like Bacillus endophyticus to draw riboflavin (Subramanian et al., 2017). However, estimation of each one of the energetics components in annelids is cumbersome and unwieldy for the following reasons: (i) osmotrophism in gutless annelids, (ii) contribution to C by symbiotic microbes, (iii) consumption of egested feces, (iv) anaerobiosis, (v) aestivation/hibernation, (vi) regeneration and (vii) clonal reproduction. Food (C) and acquisition: Besides the conventional heterotrophic acquisition of food by a vast majority of annelids, some gutless annelids are osmotrophic and/or chemoautotrophic. 1. A vast majority of polychaetes and oligochaetes are microphages and feed on bacteria, fungi and protozoa adhering to sediments, sands and other substrates. 2. Terrestrial oligochaetes especially, the earthworms undergo inactive periods of seasonal aestivation (for definition, see Pandian, 2017) for a few months (e.g. see Dash, 1987). In deep sediments of Lake Baiwa (Japan) with no oxygen during four summer months from July to September, the tubificid Rhyacordrilus hiemalis undergoes anaerobic aestivation (Narita, 2006). Polychaetes inhabiting intertidal zone undergo a short or longer period of intermittent tidal aestivation. During these periods, facultative anaerobes suppress aerobic respiration but subsequently switch over to energetically costlier anaerobiosis. Apart from the inaccessibility to food (and thereby reducing C), energy expended on R becomes complicated in these aestivating annelids. 3. Many annelids have the ability to regenerate the lost tissue(s)/organ(s). Some of them can regenerate the fraction of lost anterior or posterior body, i.e. unidirectional cloning. Others can undergo architomic or paratomic asexual reproduction namely multi-directional cloning. Apart from switching the mode of food acquisition, the loss of both feeding palps reduces the feeding duration in a spionid Pseudopolydora kempi japonica but not in another spionid Rhynchospio glutaeus (Lindsay and Woodin, 1995). In yet another spionid Pygospio elegans, C limits asexual reproduction (Wilson, 1985). Hence, regeneration may considerably alter C but reduced C can inhibit cloning.

Introduction  21

Consumption (C): Feeding behavior ranges from herbivory (e.g. naidids feeding on phytoplankton) to carnivory (e.g. leeches) and the feed ranges from suspended to deposited food particles. Hence, estimation on C is more difficult, cumbersome and unwieldy. However, new methods to estimate the microbial feed of oligochaetes (Jones and Mollison, 1948) and ingestion rate of polychaetes (Cammen, 1980a) have been developed. In general, ingestion rate is inversely related to body size and nutritional value of the substrate. In oligochaetes, the rate increases from 80 mg/g worm/d fed on dung in Allolobophora caliginosa to 7,000 mg/g/d in soil feeding Millsonia anomala (see Dash, 1987). Hirudineans are carnivores and feed on animal preys or suck their blood. Antiserum and precipitation test of the gut contents of Glossiphonia complanata consist mostly (85%) molluscs, oligochaetes and chironomids (see Dash, 1987). The gnathobdellid leech Limnatus nilotica requires seven–nine or six feedings of blood from frogs or dogs to attain sexual maturity at the age of 17–20 months and body size of 0.5–2.0 g (Negm-Eldin et al., 2013). On a saturated feeding, a leech may consume 4- (e.g. Limnatus, Johanssonia), 5(e.g. Hirudo) and 10-times (e.g. Haemodipsa) of its own normal body weight. The duration of digestion in L. nilotica lasts from 10–20 days to > 8 months, depending on leech body size and frog’s or dog’s blood. Briefly, feeding in gnathobdellids may be regular once a day or once a few days/months, as in glossiphoniids. Apart from this unwieldy procedure for estimation of C, osmotrophism in polychaetes deter precise estimation of C. An account on chemoautotrophism was provided earlier.

1.6.1 Osmotrophism In surface waters of the ocean, total Dissolved Organic Matter (DOM) amounts to ~ 1 mg/l. Free amino acids, comprising 5% of DOM, occur at concentrations of 5 × 10–7 M/l in free water and 1.1 × 10–4 M/l in interstitial water of sediments (see Pandian, 1975), in which most of the soft-bodied annelids thrive. Indeed, sea water is an organic ‘soup’. Not surprisingly, primary productivity of the osmotrophic, heterotrophic bacteria (0.2–0.5 g C/m2/d) exceeds that (0.2–0.3 g C/m2/d) of autotrophic phytoplankton in the Pacific waters (see Pandian, 1975). As early as in 1909, Putter rightly made the first claim that DOM may also be absorbed across the body surface and used as energy source by animals, as well. In animals, the uptake rates of DOM follow Michael-Menten kinetics, with rates increasing to a threshold as a substrate concentration increases. An argument against Putter’s claim is that if DOM can be absorbed through body surface against concentration gradients, the DOM from body fluids can also be leaked through body surface into the ambient sea water (see Pandian, 2017). Ferguson (1971, 1982) has measured both influx and efflux of free amino acids in many invertebrates including polychaetes and found that the net influx of amino acid is

22  Reproduction and Development in Annelida

overwhelmingly inward, with a single exception of glycine. Siebers (1976) has made a more detailed study on absorption of neutral and basic amino acids across the body surface by Enchytraeus albidus and Nereis diversicolor. He has found that neutral amino acids (glycine, L-valine) are transported against the gradient in a saturable process but not the basic amino acids like L-arginine and L-lysine. The uptake of amino acids alone contributes up to 25% of the nutrient requirements in Nais elinguis (Petersen et al., 1998). Not only sugars (cf Southward and Southward, 1980) and amino acids but also fatty acids are also absorbed by the polychaete Cirriformia spirobranchia (Testerman, 1972). Notably, the uptake of DOM by polychaetes is reduced under anaerobic conditions (Jorgensen and Kristensen, 1980b). The uptake of DOM from ambient sea waters by the polychaetes has remained a hot area of research. Between 1963 and 1982 alone, as many as 17 groups of researchers have examined the ability of 22 polychaete species to assess the net influx of DOM into the worm’s body. Most of these authors have concluded that a smaller or large fraction of metabolic requirement is accounted by the net uptake of DOM (Table 1.6). An objection raised by Jorgensen (1976) is that the interstitial amino acid concentrations of 50 µM may facilitate substantial uptake but a negligible one in the overlying surface water with a concentration of 0.5 µM. Secondly, a few polychaetes that have been investigated are burrowers (e.g. Glycera americana, Ferguson, 1982) or tubedwellers (e.g. Lanice conchilega). The DOM content of water in the burrow and tube may vastly differ from that of interstitial water, as water in the burrow/tube is being continually exchanged by ventilation. The third but a more serious objection is the failure of some authors to establish relevant to environmental concentration for the target substrates (e.g. Siebers, 1982). However, these objections may not hold water for following reasons: 1. Of ~ 21 group of authors, 11 groups have confirmed the net uptake. 2. There are > 100 gutless tubificids, which mostly depend on osmotrophism alone. Table 1.6 Polychaete species that are reported to uptake DOM Net uptake of DOM measured 1. Capitella capitata (Stephens, 1975), 2. Cirratulus hedgepethi, 3. Diopatra cuprea and 4. Glycera americana (Ferguson, 1982), 5. G. dibranchiata (Stevens and Preston, 1980, Preston and Stevens, 1982), 6. Marphysa sanguinea and 7. Pareurythoe californica (Costopulos et al., 1979), 8. Nereis diversicolor (Ahearn and Gomme, 1975, Stephens, 1975), 9. N. succinea (Jorgensen and Kristensen, 1980a, b), 10. N. virens (Jorgensen, 1979, Jorgensen and Kristensen, 1980a, b), 11. Stauronereis rudolphi (Testerman, 1972) Uptake of DOM alone measured 1. Clymenella torquata (Stephens, 1963), 2. Eunereis longissima, 3. Goniada sp and 4. Nephtys sp (Southward and Southward, 1972), 5. Lanice conchilega (Ernst and Goerke, 1969), 6. Lumbrinereis spp, 7. Nainereis dendritica and 8. Podarke pugettensis (Testerman, 1972), 9. Neanthes arenaceodentata (Reish and Stephens, 1969), 10. Nereis limnicola (Stephens, 1975)

Introduction  23

Feces (F): Polychaetes have the ability to digest a wide variety of organic substances (Cammen, 1987). They extract the required carbon equally from both the carbon-rich microbes and carbon-poor detritus by increasing the ingestion rate of the latter by ~ 30 times (e.g. Nereis succinea, Cammen, 1980b). Similarly, the earthworms also increase the ingestion rate to acquire adequate nutrients. The Gut Load (GL), the amount of (dry) substrates ingested as a percentage of body weight, varies with nutrient quality; for example, the GL increases from 100% of moist loam soil to 600% of soil with cellulose and to 700–800% of soil. The feces of oligochaetes are voided as worm casts containing semi-digested organic material of organismal origin/urine and soil and thereby enrich the agriculture fields (Dash, 1987). The egestion rate ranges from 100 to 460% of body weight/d for three tropical earthworm species (Dash et al., 1984, Dash et al., 1986). The worm cast production also ranges from 2 to 247 t/ha in different sites and thereby brings up layers of soil between 1 mm and 5 cm (see Dash, 1987). In aquatic oligochaetes, the quantification of F is difficult and methods adopted for the estimation of F have yielded widely different values. For example, in the inverted method developed by Appleby and Brinkhurst (1970), the worms are kept in an

Figure 1.7 Energetic components in annelids. C = Consumption, F = Feces, U = Urine, R = Respiration, M = Mucus, P = Production, S = Somatic growth, G = Gonad growth, C1 = cocoon, Ab = Absorption, As = Assimilation. Due to dual sexual systems and cocoon production, clitellates invest relatively less on gamete production than the gonochoric polychaetes.

24  Reproduction and Development in Annelida

Figure 1.8 A. Defecation rate of Limnodrilus hoffmeisteri as estimated by upright and inverted methods (drawn using data reported by Kaster et al., 1984). B. Absorption efficiency of polychaetes as function of food nitrogen content. ■ Observed values, ● Predicted values (drawn from protocol record of T.J. Pandian).

inverted (posterior end down, anterior end up) position but in that described by Kaster et al. (1984), they are kept in normal oriented upright position. Figure 1.8A shows that F estimates for the tubificid Limnodrilus hoffmeisteri averages to 0.4 mg (dry) F/mg (dry) worm/h in the inverted method but 0.6 mg/g/h in the upright method. The latter value on F is 50% more than that of the former. In general, the recovery of F of aquatic animals is not only cumbersome but also incomplete for following reasons: (i) loss of soluble material, (ii) loss owing to decomposition, (iii) difficulty in distinguishing food from F especially in deposit-feeding polychaetes (e.g. Pectinaria gouldii, Gordon, 1966) and (iv) chances of F being reingested by polychaetes, which switch from suspension- to deposit-feeding (e.g. Nereis diversicolor, Harley, 1950). Hence, the need for an easier indirect method to estimate absorption efficiency, Ae (A = C – F = absorption, Ae = A/C × 100) is obvious. Pandian and Marian (1985b) have considered nitrogen (N) content of C as an indicator of Ae. Considering 11 relevant publications, which have reported adequate information of N content of C ranging from 0.4% (sediment, Pollack, 1979) to 9.8% (95% Mytilus edulis + 5% agar, Neuhoff, 1979), they have found an almost linear relationship between N content of food and Ae (Fig. 1.8B). The obtained trend includes Ae values reported for Arenicola marina feeding on sediment containing 0.4% N to Nereis virens fed on mussel tissues containing 9% N. The values are also related to polychaetes of different weight classes (1 to 1,156 mg) and exposed to different temperatures (12 to 29ºC). The contributions made by variations in Ae by temperature and body size on N-Ae relationship are negligible. Earlier, Tenore (1983) had considered two types of Spartina alterniflora detritus containing 0.9 and 1.0% N but

Introduction  25

4.45 and 2.61 Kcal/g dry weight, respectively and found that energy content rather than N content of food determines the incorporation of S. alterniflora detritus by Capitella capitata. However, he has not estimated Ae of C. capitata fed on these detritus. Incidentally, the indirect method of N content of food as a marker of Ae holds good for fishes, aquatic insects and reptiles (Pandian and Marian, 1985c, d, e). Urine (U): In polychaetes, the main excretory products are ammonia, urea and uric acid (Pandian, 1975). The terrestrial oligochaetes excrete ammonia and urea and their urine contains some mucus protein (Bahl, 1947). However, quantitative value for the substance or energy content of U for annelids is not yet available. The only available value seems to be that reported for blood sucking Haementeria ghilianii. In this leech, Sawyer et al. (1981) have estimated the ingested blood retained after urine elimination. Considering their consecutive three values, the substance lost on U by the leech accounts for 23.2% of C. Respiration (R): As in other animals, oxygen uptake ultimately liberates biologically useful energy for body functions in annelids too. The fraction (C – [F + U] = As) of assimilated energy ranges from 55 to 75% in annelids. On an average, ~ 65% of the As is used for metabolism (R) in aquatic polychaetes (Cammen, 1987) and terrestrial oligochaetes (Dash, 1987), leaving ~ 35% of As for body growth. A reason for the high (75%) R value is that some annelids switch over to energetically less efficient anaerobiosis under oligoxic/anoxic conditions.

1.6.2 Anaerobiosis Many polychaetes utilize mostly glycogen and various amino acids as a major energy source (Hochachka et al., 1973). In many polychaetes, the simultaneous catabolism of glycogen and amino acids maintains the redox balance in the cell during anaerobiosis. The first phase of glucose oxidation in anaerobic glycolysis is through the Embden-Myerhoff-Parnas (EMP)pathway. In this pathway, each molecule of the 6'-C glucose is degraded to two pyruvate molecules, which are then converted into lactate. In some animals, the duration of anaerobiosis is too short to permit accumulation of lactate without excess reduction in pH of tissue fluids. When a favorable condition returns, a considerable fraction of the accumulated lactate is converted back into pyruvate and oxidized to water and CO2. This oxidation results in an increased ‘oxygen demand’ by tissues over normal metabolic requirement and constitutes the so called ‘oxygen debt’ (Neoamphitrite figulus, Theede, 1973). To prevent lethal reduction of the pH of the body fluids, pyruvate may, however, be excreted (e.g. Nereis spp, Schottler, 1979) and/or reutilized by gluconeogenesis (e.g. Nereis pelagica, Theede, 1973). Not only pyruvate but also other end products are excreted as such. For example,

26  Reproduction and Development in Annelida

propionate and acetate excreted by the anaerobic Arenicola marina is in the range of 18–27 µmol/g live weight/d (Surholt, 1977). When anaerobic duration is prolonged in the facultative anaerobic and obligate anaerobic worms, the carboxylation of pyruvate yields oxaloacetate (OXA), which is converted to fumerate and finally to succinate through the malate route. In this new pathway of glucose degradation and phosphoenolpyruvate (PEP) to OXA, succinate (e.g. swamp worm Alma emini, Mangum et al., 1975) and alanine are produced as major end products of prolonged anaerobiosis (see Pandian, 1975) and finally to volatile fatty acids (propionate being the major product) and acetate (Schottler et al., 1983). For example, the opheliid Euzonus mucronata can survive the periodic exposure to anoxic condition for 18–20 days, producing succinate and propionate as major end products of glycolysis (Ruby and Fox, 1976). Metabolic pathways terminating with succinate and propionate have two advantages over the EMP glycolytic pathway: 1. They produce twice as much ATP/mol substrate and reduce by 50% the accumulation of potential toxins. 2. On restoration from anaerobiosis, succinate and propionate can readily be converted to OXA, which directly leads into the Krebs cycle, while lactate must first be converted to pyruvate and then to OXA before entering the Krebs cycle (Cammen, 1987). Growth (P): Of ~ 35% of As energy stored in the body for growth, small or larger fractions are used for body growth (S), reproductive output (G) and mucus (M). The serpulids construct a calcareous tube using organic matrix and divert as much as 65% of As for tube construction (e.g. Mercierella enigmatica, Dixon, 1980). The sabellids secrete mucus for construction of a sand tube. Filter feeding polychaetes also secrete mucus to capture food particles. Of the total mucus secreted by M. enigmatica, 94% is secreted by the food capturing structure, the ‘crown’ and the remaining 6% used by the body surface and gut, i.e. each 3%. The mucus of M. enigmatica consists of mostly mucopolysaccharides (Dixon, 1976). Yet, no estimate has been made on the fraction of As secreted as M in the energy budget of M. enigmatica. However, the nitrogenous excretion of earthworms is through mucus. Their urea consists of 64% mucus, 16% urea and 20% ammonia. Dash and Patra (1979) have estimated that the quantum of excretion by the earthworms Lampito mauritii and Ocnerodrilus is in the order of 142.3 g mucus/m2/y, amounting to 6.6 g N/m2/y; evidently, the chemical nature of mucus of the earthworms is different from that of polychaetes. Reproduction (G): For the terrestrial oligochaetes, assimilated energy allocated for reproduction averages to < 2% (Dash, 1987). Corresponding values are not available for the aquatic oligochaetes and polychaetes. Values on the fraction of P energy allocated for R ranges from 12% in Harmothoe imbricata (Gremare and Olive, 1986) to 30–40% in many other polychaetes (Cammen, 1987). A single value reported for the serpulids M. enigmatica indicates that 12% of the

Introduction  27

assimilated energy is allocated for G, the gamete production (Dixon, 1976, 1980). Understandably, the broadcast spawning, gonochoric polychaetes allocate a greater fraction of the assimilated energy for reproduction than the simultaneous hermaphroditic oligochaetes, which ensure a relatively higher survival of their progenies through internal fertilization and safer deposition of developing eggs within the protective cocoon. In fact, the increased cost of developing and maintenance of dual reproductive systems reduces energy allocated for G by 50% in hermaphroditic polychaetes too. For example, the number of eggs spawned by seven polychaete species of Ophryotrocha averages to 9.4 eggs/d and 4.45 eggs/d for gonochorics and hermaphrodites, respectively (calculated data from Premoli and Sella, 1995). Apparently, the oligochaetes have chosen to invest a greater fraction of assimilated energy to meet the costs of dual reproductive system and cocoon.

1.7  Life Span and Generation Time Generation Time (GT) is the duration of time required by an animal from its egg stage to egg releasing stage. The investment on GT as a percentage of Life Span (LS) indicates the remaining fraction of LS for investment on reproduction. Hitherto, annelidan reproductive biologists seem to have not paid adequate attention on GT/LS. As a result, even the available bits and pieces of information are widely scattered. Incidentally, there is time lag between sexual maturity and oviposition/spawning. For example, the lag between the appearance of clitellum, a morphological marker of sexual maturity in clitellates, and oviposition ranges from 6 days in Perionyx excavatus to 19 days in Metaphire houlleti (Joshi and Dabral, 2008). Timm (1984) is perhaps the first to consolidate the relevant information on potential age for 36 aquatic oligochaete species. From Table 1.7, the following may be noted: 1. The 20 criodrilid, lumbriculid, naidid and tubificid species cultured at prevailing laboratory temperature underwent sexual reproduction only (cf Table 4.1). The estimated maximum age was 7.0, 9.5, > 11 and 14 years for the lumbriculid Stylodrilus heringianus, tubificid Tubifex tubifex, naidid Spirosperma ferox and criodrilid Criodrilus lacuum, respectively. 2. In them, the maximum age was decreased in the following order: criodrilid < naidid < tubificids < lumbriculid. 3. The age was consistently shorter for these worms (except in Potamothrix moldaviensis) from the fields, where they underwent sexual and/or asexual reproduction. However, no description was given as to how the age of these worms, especially in the fragmenting ones was determined. 4. The maximum age was also decreased in all the species, when reared at 20–25ºC. In asexually reproducing naidids Stylaria lacustris, the doubling duration decreased from 11.1 days at 10ºC to 5 d at 20–25ºC (Schierwater and Hauenschild, 1990). Further, the annual potential

28  Reproduction and Development in Annelida

Table 1.7 Potential life span (y) of representative aquatic oligochaetes. In laboratory, reproduction was exclusively sexual. In the fields, seasonal temperature ranged from 5ºC during winter to 15ºC during summer (from Timm, 1984, modified) Species

Field

Laboratory

At 20–25ºC

> 14

–

> 11 > 8 5 ~ 3.5 ~ 4.5

8 ~4 3 3.5 2.5

Criodrilidae Criodrilus lacuum

– Naididae

Spirosperma ferox Psammoryctides barbatus Ilyodrilus templetoni Potamothrix hammoniensis P. moldaviensis

> 12 > 12 8 ~ 4.5 ~ 3.5 Lumbriculidae

Stylodrilus heringianus Rhynchelmis limosella

> 12 > 6.5

7 3

3.5 –

9.5 6.3 5.5

3.8 4.3 4.2

Tubificidae Tubifex tubifex* T. tubifex** T. tubifex***

> 11 8.5 6.5

* from limnophilus, Roosna Alliku spring, **from reophilus, *** from limnophilus, Lake Peipsi-Pihva

number of generations of S. lacutris decreased from 30 with unlimited food supply to 20, 14 and 12, when food supply was limited to 50, 20 and 10%, respectively. These values were 42, 24, 13 and 9 for Nais sp and 66, 39, 23 and 18 in Chaetogaster diastrophus (Lohlein, 1999). Consequent to the differences in temperature and food supply in the fields, the mean maximum age of these worms was ~ 4 years only. Yet the LS of annelids in the natural fields, where they are subjected to an intense predation, ranges between a few days and months. Table 1.8 summarizes available information on LS and GT of representative hirudineans, earthworms, terrestrial and aquatic oligochaetes and polychaetes. It includes information for gonochoric (Dinophilus gyrociliatus) and hermaphroditic species as well as sexually reproducing (e.g. leeches) and/ or asexually reproducing oligochaetes and polychaetes. The following may be noted: 1. Adequate information on gonochoric polychaete is not yet available. 2. There are fast and slow growing enchytraeids (e.g. Marionina clevata, Springett, 1970) and tubificids characterized by short (semelparous) and long (iteroparous) (e.g. Tubifex tubifex, Limnodrilus hoffmeisteri, Poddubnaya, 1984) LS. Despite these constraints, the data assembled in Table 1.8 permit for the first time the following new findings: The percentage investment on GT is decreased from 40–46% in large (10–23 g) sanguivorous leeches to ~ 30% in

Neanthes limnicola

Capitella capitata

Streblospio benedicti

Dinophilus gyrociliatus

WW

O. diadema YY, YW

~ 8 mm

< 1 cm

5 mm

O. adherens

> 60

~ 35

~ 35

♂ , Boreal 1–2ºC + ♂ +

Fong and Pearse (1992)

♀, ♂, viviparous, California

548

171

31

Table 1.8 contd. ...

Martin and Bastida (2006)

47

♀, ♂, sexual, Argentina

195

Levin and Bridges (1994)

♀, ♂, lecithotrophic 32

13.5

43 420

Levin and Bridges (1994)

♀, ♂, planktotrophic

Akesson and Costlow (1991)

Akesson (1982)

Paavo et al. (2000)

Ocklemann and Akesson (1990)

Bouguenec and Giani (1989)

Tondoh (1998)

Fernandez et al. (2010)

Gonochoric

♂ + ♂ + ♂ +

Boreal at 10ºC, ♂ +

♂ +

Twins, earthworm, Spain

Singleton, earthworm, Spain

25

15

14

10

15

5

Polychaetes

10

22

34

29

Davies and McLoughlin (1996)

Khan (1982)

Reference

9.5

11

40

46

Remarks

Terrestrial oligochaetes

~ 26

30

GT/LS (%) Hirudineans

38

70

245

350

196

> 1098

3 mm

Ophyrotrocha socialis

165

490 56

142

280

446

GT

490

254

2.8 g

Aporrectodea trapezoides

713

Enchytraeus variatus

23 g

Hirudo medicinalis

955

196

10 g

Johanssonia arctica

LS

Hyperiodrilus africanus

Size

Species

Realized life span (LS) (d) and generation time (GT) of some annelids

Table 1.8

Introduction  29

< 25 mg

T. tubifex (Italy)

Limnodrilus hoffmeisteri

T. tubifex (Boreal)

200

~ 2.3 mg

Tubifex costatus

62 26

90

349

19

54

30

20

19

10

15

93

72

380

GT/LS (%)

Remarks ♂ + ♂, Naidid + ♂ + ♂, Marine, sexual + asexual + ♂, Sexual/Parthenogenic + ♂, Sexual/Parthenogenic + ♂, Sexual/Parthenogenic + ♂, Sexual/Parthenogenic + ♂, Sexual/Parthenogenic +

Aquatic oligochaetes

150

52

60

60

12

35

20

GT

95

200

62

364

< 2 cm

Enchytraeus variatus

60

LS

Branchiura sowerbyi

Size

Lumbricillus rivalis

Species

...Table 1.8 contd.

Poddubnaya (1984)

Pasteris et al. (1996)

Brinkhurst (1964)

Bouguenec and Giani (1989)

Lobo and Alves (2011)

(see Lindegaard et al., 1994)

Reference

30  Reproduction and Development in Annelida

Introduction  31

medium sized (2.8 g) sediment/detritus-feeding earthworm, to ~ 19–30% in tubificids and to 5–15% in small (2–5 mm body length) terrestrial oligochaetes and hermaphroditic polychaetes. However, it is not clear whether the GT is determined by other factors like (i) LS (from a few days in aquatic oligochaetes to a few years in leeches), (ii) feeding habits (detritivore, herbivore [naidids] and sanguivore) or (iii) egg size (ranging from a few µm to mm, p 73). One or more of the above cited factors may determine GT. In this area, researches are required. Of course, GT may be regulated by juvenile hormone (JH) and the like in crustaceans (Pandian, 2016).

1.8 Gametogenesis In polychaetes, for example in temperate Kefersteinia cirrata, oogenesis commences with the accumulation of primary oocytes and proceeds through prophase to deplotene. At this stage, the progress of meiosis is arrested and oocyte nucleus expands to form the germinal vesicle. After a period, vitellogenesis begins usually during autumn-winter months— as in many temperate echinoids (see Pandian, 2018). Oogenesis as well as spermatogenesis and spermiogenesis are completed between December and February and the worm is ready for spawning by spring (Olive and Pillai, 1983). Oogenesis: In polychaetes, two patterns of oogenesis have been described (Eckelbarger, 1983, 1986). In the intra-ovarian pattern, almost the entire process of oogenesis including vitellogenesis occurs within the fairly large and structurally complicated ovary. The maturing oocytes receive yolk precursors from (i) closely associated nurse cells (abortive or sibling oocytes), (ii) follicle cells, (iii) closely associated circulatory system or (iv) a combination of these sources. In about eight families (e.g. Capitellidae, Sabellariidae, Orbiniidae, Eckelbarger, 2005), in which the intra-ovarian oogenesis is reported to occur. There are significant differences regarding the type of precursors utilized during vitellogenesis, their metabolic pathways and chemical nature of yolk bodies themselves. Besides, there is no apparent relationship between the ovarian type and oogenic mode, egg size or larval development mode. In the extra-ovarian pattern, the ovary is small, structurally simple and transient in nature. The small pre-vitellogenic oocytes are released either solitarily (e.g. Sabellidae, Serpulidae, Oweniidae, Glyceridae) or in clusters into the fluid-filled coelom; they may also be released as follicles into the coelom (Alciopidae, Nereididae, Phyllodocidae, Terebellidae, Cirratulidae, Ampharetidae, Pectinariidae). The clusters may subsequently be separated, as in Sphaerodoridae and Pholoidea or rarely remain intact in some syllids, tomopterids and onuphids until vitellogenesis is completed. The

32  Reproduction and Development in Annelida

oocytes may (i) develop without an association with accessory cells, (ii) be accompanied by [a] coelomocytes specialized for triglyceride synthesis or [b] eleocytes involved in yolk precursor production, or (iii) be closely associated with nurse cells playing nutritive role (Eckelbarger, 2006). In Nereis virens, for example, the eleocytes extra-ovarially synthesize yolk protein precursors, the vitellogenins and subsequently transport into the oocytes by receptormediated endocytosis (Hafer et al., 1992). Spermato- and Spermio-geneses: Typically, gonial cells, produced from the testes, are dropped into the coelomic cavities (seminal vesicles), where they undergo a series of variable but species specific number of mitosis prior to entry into meiosis. In oligochaetes, spermiogenesis involves (i) nuclear shaping, (ii) condensation of chromatin, (iii) formation of acrosome, (iv) reduction in the number of mitochondria and (v) development of a long flagellum (Ferraguti, 1984). A typical sperm consists of (a) an apical acrosome with an acrosomal tube in clitellates but not in other annelids, (b) a condensed thin nucleus, (c) two (e.g. Tubifex) to eight (e.g. Spargnophilus) tightly packed straight (e.g. Lumbricus) or coiled (e.g. Phreodrilus) mitochondria (d) a mid-piece (Erseus, 1999) (absent in polychaetes, however see Blake and Arnofsky, 1999) (Fig. 1.9) and (e) a central long, thin flagellum. Shaping of the sperm head differs between broadcasting polychaetes and brooding sabellids/serpulids. Of 10 species investigated, seven brooding species belonging to Sabellidae, Serpulidae and Fabricinae have an elongate head and the remaining three broadcasting serpulids spherical head. Among 23 sabellinaeids, not only the 11 broadcasters but also five brooding species have spherical head. Briefly, broadcasting polychaete species consistently have spherical head

Figure 1.9 Representative sperms of some polychaete families. A. Eunicidae, B. Sabellidae, C. Phyllodocidae, D. Terebelliformiae, E. Spionidae, F. Scolecidae, G. Cirratuliformia and H. Amphinomidae (rough sketches made from Rouse, 2006).

Introduction  33

but the brooders may have elongate or spherical head (Rouse, 1999). The vestimentiferan polychaetes have attracted studies on sperm ultrastructure. In Riftia pachyptila, Ridgeia piscesae and Siboglinum ekmani, the sperm structure is essentially similar. Their mature sperm are filiform with an elongate nucleus and flagellum. Two to three mitochondria wrap around the nucleus. The acrosome is helical and lies at the apex of the nucleus in S. ekmani but on the side of nucleus in R. pachyptila (see Rouse, 1999). Notably, the sperm are aflagellate in members of Alvinellidae, Histriobdellidae and Pisionidae (Westheide, 1988). Sperm of Ophryotrocha are immotile and resemble spermatids (Pfannenstiel and Grunig, 1990). Parasperm: In a few eccentric species of aquatic of tubicine oligochaetes, two types of spermatozoa are produced (Ferraguti, 1984): (i) fertilizing eusperm with regular haploid DNA content and (ii) protective and transporting parasperm with a much reduced DNA content. The spermatogonia are released from the testis into the coelomic cavity of the tenth segment (seminal vesicles) as cysts, each consisting of a group of four cells interconnected by cytoplasmic bridges. The cysts undergo three spermatogonial divisions and form 8, 16 or 32 cells. In the euspermic line, primary spermatocytes and euspermatids are produced through regular meiosis, giving rise to 128-cell cysts. On the other hand, paraspermatid cysts are formed by a much greater number (550–3,000) of cells. Consequently, they are larger (100–150 µm) than euspermatid cysts (70–80 µm). A mitotic spindle is never formed in these fragmenting cysts of paraspermatocytes. As a result, they undergo a peculiar nuclear fragmentation. During this fragmentation, the DNA is distributed unevenly among spermatids, giving rise to a great and variable number of parasperm with variable DNA contents (Boi et al., 2001). The eusperms are characterized by the presence of acrosome, smaller mitochondria (half the size of parasperms), long (30 µm) body (3 µm in parasperm) and shorter tail. Following the gametic exchange between hermaphroditic partners, sperms are bundled as spermatozeugmata in spermatheca. The bundles are made up of parallel core of eusperm surrounded by parasperms. The observations of Boi and Ferraguti (2001) indicate the commencement of spermatogenesis on the third week and appearance of spermatids on the sixth week. The peaks of spermatid appearance are in successive waves and sperm production is a cyclic event. In members of Arenicolidae, Maldanidae, Syllidae and Terebellidae, sperms are bundled into spermatozegumata. In many other annelids, the sperms may be tightly packed into a spermatophore, which is a capsule containing sperms with their head toward the center. In polychaetes, spermatophores have been reported from Spionidae (e.g. Spio filicornis, Greve, 1974), Capitellidae, Arenicolidae, Hesionidae, Syllidae and Histriobdellidae, Nerillilidae, Protodrilidae and Myzostomidae (see Schroeder, 1989). Though a spermatophore can be a device to ensure fertilization of eggs during cocoon deposition, its function is doubtful. In fact, the experimental

34  Reproduction and Development in Annelida

Table 1.9 Effect of spermatophore removal on reproduction in the earthworm Eisenia foetida (from Monroy et al., 2002, modified and added) Traits

Spermatophore Intact

Removed

Cocoon (no./worm)

15.7

13.9

Cocoon size (mg)

166

146

Cocoon viability (%)

68

81

21.7

25.9

Cocoon survival (no./worm) Hatchling (no./worm)

64

67

Hatchling size (mg)

10.7

11.3

Hatchling (no./cocoon)

2.0

2.3

removal of spermatophore in the lumbricid Eisenia foetida slightly increases the reproductive success (Table 1.9). Calculations of cocoon survival and hatchling per cocoon in the worm indicate that in each cocoon 12% of additional eggs have been fertilized after the removal of spermatophore. Sperm types: In polychaetes, the sperms are grouped into three types: (i) ectaquasperm, (ii) ent-aquasperm and (iii) introsperm (Jamieson, 1986). The ectaquasperms are liberated freely into the ambient water, in which fertilization with con-specific eggs occurs. In about 30 species, it is characterized by (a) small cylindrical acrosome resting in a depression on the anterior end of the nucleus, (b) spherical or ovoid nucleus, (c) a few rounded cristate mitochondria and (d) a free axoneme with the 9 + 2 arrangement of microtubules (Blake and Arnofsky, 1999) (e.g. Amphinomidae: Eurythoe complanata, Sabellidae: Idanthyrsus pennatus, Serpulidae: Galeolaria caespitosa, Chaetopteridae: Chaetopterus variopedatus). The ent-aquasperm is also shed into the ambient water but it is drawn in by the inhalant or feeding current of the female/hermaphrodite. In them, internal fertilization invariably occurs (e.g. Sabellidae: Fabricia, Oriopsis, Maldanidae: Micromaldane). In contrast, the introsperm never enters the water in aquatic species and occurs in all terrestrial annelids. Of > 28 polychaete species, Blake and Arnofsky (1999) have found the confirmed or probable coexistence of spermatophore and introsperm in 20 species and lack of spermatophore in the remaining eight ecto-aquaspermic species. Introsperm may be transferred from male to female by copulation in some hesionid and saccocurid polychaetes (see Schroeder, 1989) but by pseudocopulation, in which the apposed male and female shed gametes directly into a ‘cocoon’ in dorvillid Ophryotrocha with aflagellate sperms; however, sperms transferred into the spermatheca of a mating partner are subsequently shed into the eggs contained within a cocoon in a few polychaetes and most oligochaetes (Jamieson, 1986). Within the interstitial polychaete genus Microphthalmus, Westheide (1967, 1979) describes

Introduction  35

a gradation of simple sperm transfer by spermatophore in M. aberrans to hypodermic impregnation into the female opening of the receptacular tissue in M. listensis. In still others, sperms, as in earthworm Lumbricus terrestris (Koene et al., 2005) or spermatophore, as in some hesionids (e.g. Westheide, 1967) and leeches (see Jamieson, 1986) are transferred directly into the body surface of the mating partner and the sperm pierce through the body wall to reach the eggs. In L. terrestris, 40–44 copulatory setae pierce into the partner’s skin causing damage, while injecting the semen drawn from its setal glands. Spermatheca: Named as seminal receptacles in polychaetes and uterus in hirudineans, the spermatheca serves to receive and store sperms/ spermatophores. For detailed description of its structure, Adiyodi (1988) may be consulted. In oligochaetes, the spermatheca is generally paired saccular organs with their ducts opening to the outside. Their number ranges from zero in parthenogenic lumbricid Bimastos to one pair (Naididae, Enchytraeidae, most Tubificidae and Moniligastridae) to seven pairs in megascolecid Perionyx polytheca. In hirudineans, they are a bilobed but unpaired structure. In some polychaetes (e.g. spionid Polydora ligni), the sperms are stored in pockets of female nephridium (see also Rice, 1980). The stored spermatozoa remain motionless for varying periods and may be compacted into a cylindrical bundle called spermatozegumata. In polychaetes, the stored sperms may undergo some morphological changes, suggestive of a capacitation-like process (e.g. Pisione alikunhi, Alikunhi, 1951) and P. remota (Strecher, 1968).

1.9  Reproductive Modes and Dispersal With a soft body and low motility, annelids are prone to predation. Unlike molluscs with external protective shell(s), they do not have structural and/ or chemical (however see p 48–49) defense mechanism to avoid predation. As a consequence, a key driving force in their evolution and speciation seems to have been an extraordinary diversification of reproductive modes. Interestingly, gonochorism in consort with multiplication of reproductive modes has facilitated a greater speciation in polychaetes with ~ 13,000 species. Conversely, the consistent presence of hermaphroditism and/or parthenogenesis along with internal fertilization and direct development within the cocoon in both aquatic (e.g. tubificids) and terrestrial (e.g. earthworms, see Fig. 8.10) ‘herbivorous oligochaetes’ has reduced species diversity to ~ 2,000 species (see Table 1.2). Carnivory/sanguivory has further reduced species diversity to ~ 800. It is in this context, reproductive modes in annelids become important and interesting. Polychaetes display fascinating and incredibly diverse reproductive modes (Fischer and Fischer, 1995). Broadly, the eggs may be freely spawned or brooded. For the first time, Wilson (1991) made an extensive survey of

36  Reproduction and Development in Annelida

taxonomic distribution of these reproductive modes across 307 species belonging to 36 families and 10 orders. He brought them under the following six groups: 1. Free spawned (fs) eggs, 2. Freely released embryos encapsulated in gelatinous mass (gel), 3. Brooding eggs outside the body, say, on substratum (br–ext), 4. Eggs brooded inside the tube (br–tube), 5. Brooding encapsulated embryos inside the tube (br–enc–tube) and 6. Eggs brooded inside the body (br–int). The first group was further divided into three sub-groups namely 1. fs into (i) with feeding and dispersing planktotrophic (PLK) larvae, (ii) with dispersing lecithotrophic (LEC) larvae and (iii) direct developers (DIR). Each of the remaining five groups were divided into 2. gel into (i) PLK, (ii) LEC and (iii) DIR, 3. br–ext into (i) PLK, (ii) LEC and (iii) DIR, 4. br–tube into (i) PLK, (ii) LEC and (iii) DIR, 5. br–enc–tube (i) PLK, (ii) LEC and (iii) DIR as well as 6. br–int into (i) PLK, (ii) LEC and (iii) DIR. In all, as many as 18 reproductive modes were recognized. Expectedly, Wilson’s report was more of taxonomic. Analysis of his data on a holistic polychaete level, a different picture emerged (Table 1.10), from which the following may be inferred: 1. Surprisingly, > 47% of polychaetes brood their eggs/embryos; the remaining 53% freely spawn their eggs (40%) or release embryos in encapsulated jelly mass (13%), 2. Of 307 species surveyed, 126 species, i.e. 41% of them develop through a feeding and dispersing PLK stage during the period of indirect development, 3. Another 23.5% (72 species) of them are LEC; hence they pass through the short larval period of dispersal in the pelagic realm, 4. Of the remaining 35.5% undergo direct development. Incidentally, Blake and Arnofsky (1999) listed the number of spionids that (i) broadcast thick eggs and (ii) brood thin eggs. An estimate of these two Table 1.10 Distribution of reproductive modes in polychaetes (from Wilson, 1991, modified). PLK = Planktotrophic, LEC = Lecithotrophic, DIR = Direct development; all values in parantheses are in % Mode

PLK

LEC

DIR

Total 123 (40.0)

Freely spawned eggs Freely spawned (fs)

79

34

10

Gel encapsulated (gel)

10

16

13

39 (12.7)

89 (28.9)

50 (16.3)

23 (7.5)

162 (52.7)

Subtotal

Brooded eggs External (br-ext)

6

7

15

28 (9.1)

Tubular (br-tube)

7

11

39

57 (18.6)

Encapsulated tube (br-enc-tube)

24

3

15

42 (13.7)

Internal (br-int)

0

1

17

18 (5.9)

Subtotal

37 (12.0)

22 (7.2)

86 (59.3)

145 (47.3)

Total

126 (41.0)

72 (23.5)

109 (35.5)

307

Introduction  37

Table 1.11 Development and dispersal of polychaete larva. A resurvey of information presented in Appendix of Carson and Hentschel (2006). E = Epitokous, MA = Mobile adults, AS = Asexual Item

Development Mode

Subtotal (no.)

1.  Indirect development 1.1  Free spawner (fs), planktotrophic (PLK)

1.1.4 1.1.5 1.1.6 1.1.7 1.1.8 1.1.9 1.1.10 1.1.11

Pelagic, PLK PLK, Trochophore E, Benthic spawning, pelagic PLK > 3 w- mo fs, PLK, mitraria fs, PLK, rostraria fs, PLK, aulophore Pelagic, PLK, MA Pelagic, PLK, AS fs, LEC, larval duration > 10 d Pelagic, gel. Encapsulated Pelagic, gel. tube

1.2.1 1.2.2 1.2.3 1.2.4

E, LEC fs, LEC, pelagic for < 7 d fs, LEC, pelagic for < 7 d + rafter pelagic, LEC, MA

1.3.1 1.3.2 1.3.3

bs bs, MA E, benthic egg mass

1.1.1 1.1.2 1.1.3

6 2 1 24 5 1 2 1

High High High High High High High High 56

8 43 3 2

Tube brooded egg mass Brooded, pelagic for 1 d Brooded, direct development Tube brooded, direct development, MA E, brooded, benthic, AS Internal/External brooding Brooded benthic larva

8

2.2.5 2.2.6 2.2.7

Medium Medium Medium Low

82 33 4 1 20 1 3 4

2.2  Brooded but no pelagic PLK

Medium High Medium Medium Medium

3 2 3

2.  Brooded development 2.1  Brooded + pelagic PLK

2.2.1 2.2.2 2.2.3 2.2.4

High/Medium High High High High

1.3  Benthic spawner (bs)

Internal pelagic, PLK Brooded PLK for > 13 d Brooded but pelagic PLK larva Externally brooded PLK ~ 8 d Brooded but released at 3 setiger Brooded, pelagic LEC

Dispersal

173 109 35 20 12

1.2  Free spawner (fs), lecithotrophic (LEC)

2.1.1 2.1.2 2.1.3 2.1.4 2.1.5 2.1.6

Total (no.)

Medium/Low Medium High High Medium Medium Medium Medium

49

Low

8 1 2 12

Low Low Low Low

10 15 1

Low Low Low Table 1.11 contd. ...

38  Reproduction and Development in Annelida

...Table 1.11 contd. Item

Development Mode

Subtotal (no.)

3.  Direct developers (Dd) 3.1 3.2 3.3 3.4 3.5 3.6 3.7 3.8 3.9 3.10

Egg mass, short swimming, neochaete larva Eggs in jelly mass, MA Egg mass Direct developers Dd, presumed no pelagic stage Dd, MA Dd, MA, AS Brooded up to direct development LEC Viviparous

Total (no.) 99

Dispersal Low

5

Medium

1 33 7 14 5 25 2 3 4

Medium Low Low Low Low Low Low Low Low

groups suggests that 47% of them brood their eggs and 53% are broadcast spawners. Understandably, the sedentary tubiculous spionids opt more for brooding rather than broadcast spawning. Unlike brooding limited to embryos and release of the larvae in crustaceans, some brooding annelids (e.g. 82 species Table 1.11) continue to brood the larvae too. The larvae are released from three up to 15–24 chaetigers stage. In polychaetes, the planktotrophic larval duration may last to 50 days or longer. In Fig. 1.10A, relevant data reported by Blake and Arnofsky (1999) on egg size of some spionids are plotted against larval duration. Unusually, the duration increases with increasing egg size, although individual values are scattered. Understandably, the larvae arising from larger eggs tend to postpone the departure from brood at 3rd to 15th–24th chaetiger stage (Fig. 1.10B). On plotting the values reported by Prevedelli and Simonini (2003) for body size (length) of three nereidids, a dorvilleid and a dinophylid against egg size, a direct relationship becomes apparent (Fig. 1.10C). At interspecies level, the same holds true for the relationship between body size and fecundity for free spawners and brooders (Fig. 1.11). Rouse and Fitzhugh (1994) have listed values for body size, egg size and fecundity of some sabellids that broadcast spawn and brood eggs intra-tubularly or extra-tubularly. On plotting mean values for the spawners and brooders, the emerging trends indicate that egg size increases with increasing body size but fecundity decreases with increasing body size (Fig. 1.10C). Hence, the limited available information in sabellids seems not to fall in line with that of nereidids and others. Incidentally, more information is available on the effects of body size on fecundity and egg size of brooders and spawners belonging to sabellinids and fabricinids (Rouse and Fitzhugh, 1994). To establish phylogenetic relation among brooders and spawners in the subgroups,

Introduction  39

Figure 1.10 Effects of egg size on A. planktotrophic duration and B. release of chaetiger stage in polychaetes (drawn using data compiled by Blake and Arnofsky, 1999). C. Effects of body size on egg size (▲▁▲) and fecundity (△▁△) in 3 nereid species (1 = Marphysa sanguinea, 2 = Perinereis rullieri, 3 = P. cultrifera), a dorvilleidid (4 = Ophryotrocha labronica) and a dinophilid (5 = Dinophilus gyrociliatus) (drawn using data reported by Prevedelli and Simonini, 2003). Effect of body size (length) on egg size (■▁ ▁ ■) and fecundity (□ ▁ ▁ □) of broadcast spawning (x, mean of 6 species), extra-tubular brooding (x, 3 species) and intra-tubular brooding (x, 3 species), terebellids (drawn using data compiled by McHugh, 1993). Effects of body volume on D. egg volume, E. fecundity and F. total egg volume in broadcast spawning, extra-tubular brooding, intra-tubular brooding fabricinid and intra-tubular brooding sabellinid polychaetes (modified from Rouse and Fitzhugh, 1994).

40  Reproduction and Development in Annelida

Figure 1.11 Fecundity as function of body length in A. free spawning and B. brooding polychaetes. Note the values shown by ● are simultaneous hermaphrodites, ■ gonochorics and □ with feeding larvae (drawn using data reported by Kupriyanova et al., 2001 and also data (▲) reported by Prevedelli and Simonini (2003) are also included).

an attempt has been made by Rouse and Fitzhugh, who have found that body size (volume) holds linear relations at different levels with egg size (Fig. 1.10D). However, linear trends for the relation per se become apparent (Fig. 1.10D), when values reported for species with unknown life history are eliminated and statistical analysis is not applied. Accordingly, the level and degree of the slopes of these trends differ. Notably, the slope decreases in the following order: broadcast spawners < sabellinid intra-tubular brooders < extra-tubular brooders < fabricinid intra-tubular brooders. This is also true for the relations between body size vs fecundity (Fig. 1.10E) as well as body size vs egg size (Fig. 1.10F). It must be noted that all these linear relationships are based on log values. Different trends may become apparent, when the normal values are considered (see p 75). Remarkably, sabellinids larger than log 1.33 mm3 alone are broadcast spawners. Fabricinids smaller than log – 1.67 mm3 are all intra-tubular brooders. Both extra-tubular as well as intra-tubular sabellinids and fabricinids have a wide size range from log – 1.33 to log 3.0 mm3. It is not known whether the presence of nurse eggs/larvae in the intra-tubular brooding is responsible for the widening of the size range in these polychaetes. Interestingly, Garraffoni et al. (2014) have also listed available values for egg size, fecundity and adult size of terebellids characterized by indirect (seven species) or direct (five species) life cycle. The mean values for the direct developers are 0.1 mm3 for egg size, 242 eggs for fecundity and 8,608 mm3 for adult size, in comparison to 0.006 mm3, 408 eggs and 10,488 mm3 for egg size, fecundity and adult size of indirect developers, respectively. Although, the differences between these mean values are justifiable for terebellids with direct and indirect life cycles, the individual values range very widely; for example, the adult size of direct brooders ranges from 85 to 49,480 mm3.

Introduction  41

For polychaetes, relevant information on dispersal is widely scattered. Thankfully Carson and Hentschel (2006) have summarized the relevant information from ~ 200 samples holding 501 polychaete species that inhabit 10–120 m depth of the Santa Catalina Island, off Los Angeles, California and other surrounding six island shelves with an area of 2,739 km2. This publication includes appended data, which have been subjected to more analyses in this account to draw additional information. Of these 501 species belonging to 226 genera and 49 families, the life history is known only for 354 species and for the remaining 152 species belonging to 78 genera and 24 families, it is not yet described, especially for Parasonidae (27 species), Sabellidae (18 species), Ampharetidae (14 species) and Lumbrinereidae (12 species). In 1991, when Wilson has made the first survey, the history has already been known for 307 species. After a passage of 16 years, the history is described at the rate of three polychaete species/y (see also Giangrande, 1997). Clearly, there is an urgent need for more research input into this area. Carson and Hentschel (2006) have divided the 354 polychaete species into (i) high-dispersal ‘open marine’ category comprising of species that are capable of dispersing and exchanging larvae with different populations over tens of kilometer areas, (ii) medium-dispersal category denoting species capable of dispersal on the scale of one kilometer and (iii) low-dispersal ‘closed’ category that includes species dispersing in the range of < 1 km, as their larvae are cued to settle close to their parents (Table 1.11). In this categorization, they have considered not only the established PLK, LEC and DIR but also other complicating life history factors like epitoky (E) (vertical distribution), Mobile Adults (MA) (horizontal distribution) and asexual reproduction (AS) (local distribution). From the present analysis, the following are inferred: 1. Not surprisingly, the diversification of reproductive modes have gone to 41 recognizable types, 2a. Despite the incidence of epitoky (benthic spawning + PLK), mobile adults and/or asexual reproduction, the PLK lasting from > 10 days to months ensures high dispersal of polychaetes that freely spawn eggs/embryos in encapsulated jelly mass. 2b. Levin (1984) has also reported that Polydora ligni (7–42 days larval duration) and Pseudopolydora paucibranchiata (> 7 days larval duration) have dispersal capability over longer distance. Streblospio benedicti and Capitella with < 3 days larval duration have the ability for small scale dispersal only. However, LEC followed by pelagic larval duration of < 7 days ensures only medium level of dispersal. 2c. Benthic spawning, even with MA also limits the dispersal to medium level. 3a. Brooding + pelagic PLK for more than 8 days and larvae released at three chaetigers stage ensures medium dispersal. 3b. However, brooding resulting in direct development with zero or 1 day pelagic larval duration limits the dispersal to low level. 4. Despite the presence of neochaete larva, MA, MA + AS, DIR results in low dispersal. 5. Of 354 species, 28.5% (109 species) polychaete species are broadcast spawners followed by

42  Reproduction and Development in Annelida

PLK larvae with potential for high dispersal. The remaining (18.1%) free spawners (LEC 56 + bs 8) are characterized by medium dispersal, as their LEC larval duration lasts for 0–3 days only. Of 82 brooder species, 33 of them pass through a period of PLK larval duration lasting for ~ 8 days. They are also characterized by medium dispersal. Brooding with no pelagic larval duration (49 species) and DIR (99 species) limit the dispersal to low level. Briefly, 49% of polychaetes are free spawners and are characterized by high or medium dispersal potential; another 9.3% (33 species) brood their eggs/ embryos/larvae but release their larvae for (medium level) dispersal. Of the remaining 42%, some are brooders with no pelagic larval duration (14%) and others (28%) are characterized by DIR with low dispersal capacity. From Wilson’s (1991) survey, it is shown that 47 and 53% of polychaete species are brooders and broadcast spawners, respectively. The survey by Carson and Henstchel indicates that only 42% are brooders. Notably, the annelid larvae with high potency for dispersal are capable of dispersing on the scale of tens of kilometers only.

1.10  Fertilization Site and Success The broadcast spawning polychaetes reproduce by releasing gametes into water, where fertilization and subsequent development occur. In them, the ambient sperm concentration, into which spawned eggs are released, is a key factor in deciding Fertilization Success (FS). Motile polychaetes may form mating aggregation and often display a high degree of synchronized spawning, as in palolo worm Eunice viridis (Caspers, 1984). In eggs of gregarious species, too many sperm may lead to polyspermy. Hence, laboratory studies are required on gamete longevity and optimal egg-sperm ratio that ensures the highest FS. The experimental study on a representative gregarious serpulid Galeolaria caespitosa provides some basic information on FS. From a well designed laboratory study, Kupriyanova (2006) has found that 1. despite variations between 58 and 66 µm, the egg size remains equal at 61.3 µm in worms weighing 10 to 50 mg (Fig. 1.12A). 2. Likewise, swimming velocity of sperm averages to 107.3 µm/second (s) in males weighing up to 25 g (Fig. 1.12B). 3a. Expectedly, FS increases sigmoidally with increasing sperm concentration up to 108/ml and 3b. Achieves the highest FS of 75% and > 80% at 108/ml sperm concentration, when given 1 and 10 minutes duration for fertilization, respectively (Fig. 1.12C). 4. Sperm velocity progressively decreases from ~ 145 µm/s and perceptibly from > 100 µm/s 2 hours after activation and finally too < 5 µm/s 8 hours after activation (Fig. 1.12D). More or less similar decreasing trends are noted for the decrease in FS (Fig. 1.12E, F). Surprisingly, the longevity of eggs and sperms is almost

Introduction  43

Figure 1.12 Experimental fertilization in the serpulid Galeolaria caespitosa. A. Effect of body weight on egg size, B. Effect of body weight on sperm velocity, C. Effect of sperm density on fertilization success, D. Sperm swimming velocity after activation, E. Fertilization success after sperm activation and F. Fertilization success after egg activation (compiled and modified from Kupriyanova, 2006).

equal. In general eggs remain fertilizable for longer durations than the sperms. Williams and Bentley (2002) have undertaken a comparative study on the gametic longevity of an errant Nereis virens and sedentary Arenicola marina. In them, fertilizable egg age lasts for 72 days to ensure 100% FS. However, the age of sperm is just 30 hours and 60 hours to ensure 100 and 60% FS in sea water, respectively. In the siboglinids inhabiting thermal vents, cleavage commences 24 hours after fertilization. On in situ incubation of eggs, cleaved embryos average to 65% (range: 40–84%) in the laboratory and 90% (74– 98%) in Riftia pachyptila and Lamellibrachia luymesi, respectively. But, they average to 86% (76–91%) in the field for R. pachyptila (Hilario et al., 2005). Understandably, FS in these in situ and ex situ incubated eggs ranges between 40 and 98%. Apart from these gonochorics, experimental FS values are also available for hermaphroditic annelids, in which 100% FS is expected. As already indicated, the removal of spermatophore in the earthworm Eisenia foetida increases FS in 11% of more eggs within a cocoon (see Table 1.9). In selfing sabellid polychaete Laonome albicingillum, FS is 83%, in comparison to 88% in outcrossing individuals (Hsieh, 1997). In another polychaete Ophryotrocha diadema, grouping of dozen individuals doubles the percentage of non-egg layers (20%) against just 10% of non-egg layers in isolated individuals (Henshaw et al., 2015). Evidently, FS remains at 80 and 90% in grouped and isolated individuals, respectively. From field observation, only very few FS values are reported for polychaetes. In epitokers, aggregation of spawners synchronizes spawning and chemical

44  Reproduction and Development in Annelida

cue-guided swimming sperm may lead to > 90% FS. Behavioral strategies also bring the mating partners close together and ensure high FS in ambient waters. The ‘nuptial dance’, during which gametes are released, may bring the mating partner close together to ensure high FS, as in Autolytus prolifer. In others, pseudocopulation, during which the gonopores of the mating partners are brought together, may bring the partners still closer to achieve higher FS. However, no information is yet available on this aspect. For benthic spawners, FS is low (40–60%). A series of publications (Hardege et al., 1996, Watson et al., 1998 and Williams and Bentley, 2002) have reported that the lugworm A. marina female lays eggs within its burrow, where the laid eggs remain fertilizable for 72 hours. Male lugworm releases sperm bundles on the sediment surface. Carried by incoming tides over the sediment surface and guided by volatile organic pheromone, the sperms are drawn into the burrow at the density of 106/ml in the water column. Fertilization is successful but varies widely from 0 to 90% with most values falling within 40–60%. In 45% of polychaetes (165/354 species, see Table 1.11), fertilization is external. It is also external in another 9% brooding polychaetes, in which larvae are released after brooding embryonic and/or a short or longer larval stages. In these 9% brooders, the sperms are either delivered on the female’s body or the female draws the sperms using chemical cues, as in Arenicola marina or filtering the ambient water by the branchiae and other structures for food acquisition. For example, sperms are gathered in the short oral tentacles of the female before mature eggs are passed over the tentacles for fertilization in Nicolea zostericola, which broods its embryo in an extratubular cocoon (Eckelbarger, 1984). This is also the case in the intratubular brooder Neolepoa septochaeta (see McEuen et al., 1983). In all, fertilization is external in 54% of polychaetes. In other brooders characterized by direct development/viviparity, fertilization seems to be internal. Polychaetes engage an array of organs that can be termed as ‘penis’. The penis is inserted into the female’s genital pores (e.g. Saccocirrus eroticus) or female’s body (e.g. Stratiodrilus novaehollandiae). In females with no sperm receptacle, the sperms are transferred by hypodermic injection. Sella and Ramella (1999) list five dinophylids, which engage hypothermic injection for sperm transfer. Fertilization is also internal and occurs in the ovisac or cocoon. In oligochaetes pseudocopulation is not uncommon, during which mutual transfer of sperm/spermatophore occurs. In Eisenia foetida, for example, the spermatophores are implanted between the 21st and 24th segments, adjacent to the spermathecal anlage. In 86% of the mating partners, only one spermatophore is implanted (Monroy et al., 2003). The preferred segments for spermatophore implantation in the leech Haementeria parasitica are also close to those segments, in which eggs are borne and thereby the distance to be travelled by the sperms to reach the eggs is minimized (see Sawyer et al., 1981). In most glossiphoniid, piscicolid and erpobdellid leeches, which lack penis, sperm are transferred by a hypodermic injection. In other earthworms

Introduction  45

like Lumbricus terrestris, the copulatory setae pierce into partner’s skin and inject sperm as well as an allohormone that inhibit sperm digestion (Koene et al., 2005). At concentrations of 7 × 104/ml and 1.5 × 106/ml, sperm from more than one male G. caespitosa increases FS from ~ 25–30% to ~ 90–95% (Marshall and Evans, 2005). Whereas information on FS for gonochores (e.g. Arenicola marina) and protandric hermaphrodites (e.g. G. caespitosa) is limited, available information on FS of simultaneous hermaphroditic annelids is also limited. Available publications elucidate the adjusted resource allocation in hermaphrodites (e.g. polychaetes: Ophrytrocha diadema, Schleicherova et al., 2006, Cannarsa et al., 2015). For example, on enforced isolation, the number of cocoons ♂ generated is 12/♂ + and 2/ + in the oligochaetes Eisenia andrei and E. foetida, respectively, clearly indicating that the latter is able to reduce allocation with enforced isolation and self-fertilization. However, hatching is limited to 2.5 cocoons per female in E. andrei and 1.5 cocoons per female in E. foetida. Interestingly, Hsieh (1997) is perhaps the only author, who has reported FS (in unit of egg production) as a function of number of eggs used by self- and cross-fertilization in simultaneous sabellid polychaete Laonome albicingillum. From Fig. 1.13, the following may be noted: 1. With increasing number of eggs made available/used for fertilization, FS increases in both self- and cross-fertilization. 2. But outcrossing worms are able to achieve 50 to 95% FS, while selfers achieve 50 to 75% FS only. Competition for mating in E. andrei also increases FS. For example, FS is increased from ~ 60% with a single

Figure 1.13 Fertilization success in self- and outcross-fertilization in the hermaphroditic sabellid polychaete Laonome albicingillum (drawn from data reported by Hsieh, 1997).

46  Reproduction and Development in Annelida

mating partner but > 80%, when the number of mating partners is increased to two–six (Porto et al., 2012).

1.11  Annelidan Larvae As indicated, a majority of polychaetes display indirect development involving typically the trochophore larva, or one or other modified form of trochophore. Many genes, whose expression is known to regulate one or other events of embryonic and larval development, have been identified; for more details, Irvine and Seaver (2006) may be consulted. In his narrative account, Rouse (2006) has listed 10 annelid larval forms, and described a range of ciliary bands and tufts that are used for feeding and locomotion. Accordingly, the ciliary bands are designated as (i) Prototroch with a single equatorial compounded ciliary band dividing the larva into an anterior episphere and posterior hyposphere, (ii) Akrotroch with complete ciliary ring around the episphere, as in eunicids and other six families, (iii) Meniscotroch with a crescent shaped area of stout cilia on the episphere, as in glycerids and 14 other families, (iv) Metatroch characterized by the post-oral ciliary band, beating from posterior to anterior direction, e.g. Amphinomidae and 10 other families, (v) Neurotroch with a longitudinal ciliary band running from behind the mouth to near the anus, as in siboglinids and 30 other families, (vi) Mesotroch, in which the transverse ciliary ring adorns the larval midbody, as in Chaetopterus, (vii) ventral Gastrotrochal and dorsal Nototrochal ciliary rings on segment, as in spioniform larvae and (viii) Telotroch with locomotory ciliary ring located at the posterior end of larva, as in hesionids and 11 other families. Table 1.12 briefly summarizes the morphology and occurrence of 10 different trochophore and trochophore-like larvae reported from polychaetes. During development, some spionids, draw nutrients from the so called nurse eggs and larvae within a brood. Nurse eggs serve as source of extra embryonic nutrients for developing embryos. On activation, these eggs too release polar bodies but subsequently are compartmentalized with a loss of nuclear DNA (e.g. Boccardia proboscidae, Gibson et al., 1999, Smith and Gibson, 1999). As a climax, larvae (e.g. Streblospio benedicti, see Blake and Arnofsky, 1999) ingest fellow larvae prior to hatching and release from brood. As much as 95% of the eggs brooded by Polydora cornuta are nurse eggs and are ingested by adelphophagic larvae hatched at 3rd chaetiger stage (Mackay and Gibson, 1999). Some P. cornuta females switch between adelphophagic and planktotrophic larvae arising from a single brood or in successive broods. Both nurse eggs and adelphophagic larvae are an adaptive mechanism to accelerate early cleavages in smaller eggs with less yolk that

Introduction  47

Table 1.12 Description of polychaete larvae (condensed from Rouse, 2006, all figures are sketches from Rouse, 2006 and others) Larva

Description

Proto-trochopore

Completely or nearly completely ciliated trochophore, e.g. oweniids

Trochophore

Typical larva with opposed-band method of feeding by involving ciliary bands of prototroch and metatroch

Meta-trochophore

Normal trochophore with clear signs of segmentation. Non-functional parapodia, if present, e.g. Lanice conchilega

Nectochaeta

Trochophore with functional parapodia, e.g. phyllodocids

Aulophore

Metatrochophore is long living in a tube, e.g. terebellids, pectinariids

Chaetosphaera

Swims by undulation as well as cilia. Can roll up into a ball, e.g. spioniformids, sabellarids

Nectosoma

More like chaetosphaera larva but cannot roll up into a ball, e.g. Poecilochaetus

Rostraria

A pair of distinctive tentacle is used for feeding via ciliary bands, e.g. amphinomid

Mitraria

Normal trochophore but undergoes ‘catastrophic metamorphosis’ by casting off much of its body prior to settling as juvenile, e.g. Oweniidae, Myricola

Erpochaete

More a ‘juvenile larva’ that creeps over the sediment using its chaetae

Figure

facilitates the embryo to grow faster (see Pandian, 2016). Interestingly, Schneider et al. (1992) have shown that with ~ 90% yolk content, egg size of Platynereis massiliensis is 10 times that (with 64% yolk content) of the sibling species P. dumerilii. As a consequence, the cell cycles upto 5th cleavage are 3.7 times slower in P. massiliensis than in P. dumerilii. Hence, it is advantageous

48  Reproduction and Development in Annelida

to have smaller eggs, whose embryos/larvae draw extra-embryonic nutrients from nurse eggs and larvae.

1.12  Defense and Parental Care The soft-bodied, slow motile/sedentary annelids are not readily prone to predation, as it has been considered earlier. Many annelids provide protection to their eggs/embryos by (i) encapsulating them in jelly matrix or gelatinous envelope, (ii) synthesis and accumulation of deterrents in their larvae and adults, (iii) providing internal or external structures for brooding eggs/ embryos/larvae as well as provision of hard cocoon or soft cocoon followed by parental care. Jelly matrix: In general, polychaetes are broadcast spawners. However, some of them shed pear-shaped or spherical egg masses, in which eggs are embedded in a jelly matrix and attach them to suitable substrate or the parent’s tube. In echinoderm eggs, the jelly facilitates floatation of pelagic eggs and increases the chances for sperm-egg collision and hence fertilization success (see Pandian, 2018). In polychaetes, the jelly matrix is considered to protect the developing embryos, supply nutrients but limit dispersal of the offspring to settle in and around the ‘suitable’ habitat (Sato et al., 1982). From an experimental study, Sato and Osani (1996) have found that (i) de-jellied unfertilized eggs of the polychaete Lumbrinereis latreilli lose the capacity for sperm binding and hence become unfertilizable, (ii) fertilizability can be restored by addition of jelly to the de-jellied eggs. An electron microscopic study has revealed that the sperm-egg binding occurs only in the presence of jelly and an unknown interaction between the jelly matrix and spermatozoa may be a pre-requisite to induce acrosome reaction. Incidentally, there are others, in which the jelly layer is formed shortly after fertilization (e.g. Nereis falcaria, Read, 1974). In Platynereis dumerilii, a huge jelly coat is formed immediately after fertilization (Kluge et al., 1995). Chemical defense: The recent past has witnessed increasing number of publications on chemical defense of polychaete worms. For example, the LEC spionid Streblospio benedicti contains at least 11 chlorinated and brominated hydrocarbons (alkyl halides) and the brooding capitellid Capitella sp I contains three brominated aromatic compounds. Quantifying these alkyl halides, Cowart et al. (2000) have found that S. benedicti contains 1.8, 8.3, 4.7 and 28.9 ng/mm3 in the pelagic LEC larva, post-release, new recruit and adult, respectively. This increasing trend suggests that the halides are synthesized during post-embryonic developmental stages. With contrasting life history of Capitella sp I, the halogenate compound contains 1150 and

Introduction  49

126 ng/mm3 in the LEC larva and adult, respectively. At these concentrations, the haloaromatics are known to deter predation. Investigating the chemical and structural defense by external strategies in six tubiculous sabellids, Giangrande et al. (2014a) have reported toughness of the branchial crown, structure and strength of the tube as well as antibacterial lysozyme activity in the mucus. The tube strength, as measured by tearing weight, decreased from 800 g in hard substratum-inhabiting Sabella spallanzanii to 200 g in S. spectabilis. However, the anti-bacterial lysozyme activity in the mucus of the soft substratum burrowing S. spectabilis remains high. In another interesting study, Iori et al. (2014) have observed that on being attacked by the fish Oryzias melanostigma, the oenonid Halla parthenopeia, after emitting purple mucus, quickly moves away (similar to inking and clouding by sea hares and cephalopods, Pandian, 2017) and subsequently release transparent mucus. The purple mucus is toxic, due to the presence of halochrome a 1, 2 anthraquinone. Brooding: In aquatic animals, affording protection of eggs, embryos and/ or larvae is not uncommon. The afforded protection ranges from simple embryo and/or larval guarding to brooding them until a certain larval stage or complete development. Brooding may occur in burrows (e.g. Aphelochaeta, Petersen, 1999). In Ophrytrocha puerilis puerilis, female spends more time in brooding than male (Berglund, 1986). Nereis acuminata reproduces monogamously and exhibits male parental care, a rare reproductive mode in marine invertebrates (Weinberg et al., 1990). In others like Neanthes arenaceodentata and Platynereis massiliensis, females lay eggs and die, males fertilize the eggs, and protect and ventilate the fertilized eggs (see Premoli and Sella, 1995). Brooding occurs within a flexible cocoon in O. adherens; O. socialis produces a system of branching mucous tubes, in which eggs are laid. In O. hartmanni, mucous tubes are ventilated by parents (Paavo et al., 2000). However, quantification of guarders and brooders has not been made. There are hints for other taxonomic groups. For example, brooding occurs in 3 and 27% of ophiuroids and gastropods, respectively. Notably, brooding occurs in one or more species in all the listed phyla (Table 1.13) and within each phylum in all classes (e.g. echinoderms). About 42–47% of polychaetes are brooders, in which ~ 36% brooding lasts until the directly developed young ones are released. Brooding ranges from ~ 1% in echinoderms to 96% in crustaceans. However, it is terminated with a release of young ones in ~ 9% of annelids but ~ 19% of crustaceans. It seems that aquatic invertebrates can ill-afford viviparity; for example, true viviparity occurs in a single ophiuroid Amphipholis squamata in echinoderms and six species of annelids. On the other hand, it occurs in 577 species of teleostean fishes (Pandian, 2013). Cocoon: A distinguishing feature of clitellate annelids is the presence of specialized segments comprising the clitellum (Sayers et al., 2009). The clitellum secretes proteinaceous egg case, the cocoon, into which eggs

50  Reproduction and Development in Annelida

Table 1.13 Brooding and direct development in some aquatic vertebrates and invertebrates (from Pandian, 2011, 2013, 2016, 2017, 2018) Taxon

Species (no.)

Brooded (%)

Teleostean fishes

32,000

22

Direct Development (%) 1.8

Crustacea

52,000

96

~ 10 +

Mollusca

~ 1,00,000

~ 34 +

~ 25 +

Annelida

17,000

~ 42

Echinodermata

7,000

~ 1  +

~ 28 + 1 species only

are deposited. In clitellates, the cocoon provides a microenvironment for embryonic development and prevents desiccation/imbibition, predation (however, see Young, 1988) and microbial invasion. It is highly resistant to physical and chemical decay and renders the preservation of fossilized spermatozoa within the 50-million year old cocoon (Bomfleur et al., 2017). In the glossiphoniid leech Theromyzon tessulatum, the cocoon formation is initiated with secretion of a thin, external mucous layer, into which fibrous proteinaceous matrix is deposited, forming a sheath surrounding the clitellum (see also Westheide and Muller, 1996). After the eggs are shed through the female pore, the sheath and its contents are passed over the worm’s head and sealed at either end with glue-like plugs, called opercula. In leeches internal fertilization occurs either in the ovisac or cocoon (see Sayers et al., 2009). Clitellates secrete three types of cocoons: (i) mechanically strong, (ii) hard-shelled cocoons that are abandoned, leaving the embryos to develop independently on nutritive cocoon fluid (albuminotrophy), thermally and chemically resilient (iii) membranous and (iv) gelatinous cocoons. The large yolky (oviparous) eggs, deposited in the membranous cocoons, are brooded by the parent (Rossi et al., 2016). In the aquatic leech T. tessulatum, cocoon formation is a simple but dynamic series of coordinated events. Just a week prior to egg laying, cell type I proliferates and differentiates into cell types II and III, depending on the position in cell type I within the clitellum. Type II cells secrete alcian blue-staining granules that form strong, malleable and bioadhesive opercula (see also Rossi et al., 2013). Type III cells secrete azocarmine-staining granules that build the cocoon wall. Type I and V cells make minor contributions and type IV plays supporting/signaling role (Sayers et al., 2009). The jawless leeches (Erpobdellidae) produce relatively more (~ 1,000) but smaller (~ 50 µm) eggs, enclosed in a cocoon, which is cemented on a substratum and left totally uncared (Table 1.14). In others like glossophoniids, the cocoons contain < 60 large (~ 600 µm) eggs, supplied with albumin and protected. Besides supplying albumin, Helobdella stagnalis protects and feeds the young one. With the evolution of parental care in the glossophoniids leeches, mortality of eggs, hatchlings and juveniles have been almost totally reduced to zero.

Introduction  51

Table 1.14 Increasing level of parental care from erpobdellid to glossophoniid leeches (from Kutschera and Wirtz, 2001, modified) Features

Erpobdella octoculata

Glossiphonia complanata

Helodella stagnalis

Cocoon (no./season)

~ 120

~ 3–4

~ 5–6

Fecundity (no./season)

~ 1,000

~ 60

~ 50 500

Egg size (µm)

50

600

Albumin

No

Yes

Yes

~ 10 min

30 d

50 d

High

No

Ventilation Egg mortality Care for hatchling Hatchling mortality Care for juvenile Juvenile mortality

No

Yes

High

Low

No

Yes

High

Low

~ 0 Yes ~ 0 Yes + Fed ~ 0

1.13  Metamorphosis and Settlement It must first be recognized that although metamorphosis and settlement are closely related, they are different temporally separate processes. The former is defined as the process, by which a larva undergoes a series of changes to terminate the larval phase. But the settlement is the process, by which a planktonic larva explores and selects a suitable substratum, toward which it moves to finally settle (see Qian, 1999). Larval settlement on large spatial scale is primarily determined by hydrodynamics; however, the successful settlement by competent larvae (starved and aged larvae may not be competent) on smaller spatial scale is mediated by abiotic and biotic cues. These cues may originate from host plants/animals, bacterial microfilms or habitats. The ease with which the microscopic settling larvae can visually be recognized by the formation of milky white calcareous tube in serpulids has facilitated many publications. The segmented larvae of some terebellids and many polynoids undergo metamorphosis as plankton and remain in the pelagic realm for days and weeks, and eventually settle and start benthic life (see Qian and Uwe-Dahms, 2006). In some spionids, sabellarids and oweniids, the metamorphosing larvae possess enlarged erectile anterior setae that serve as floating device (Bhaud and Cazaux, 1990). Settlement of pelagic larvae is of both academic and economic importance. As foulers, the settling larvae create problems in harbors, ships and on coolant screens in power stations. Sabellarids and serpulids often settle gregariously and form colonies. In this regard, Phragmatopoma californica, Sabella alveolata, Spirobranchus polycerus, Hydroides dianthus and H. ezoensis have received much attention (Qian and Uwe-Dahms, 2006).

52  Reproduction and Development in Annelida

Table 1.15 Factors that induce or inhibits settlement of polychaete larvae Species

Factor

Reference

Polydora ligni

Starvation reduces settling ability

Qian and Chia (1993)

Hydroides elegans

Aged larva loses settling ability

Qian and Pechenik (1998)

Spiorbis borealis

Dark substratum attracts settlement James and Underwood (1994)

Capitella sp

Organic rich sediments (but not hydrogen sulfide, Cuomo, 1985) attracts settlement

Dubilier (1988)

Capitella sp

Juvenile hormone arising from sediment attracts settlement

Biggers and Laufer (1992)

S. borealis

Fucus serratus attracts settlement

Williams (1964)

S. rupestris

Lithothammoni attracts settlement

O’Connor and Lamont (1978)

Spirobranchus giganteus

Diploria strigosa attracts settlement

Marsden et al. (1990)

H. elegans

Bugula neritina attracts settlement

Bryan et al. (1998)

H. elegans

Levels of glycojuvenate secreted by rod bacteria attract settlement

Lau and Qian (1997)

Janua brasiliensis (experimental)

Extracellular polysaccharides and glycoprotein attract settlement

Kirchman et al. (1982)

Phragmatopoma californica (experimental)

Fattly acids: cis eicosapentaenoic acid, palmitic acid, palmitoletic acid attract settlement

Pawlik (1986)

H. elegans (experimental)

Amino acids: glycine, glutamine, aspartic acid arising from leechate of B. neritina attract settlement

Harder and Qian (1999)

Available information on settlement is summarized in Table 1.15. Earlier, sea grass and coral species that attract the serpulid settlement had been identified. Bryan et al. (1998) have recognized that alcohol extract of dried aqueous Bugula neritina leechate carries the compound responsible for attraction of H. elegans larvae to settle. Subsequently, experimental investigations have shown that one or other polysaccharides, fatty acids and amino acids can also attract the settlement. Recent studies have shown that the microbial film (Beckmann et al., 1999), and chemical signals like the polar aliphatic amino acids emanating from the biofilm (Hadfield et al., 2014) are not individually responsible for final settlement but the sorbent-like substratum acting as a co-factor is also responsible to induce final settlement of H. elegans larvae (Harder et al., 2002). In view of the fouling problems on the ships, harbors, pipelines and screens in power stations, this aspect merits more attention.

2 Sexual Reproduction

Introduction Annelids display divergent expressions of sex. In them, sexuality ranges from parthenogenesis to gonochorism and self-fertilizing to sequential and serial hermaphroditism. Sexual reproduction in gonochoric and hermaphroditic annelids is characterized by gametogenesis, meiosis and fertilization of gametes arising from one (self-fertilizing) or more than one (multiple matings, polyandry) individuals. In parthenogenic polychaetes, females are always present and males may appear sporadically. In parthenogenic earthworms, male reproductive system is eliminated either partially or totally. Barring hirudineans, many polychaetes and oligochaetes undergo agametic reproduction. Sexual reproduction may succeed or co-exist with agametic cloning. With occurrence of parthenogens, hermaphrodites and agametic cloners, sexual reproduction in annelids is a fascinating subject for research.

2.1  Reproductive Systems Polychaetes: In majority of polychaetes, the gonads are dispersed and not discrete organs. Masses of eggs and sperm are generated as projections or swellings from undifferentiated peritoneal lining of the coelom. Most segments generate gametes, as in errant nereidids and eunicids. With specialization of the thorax and abdomen in sedentary polychaetes, the gamete generation is, however, limited to the abdominal segments. Increasingly, the number of gametogenic segments is limited (Amphitrite) to only six segments in Arenicola and further to only two ovarian segments in Spirorbis spirorbis (Daly, 1978a). Table 2.1 lists some examples for the anlage of gametogenic segments in gonochoric, protandric and simultaneous hermaphroditic

54  Reproduction and Development in Annelida

polychaetes. In the intra-ovarian polychaetes, oogenesis is completed in the projections/swellings of the coelomic lining. But gametogonia are shed into the coelom, where further genesis and maturation take place in extraovarian polychaetes (Eckelbarger, 1983, 1986). In gametogenic segments, the nephridia serve as exits for the genital and excretory products. However, the ruptured body serves as an exit in epitokous polychaetes (Barnes, 1974). Oligochaetes: Strikingly, the number of gametogenic segments is very much limited to reduce the reproductive cost in the simultaneous hermaphroditic oligochaetes. In some of them, the nephridia serve as exits for the sperms. Table 2.1 Examples for the presence of ovary and testis in the same or separate gametogenic segments in some hermaphroditic polychaetes. SH = simultaneous hermaphrodites Species, Reference

Features Gonochorics

Nereidids, Eunicids Arenicola Spirorbis spirorbis, Daly (1978a)

Almost all segments are gametogenic Only 6 gametogenic segments Only 2 ovarian segments Protandrics

Ophrytrocha diadema Schleicherova et al. (2010)

Until the age of 21st day, testes are present in the 4th and 5th segment. Becomes SH, when 14–17 segments are added. Functions as SH from 30th–40th d

O. gracilis Sella et al. (1997)

Four testicular segments from 3rd to 6th. Followed by 14 ovarian segments from 15/16 to 30th. External fertilization. Generates 11.2 cocoons/w for 13 weeks SH with gonads in different segments

Aracia sinaloae Tovar-Hernandez and Dean (2014)

First five abdominal ovarian segments followed by last eight testicular segments

Laonome albicingillum Hsieh (1997)

Anterior most 10 abdominal ovarian segments and subsequent the 33 abdominal testicular segments. Extraovarian maturation. No seminal receptacle

Neanthes limnicola Fong and Pearse (1992)

Viviparous. Oocytes collected from posterior one third of the body SH with gonads in the same segment

Branchiomma bairdi Tovar-Hernandez et al. (2009)

Gametogeneic segments are present from 7th thoracic to entire abdomen. Ovary and testis co-exist in the same segment but separated topographically

Diopatra marocensis Arias et al. (2013)

Oogenesis and spermatogenesis occur in temporally and topographically separated regions of the same segment. Spermatogenesis occurs close to the ventro-lateral branches of the blood vessel from February to April. Oocytes mature in the coelom during March–June. Self-fertilization. SH at 2 no./m2, protandric at 4 no./m2 and gonochoric at > 10 no./m2. Female ratio ranges from 0.67 to 0.80

Sexual Reproduction  55

In other oligochaetes, the gametogenic segments are located anteriorly (Fig. 2.1A). The female gametogenic segments are located posterior to the male segments. Notably, the paired gonads are developed as discrete organs with the respective ducts so that a discrete reproductive system is present. The female components include a pair of ovaries, ovisacs and oviducts as well as female pores in the 13th and 14th segments. The male component

Figure 2.1 Reproductive system in A. an oligochaete earthworm (freehand drawing), B. hirudinean Hirudo medicinalis (modified and redrawn from Mann), C. Glossiphonia complanata (modified and redrawn from Harding and Moore) and D1. naidid, D2. tubificid, D3. enchytraeid, D4. potomadrillid and D5. aeolosomatid. Segments number are indicated by Arabic numerals, T = testis, VS = vas deferens, S = spermatheca, O = ovary, G = gonad. Note the changes in distribution of reproductive components in different segments (drawn from Lassarre, 1971).

56  Reproduction and Development in Annelida

comprises testes, male funnels in the 10th and 11th segments and variable number of seminal vesicles in 9–12 segments as well as different ducts and male pores (see Diaz-Cosin et al., 2011). The number of gametogenic segments along the body varies in different families (Barnes, 1974). Figure 2.1D1–D5 shows the locations of ovary, spermatheca and vas deferens in naidid, tubificid, enchytraeid, potamodrillid and aeolosomatid. In the exceptional aeolosomatids, sexual reproduction is a very rare phenomenon. Genital organs are known only in some species. Paired gonads differentiate at the ventral coelomic epithelium with ovaries in the mid-body and testis in front of them. Usually, only one ovary is fully mature. Gonoducts are lacking. Sperms are discharged through metanephridia and eggs are shed through a simple porous body wall. Copulation has never been observed. Spawning and normal sexual development are reported only from Aeolosoma quaternarum (Bunke, 1986). For more details, Jamieson and Ferraguti (2006) may be consulted. Hirudineans are also hermaphrodites, in which the number of ovary is limited to a single pair, as in Hirudo medicinalis (Fig. 2.1B) but the pair is extended to a few segments, as in Glossiphonia complanata (Fig. 2.1C). However, the number of testicular segments is 10. Notably the anlage of the ovaries is anterior to those of testes. The oocytes leave the ovaries to ovisac, where they mature (Barnes, 1974). Regrettably, not many authors have described the exact locations of the ovarian and testicular segments in polychaetes. Available information including that of a passing remark for Neanthes limnicola is listed in Table 2.1. In Ophryotrocha spp, two–four anterior testicular segments appear first; subsequently, posterior segments are added, of which 4–16 segments harbor ovaries. In them, the anlage for the testicular and ovarian segments is clearly separated. This is also true of some SH like Aracia sinaloae and Laonome albicingillum. Notably, the ovarian segments appear first in the anterior abdominal segments. In other SH, both ovaries and testes co-exist in the same segment, although topographically separated, as in Branchiomma bairdi and temporally also in Diopatra marocensis. Strikingly, not all SH commence as SH. Manifestation of hermaphroditism is dependent on population density. For example, D. marocensis functions as SH, protandric and gonochoric at densities of 2, ~ 4 and > 10 no./m2, respectively (see Arias et al., 2013). Differences in structural organization of the reproductive system among annelids have the following implications: 1. The Primordial Germ Cells (PGCs) are responsible for the manifestation of sex. Derived from PGCs, the Oogonial Stem Cells (OSCs) manifest female sex in some individuals and that of Spermatogonial Stem Cells (SSCs) male sex in others. Understandably, mutations involving one or more genes have retained both OSCs and SSCs in hermaphroditic annelids. 2. In polychaetes, the transition from crawling errant mode of life to sedentary/sessile mode has involved reduction in

Sexual Reproduction  57

the number of gametogenic segments to facilitate the sharing of resource allocation between gametogenesis and brooding/parental care. 3. In oligochaetes and hirudineans, the cost of manifesting and maintaining dual reproductive systems is facilitated by reduction in the number of the ‘costlier’ ovary to a single pair, in comparison to that of gonochoric polychaetes, in which the minimum is six pairs, as in Arenicola. Internal fertilization and protection of eggs within the cocoon increase survival and recruitment of progenies. To escape from ‘inbreeding depression’ and to increase genetic diversity, every effort is made by hermaphroditic annelids to ensure cross fertilization (see later). On enforced isolation, some of them reduce allocation for cocoon production (e.g. Eisenia foetida).

2.2 Gonochorism Sex is a luxury, and costs time and resources but ensures recombination and genetic diversity, the raw material for evolution and speciation. Not surprisingly, the polychaetes are more diverse and speciose than oligochaetes and hirudineans. Both male-(XX-XO) and female-(ZZ-ZW) heterogametic sex determination mechanisms are reported to operate in gonochoric polychaetes. Whereas the mechanisms are stable in most gonochoric polychaetes, labile mechanisms have been demonstrated in a few polychaetes. Besides, Premoli et al. (1996) have reported heritable variation in sex ratio of gonochoric Ophryotrocha labronica and advanced a hypothesis that sex is determined by multilocus (polygenic) genetic systems in polychaetes.

2.2.1  Sex Ratio The ratio represents the cumulative end products of sex determination and differentiation processes. It may serve as a key index to assess potential natality in a population. Incidentally, no female is yet known in Parenterodrilus taeniodes (Purscke, 2006). However, sex cannot easily be distinguished morphologically in gonochoric polychaetes. Not surprisingly, many authors have refrained from reporting the ratio. However, sex can be distinguished in 42–47% of brooding polychaetes. It can also be distinguished in species, in which the color of the testis differs distinctly from that of the ovary. For example, the abdomen of a ripe Pomatoceros male appears white and that of a female bright pink or orange colored due to the difference in color of the sperms and eggs (see Barnes, 1974). Encountering immature and sporadic occurrence of hermaphroditic individuals, some authors have assigned an additional ratio for them (Table 2.2). In a few polychaetes, sex ratio remains stable at or around 0.5 ♀ : 0.5 ♂, indicating the chromosomal mechanism

58  Reproduction and Development in Annelida

Table 2.2 Sex ratio of some gonochoric polychaetes Sex Ratio ♀ : ♂ : ♂ +

Species

Reference

Chromosomal mechanism of sex determination Amphisamytha galapagensis

0.50 : 0.50

McHugh and Tunicliffe (1994)

Hediste japonica

0.50 : 0.50

Sato (1999)

Sabella spallanzanii

0.50 : 0.50

Currie et al. (2000)

Marphysa sanguinea

0.50 : 0.50

Prevedelli and Simonini (2003)

Perinereis cultrifera

0.50 : 0.50

Prevedelli and Simonini (2003)

P. rullieri

0.50 : 0.50

Prevedelli and Simonini (2003)

Glycera dibranchiata

0.55 : 0.45

Creaser (1973)

Polygenic mechanism of sex determination Bispira brunnea

0.33 : 0.37 : 0.22*

Davila-Jimenez et al. (2017)

Branchipolynoe seepensis

0.19 : 0.28 : 0.29*

Jollivet et al. (2000)

Nicolea upsiana

0.19 : 0.46 : 0.35†

Garraffoni et al. (2014)

N. zostericola

0.45 : 0.38 : 0.13†

Eckelbarger (1975)

Nereis virens > 30 cm

0.50 : 0.50

< 30 cm

0.67 : 0.33

Capitella capitata ♀×♂

♂× ♂ +

Dinophilus gyrociliatus

Creaser and Clifford (1982) Petraitis (1985b)

0.50 : 0.50 0.03 : 0.97 0.75 : 0.25

D. gyrociliatus

Prevedelli and Simonini (2003) Prevedelli and Vandini (1999)

cereals

0.50 : 0.50

tetramin

0.67 : 0.33

Ficopomatus enigmaticus

Obenat and Pezzani (1994)

spring cohort

0.39 : 0.61

autumn cohort

0.26 : 0.74

Pomatoleios kraussi

0.67 : 0.33

Nishi (1996)

Polydora ligni

0.71 : 0.29

Zajac (1991)

Halla parthenopeia

0.28 : 0.72

Osman et al. (2010)

Lumbrinereis funchalensis

0.19 : 0.81

Osman et al. (2010)

* hermaphrodites, † immature, hence sex not known

of sex determination. In others like Capitella sp I, the homogametic ZZ males change sex to hermaphrodites and a cross between ZZ male and ZZ hermaphrodite skew the ratio in favor of male. In male heterogametic (XO) Dinophilus gyrociliatus, selective fertilization of larger eggs by sperms bearing X chromosome and small eggs by sperm carrying no sex chromosome nullifies the chromosomal mechanism of sex determination and sex ratio,

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as well. In all others, sex seems to be determined by polygenic system, i.e. one or more genes harbored on autosome(s) have an overriding effect in sex differentiation, following sex determination by sex chromosome. As a result, female ratio, for example, varies widely from as low as 0.19 in Lumbrinereis funchalensis to as high as 0.71 in Polydora ligni. Apparently, the autosomic genes in them express in response to one or other environmental factors like temperature in Ficopomatus enigmaticus and algal food Halopteris scoporia. In Typosyllis prolifera, homo/heterogametic sex chromosome determines irrevocably the primary sex of male, as its primary sex ratio is 0.5 ♀ : 0.5 ♂. However, female differentiation is labile, whereas differentiation in primary males remains absolutely stable throughout the life. Hence, some females switch to males depending on density. The number of females switching to males increases from 62% in Porec population to 77% in Pula population in Yugoslavia. Further, laboratory experiments have shown that the social condition is a more important factor in females switching to males in this ‘genetically unbalanced protogynous/serial hermaphrodite’. Control of sex ratio by environmental factors is explained in more details elsewhere (Chapter 7).

2.2.2  Ovary Somatic Index Gonado Somatic Index (GSI) relates to gonad weight to body weight. The nearest equivalent value is reported for the terebellid polychaete Eupolymnia nebulosa (Gremare, 1986). In view of difference in resource allocation, values for the Ovary Somatic Index (OSI) and Testis Somatic Index (TSI) are reported during recent years (e.g. fishes, Pandian, 2015). In E. nebulosa, the OSI value increases from 9 at age 2 to 21 at 4 years (Fig. 2.2A). However, TSI remains

Figure 2.2 A. ‘Gonado Somatic Index’ as function of age in Eupolymnia nebulosa (modified and redrawn from Gremare, 1986). B. Gamete weight as function of body weight in Mercierella enigmatica (modified and redrawn from Dixon, 1976).

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Table 2.3 Ovary Somatic Index (OSI) of some polychaetes (compiled from McHugh, 1993, Rouse and Fitzhugh, 1994). G = gonochoric, fs = free spawner, ds = discrete spawner, cs = continuous spawner, D = direct development, S = semelparous, Sqb = sequential brooder Species

Reproductive Mode

Fecundity

OSI

Eupolymnia nebulosa†

G

E. crescentis

G, fs, ds

–

15 ♀, 12 ♂

128, 500

Neoamphirite robusta

G, fs, ds

829, 833

6.7 8.6

Thelepus crispus

G, D, cs

51, 555

5.9

T. crispus

G, D, cs

51, 555

0.32*

410

2.4

Terebellids (McHugh, 1993)

Ramex californiensis

G, cs, Sqb Sabellids (Rouse and Fitzhugh, 1994)

Caobangia abbotti

Intra brooder

1

0.03

Fabricinuda trilobata

G, fs, Intra brooder

2

0.02

Manayunkia aestuarina

G, fs, riverine, Intra brooder

2

0.05

M. caspica

G, fs, riverine, Intra brooder

2

0.08

M. speciosa

G, fs, riverine, Intra brooder

4

0.10

Augeneriella alata

Intra brooder

2

0.11

Fabricia stellaris

Intra brooder

4

0.14

Maternal nutrients drawn

10

0.05

7

0.29

9

0.01

9

0.16

Intra brooder

10

0.14

Demonax medius

Intra brooder

1200

0.11

Potamilla torelli

Intra brooder

202

0.06

Potamilla sp

Intra brooder

Amphiglena marita A. terebro A. mediterranea A. nathae Amphicorina brevicollaris

Perkinsiana sp Perkinsiana antarctica

♂ , Intra brooder + ♂ , Intra brooder + ♂ , Intra brooder +

♂ , asynchronous extra brooder + ♂ , asynchronous extra brooder +

518

0.63

8666

0.07

300

0.01

† Gremare (1986), * data reported by Garraffoni et al. (2014)

constant at ~ 12 in the same age classes. Dixon (1975) has also estimated the dry body weight and gametes of the serpulid Mercierella enigmatica. Contrastingly, his calculated values indicate that (i) with increasing body weight, both TSI and OSI values increase, (ii) are in the range of > 67 at the body size of 1.5 mg (dry) body weight and (iii) the TSI values begin to exceed those of OSI beyond 1.5 mg size (Fig. 2.2B). The loss of ash-free dry weight of a spawning Nephtys caeca is 45 mg and amounts to OSI of 24.3; the values are 53 mg and 30.3 for N. hombergii (Olive et al., 1985). For the hermaphroditic

Sexual Reproduction  61

oligochaetes and hirudineans, the OSI values are not available. However, it may be difficult to estimate it in a leech, for example, which engorges by sucking blood and increases its body weight. But its body weight decreases due to progressive cocoon depositions and with advancing time/age. In the hermaphroditic polychaetes, ~ 80% of the gonad consists of the ovarian component (e.g. Ophryotrocha spp, see p 89). Direct measurements on the OSI have been made for four terebellid species by McHugh (1993). These values range from 2.4 for the continuous breeder and sequential brooder Ramex californiensis to 8.6 for the broadcast spawner Neoamphitrite robusta. Rouse and Fitzhugh (1994) and Garraffoni et al. (2014) have also reported data for the total egg volume and body volume of a few sabellids and terebellids, respectively. Hence, it is possible to calculate OSI on volume basis for these polychaetes (Table 2.3). These values are several orders lower than those reported from the direct estimates made on the weight basis by Dixon (1976) and Gremare (1986). As fecundity of some sabellid species is > 10 eggs, the calculated OSI values suggests that in such species characterized by the intra-ovarian oogenesis may have low OSI values. Notably, the OSI values for Thelepus crispus directly estimated and those calculated are 5.9 (McHugh, 1993) and 0.32 (Garraffoni et al., 2014), respectively. It is also difficult to comprehend that the semelparous terebellid Amaena occidentalis has an OSI of 0.036 (Garraffoni et al., 2014). Hence, it is recommended that OSI and TSI values are directly estimated on the basis of weight to enable the comparison of reproductive performance between annelid species and others, as well.

2.3 Hermaphroditism It is defined as the expression of both female and male functions in a single individual either simultaneously or sequentially. Three patterns of functional hermaphroditism have been recognized: (i) simultaneous hermaphrodites functioning as a male or female at a time (unilateral mating partners, e.g. Ophryotrocha gracilis, Sella et al., 1997) or as both male and female within a short span of time (reciprocal mating partners, e.g. O. gracilis). They may not usually undergo natural sex change, although O. gracilis can do it. The sequentials do it once in a life time in a single direction but the serials do it more than once in either direction (e.g. Syllis amica, Table 2.4). The sequentials are further divided into (a) male to female sex changing protandrics and (b) female to male sex changing protogynics. The former is more common among aquatic invertebrates (e.g. molluscs, Pandian, 2017, echinoderms, Pandian, 2018) but the latter among teleostean fishes (Pandian, 2011). Of

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Table 2.4 Hermaphroditic polychaetes (updated and compiled from many sources including those indicated by bold letters by Schroeder and Hermans, 1975) Simultaneous hermaphrodites Archiannelidae:  Mesonerilla armoricana, M. fagei, M. roscovita Arenicolidae:  Branchiomaldane vincenti Capitellidae:  Capitella hermaphrodita, Capitomastus minimus Cirratulidae:  Aphelochaeta sp, Caulleriella parva, Chaetozone vivipara, Ctenodrilus serratus Ctenodrilidae:  Raphidrilus nemasoma, Trilobodrilus Hesionidae:  Hesione sicula, H. mazima(?), H. pantherina, Microphthalmus fragilis, M. listensis, M. oberrans, M. sczelkowii, M. similis, M. tyrrhenicus, M. urofimbriatus Nereididae:  Neanthes lighti, N. limnicola (viviparous) (Fong and Pearse, 1992), Namalycastis indica, Namanereis quadraticeps, Platynereis massiliensis Onuphidae:  Diopatra marocensis (Arias et al., 2013) Orbiniidae:  Nainereis laevigata Pectinariidae:  Lagis koreni Polygordiidae:  Polygordius triestinus Polynoidae:  Macellicephala violacea Sabellidae: Amphiglena marita, A. mediterranea, A. nathae, A. terebro, Aracia sinaloae, Branchiomma luctuosum, B. bairdi, B. cingulata, Caobangia billeti, Demonax pallidus, Euratella salmacidis, Laonome albicingillum, Manayunkia aestuarina, Ophryotrocha bacci, O. diadema, O. gracilis, O. hartmanni, O. labronica, O. maculata, O. socialis, Perkinsiana antarctica, Perkinsiana sp Serpulidae:  Apistobranchus glacierae, Filograna/Salamacina complex, Pomatoceros lamarckii, Serpula polycerus, Spirobranchus polycerus* Sphaerodoridae:  Ephesiella mizla Spionidae:  Polydora gigardi, P. hermaphroditica, P. ligni, P. commensalis?, Spio filicornis* Spirorbidae:  Bushiella abnormis, B. atlantica, B. granulata, B. kofiadii, B. quadrangularis, Bushiella sp, B. similis, Circeis armoricana, C. oshurkovi, C. paguri, Circeis sp, Eulaeospira convexis, Helicosiphon platyspira, H. biscoensis, Janua pagenstecheri, Metalaeospira clasmani, M. pixelli, M. tenuis, Neodexiospira alveolata, N. brasiliensis, N. foraminosa, N. formosa, N. kayi, N. lamellosa, N. pseudocorrugata, N. steueri, Nidificarisa nidica, N. palliata, Paradexiospira vitrea, Paralaeospira levinseni, P. malardi, P. parallela, Pileolaria sp1, P. sp2, P. berkeleyana, P. daijonesi, P. darkarensis, P. lateralis, P. marginata, P. militaris, P. pseudoclavus, P. spinifer, P. tiarata, Pillaiospira trifurcata, Protolaeospira(?) eximia, P. pedalis, P. striata, P. tricostalis, P. triflabellis, P. stalagmia, Romanchella pustulata, R. quadricostalis, R. scoresbyi, R. solea, Simplaria postwaldi, S. pseudomilitaris, Spirorbis bifurcanis, S. corallinae, S. cuneatus, S. gesae, S. inornatus, S. infundibulum, S. rothilisbergi, S. rupestris, S. spatulatus, S. spirorbis, S. strigatus, S. tridentatus, Spirobranchus cariniferus, Vinearia zibrowii Syllidae:  Bollandia anthipathicola?, Brania protandrica, Brania pusilloides, Exogone naidina, Grubeosyllis neapolitana, Haplosyllis spongicola, Myrianida pinnigera, Pionosyllis neapolitana, Sphaerosyllis hermaphrodita, S. corruscans, Typosyllis variegata, T. vittala Terebellidae:  Alkmaria romijni Tomopteridae:  Enapleris euchaeta Typhloscolecidae:  Sagitella kowalewskii (?) Table 2.4 contd. ...

Sexual Reproduction  63

...Table 2.4 contd. Protandric hermaphrodites Archiannelidae:  Nerillidium gracile, N. macropharyngeum, N. mediterraneum, N. renaudae, N. simplex, N. troglochaetoides, Nerillidopsis hyaline, Troglochaetus beranecki Dorvilleidae:  Sabellastarte spectabilis Myzostomida:  Asteromyzostomum grygieri, Contramyzostoma, Cyclocirra, Cystimyzostomum, Endomyzostoma scotia, E. neridae, Hypomyzostoma jasoni, H. jonathoni, Mesomyzostoma botulus, M. katoi, M. lanterbecqae, M. leukos, M. lobus,  M. okadai, M. reichenspergeri,  Myzostoma cirriferum, M. debiae, M. deformator, M. eeckhauti, M. fuscomaculatum, M. glabrum, M. hollandi, M. indocuniculus, M. josefinae, M. kymae, M. laurenae, M. miki, M. pipkini, M. pulvinar, M. susanae, M. tertiusi, Notopharyngoides, Protomyzostomum lingua, P. roseus, Pulvinomyzostomum inaki, P. messingi Serpulidae:  Ficopomatus enigmaticus, Galeolaria caespitosa, G. hystrix, Hydroides elegans, H. norvegica, Mercierella enigmatica, Paradexiospira vitrea, Pomatoceros lamarckii, P. triqueter, Protolaeospira translucens, Sabellastarte magnifica, Salmacina australis, S. incrustans Syllidae:  Brania clavata, Exogone gemmifera, E. verugera, Janus knightjohnsi Spionidae:  Dipolydora commensalis Protogynic hermaphrodites Serpulidae:  Sabella microphthalma Syllidae:  Syllis variegata Serial hermaphrodites Dorvillidae:  Ophryotrocha puerilis puerilis Syllidae:  Syllis amica, S. prolifera (Policansky, 1982), Trypanosyllis zebra, Typosyllis prolifera Leech:  Helobdella striata (Kutschera and Wirtz, 1986) *of two morphs, one is hermaphroditic

~ 115,000 molluscan species, 23 and 2% are simultaneous and sequential hermaphrodites, respectively. The simultaneous hermaphroditic pulmonates (24,000 species) invariably carry an ovotestis (see Pandian, 2017) but none of the annelids bear an ovotestis. Incidentally, Davison (2006) considers ovotestis as an underdeveloped organ of evolution. From a survey, Schroeder and Hermans (1975) have reported that 67 polychaetes belonging to 23 families are hermaphrodites; of them, 31, 32 and 4 species are recognized as simultaneous, protandric (Ophryotrocha spp) and protogynic (e.g. Sabella microphthalma) hermaphrodites, respectively. Among the 23 families, the incidence of hermaphroditism is more common in Hesionidae, Syllidae, Sabellidae, Serpulidae and Nereididae. After a period of > 25 years, the reviews of Premoli and Sella (1995), Giangrande (1997) have virtually repeated the same conclusions arrived by Schroeder and Hermans. For some reasons, no attempt has been made to validate and update the conclusions of Schroeder and Hermans. From an intensive survey of relevant literature up to 2017 and computer search, the incidences of hermaphroditism in 207 species belonging to 23 families are listed in

64  Reproduction and Development in Annelida

Table 2.4. This updating has revealed that the number of hermaphrodites in polychaetes is more than doubled and implies that > 1.35% (207 out of 13,002 species) of polychaetes are hermaphrodite. This is the first updated estimate of hermaphroditism in polychates and the number is likely to increase. Notably, all the 70 species belonging to the family Spirorbidae are SH (Kupriyanova et al., 2001). A computer search has revealed the presence of ~ 26 myzostomid species, all of them have been considered as protandrics. Though broadly classified into (i) simultaneous, (ii) protandric, (iii) protogynic and (iv) serial hermaphrodites, Schroeder and Hermans have grouped them into seven types but assigned ~ 20 species to one or other type with the question mark. Hence, the classification of hermaphroditic types remains very fluid. At least, a few species may be shifted from one to other type. A vast majority of polychaetes are gonochores. However, of ~ 13,000 polychaete species (Table 1.2), > 207 species (see Table 2.4) (1.35% of polychaetes, 0.6% of annelids) are hermaphrodites. Both oligochaetes (3,155 species, 18.7% of annelid species) and hirudineans (684 species, 4.0%) are hermaphrodites. This is perhaps the first estimate to indicate that the annelid comprises of ~ 76% gonochorics and 24% hermaphrodites. This estimate of 76% gonochorism in annelids may be compared with 75% in molluscs (Pandian, 2017), 92% in crustaceans (Pandian 2016) and 99% in echinoderms (Pandian, 2018). Notably, a few observations on the polychaete hermaphroditism may be mentioned. In the serpulid Spirobranchus polycerus, Marsden (1992) has reported the presence of seven opercular horned SH morph inhabiting singly or in a small group on the live hydrozoan coral Milliporea complanata and two horned gonochoric morph living in the low tidal zone. Lacking seminal receptacle, the serpulid Laonome albicingillum is a self-fertilizing SH (Hsieh, 1997). With sequential sex change, the protandrics exhibit different sex ratios. Figure 2.3 summarizes the ontogenetic pathways, through which protandric to female, protogynic to male and SH are generated by sex change in polychaetes. Notably, the female gonad is simply added in the protandric to SH. Hence, these protandrics do not undergo sex change but only adds female sex. The ratio of ♂ + : ♂ in the protandric hermaphrodites is 0.61 : 0.39 in Ophryotrocha diadema and 0.70 : 0.30 in O. gracilis (Sella and Ramella, 1999). An interesting protandric is the sabellid Sabellastarte spectabilis. The expression of sex ratio has been assessed as ♂ : ♀ : ♂ + : 0 (unknown) in small (6–8 mm), medium (9–10 mm) and large (11–13 mm) worms. The ratios increase from 0.13 to 0.30 and 0.07 to 0.31 in females and hermaphrodites, respectively. In the same sizes, it decreases from 0.56 to 0.25 and 0.24 to 0.14 in males and unknowns, respectively (Bybee et al., 2006). Notable is the persistence of males even in the largest size of protandric polychaete hermaphrodites at ratios ranging from 0.14 in S. spectabilis to 0.39 in O. diadema.

Sexual Reproduction  65

FiguRe 2.3 Ontogenetic pathways through which sex is changed or sex ratios are changed in some hermaphrodites. O represents unknown sex.

2.4 Parthenogenesis A number of authors have defined parthenogenesis (e.g. Suomalainen, 1950, Beatty, 1967). In view of polyploid automictic parthenogens, in which triploid (3n) parthenogenic eggs require activation by a haploid sperm from diploid sexual morph, parthenogenesis is defined as the generation of embryo from an egg without any genetic contribution by a sperm. For the first time, the incidences of parthenogenic annelids are assembled in Table 2.5. Of > 75 parthenogenic annelid species, 75% are earthworms. The ability of earthworms for asexual reproduction is limited (e.g. Xiao et al., 2011), in comparison to that of enchytraeids and naidids. Understandably, earthworms have opted for parthenogenesis to reduce the cost of developing and maintaining a dual reproductive system. The incidence of parthenogenesis

66  Reproduction and Development in Annelida

Table 2.5 Parthenogenic annelids (compiled from many sources) Polychaetes:  Dodecaceria concharum, D. pulchra (Gibson, 1977), Brania pusilla (Hauenschild, 1955) Tubificids:  Tubifex tubifex, Limnodrilus hoffmeisteri, L. mastix, Lumbriculus variegatus (Poddubnaya, 1984, Brinkhurst, 1986) Enchytraeids:  Cognettia glandulosa, Fridericia galba, F. ratzeli, F. striata, Lumbricillus lineatus, Mesenchytraeus glandulosus, Achaeta bilobisa?, F. bisetosa?, F. connata? (Christensen, 1961) Earthworms:  Allolobophora trapezoides, Amynthas bileatus, A. catenus, A. chilanensis, A. corticus, A. cruxus, A. diffringens, A. gracilis, A. hohuanmontis, A. hupiensis, A. shinammontis, A. tokioensis, Aporrectdodea caliginosa, A. rosea, A. trapezoides, Bimastos beddardi, B. gieseleri, B. heimburgeri, B. longicinctus, B. palustris, B. parvus, B. tumidus, B. welchi, B. zeteki, Drawida hattamimiju, D. nepalensis, Dendrobaena octaedra, D. rubida, Dendrodrilus rubidus, Eisenia rosea, Eiseniella tetraedra, Eukerria saltensis, Heliodrilus hachiojii, Lumbricus eiseni, L. terrestris, Microscolex dubius, M. phosphoreus, Metaphire hilgendorfi, Ocnerodrilus occidentalis, Octolasion cyaneum, O. occidentalis, O. tyrtaeum, Onychochaeta windlei, Pheretima agrestis, P. bicinta, P. diffringens, P. hilgendorfi, P. hupiensis, P. levis, P. loveridgei, Piutellus papillifer, P. umbellulariae, Pontoscolex corethrurus, Dichogaster bolaui?, Allolobophora muldali (PF) (Reynolds, 1974, Jaenike and Selander, 1979, Blakemore, 2003, Tsai et al., 2007).

is limited to a couple of polychaetes and to a few species in tubificids and enchytraeids. At present, there is no report on the incidence of parthenogenic hirudinean. Incidentally, Gibson (1977) has stated that Dodecaceria concharum and D. pulchra are parthenogens, as males are not found and 70% of the mature worms have oocytes. However, he has not provided any additional evidence to confirm that these polychaetes are parthenogens.

2.4.1  Parthenogenic Types In annelids, the origin of parthenogenesis can be traced to two sources: (i) the repeated polyphyletic origin of new parthenogens from sexual ancestors (e.g. Fridericia striata, Christensen et al., 1989) and (ii) monophyletic origin through accumulation of new mutations within an existing lineage. Cytological studies have revealed the presence of three major parthenogenic types: 1. Ameiotic/apomictic parthenogens. In them, synapsis and bivalent formation do not occur (e.g. Dendrobaena octaedra, Suomalainen et al., 1987). 2. Automictic/gynogenic parthenogens retain features of normal meiosis including crossing over and so on, albeit no crossing over in polyploids. Whereas meiosis in diploids is chiasmatic, polyploids undergo synaptic meiotic division in the oocytes (Christensen, 1960, 1961, 1980). However, the haploid egg pronucleus restores diploidy by fusing with that of homologous second polar body (e.g. Lumbricillus lineatus). This is also true of the family Lumbricidae and in the tubificid genus Limnodrilus (Christensen, 1984). Interestingly, the eggs of triploid and other polyploid parthenogens do require activation by sperm from diploid sexual morph. Consequently, if sperm producing morphs are absent, there will be no triploids (Christensen

Sexual Reproduction  67

et al., 1978). 3. In telytochorous parthenogens, in which pre-meiotic doubling of chromosomes at the last oogonial divisions result in endomitosis followed by the formation of chiasmatic bivalents and regular meiosis with extrusions of two polar bodies. The genetic consequences of this cytological mechanism are parallel to that of apomictics, as synapsis is limited between sister chromosomes that are exact molecular copies of one another (see Diaz-Cosin et al., 2011). As a result, diploid eggs are produced without recombination leading to production of genetically indistinguishable clones (e.g. Tubifex tubifex, Christensen, 1984). Nevertheless, electrophoretic analysis of 3n parthenogenic Fridericia galba has revealed the presence of 13 different clonal phenotypes (Christensen et al., 1992). Similarly, allozyme analysis can also discriminate selfers from parthenogens by comparing heterozygous loci of an individual and its related uniparental progeny (see Baldo and Ferraguti, 2005). Notably, of 33 lumbricid species found in North America, 17 of them reproduce primarily or exclusively by parthenogenesis, i.e. they are obligate parthenogens (Jaenike and Selander, 1979). There are also facultative (e.g. Allolobophora muldali) and amphimictic cum parthenogens (e.g. Diplocardia singularis, Dendrobaena rubida, Eisenia foetida) (Reynolds, 1974). Incidentally, despite his elaborative cytological study on oogenesis in parthenogenic worms (e.g. Christensen, 1980), Christensen has not considered the centrosomes and their derivative centrioles. Their function is important for assembling the spindle apparatus that organizes separation of chromatids. In most animals, centrioles are degraded during oogenesis and are inherited paternally through the sperm (Engelstadter, 2008). It is not clear whether centrioles are either not degraded during oogenesis or synthesized de novo in the parthenogenic eggs of annelids.

2.4.2  Parthenogenic Levels The existence of facultative parthenogens and amphimictic parthenogens clearly indicates that components of male reproductive system are not stably maintained. In his summarizing note, Gates (1971) has reported the gradual but progressive reductions in components of male reproductive system of parthenogenic oligochaetes. Accordingly, the reductions are from four to three pairs of testes, two to one pair of seminal vesicles, two to one pair of prostate, two to one pair of spermatheca and disappearance of oviducts and male pores. Reynolds (1974) has indicated that despite retention of one or more of these male components, sperms may be absent. For example, sperms/spermatophores are absent in some parthenogenic earthworms, despite the presence of testes and seminal vesicles. In Tubifex tubifex, the function of the testes diminishes and the spermatogonia are not replenished with the development of ovary. In fact, the process of spermatogenesis is not completed and ceases at the spermatid stage (Poddubnaya, 1984). In the parthenogenic earthworms too, the spermatogenesis may be arrested at spermatid stage, despite the presence of one or more components of

68  Reproduction and Development in Annelida

Table 2.6 Levels of reduction in components of male reproductive system in the Taiwanish parthenogenic earthworms (condensed and compiled from Tsai et al., 2007) Spermatheca

Prostate Glands

Seminal Vesicles

Amynthas shinanmontis Absent 10/20 One pair 1/20 Two pairs 2/20 Three pairs 7/20

Absent 3/20 Vestigial 3/20 Nodular 13/20 Normal 1/20

Vestigial 5 Medium 3 Normal 12

A. chilanensis Absent

Absent

Small

A. sheni Absent

Small

Small

the reproductive system. Interestingly, Tsai et al. (2007) have described a spectrum of reductions or loss of spermatheca, prostate glands and seminal vesicles in Taiwanese earthworms. Irrespective of the presence of seminal vesicle in Amynthas shinanmontis, the number of spermatheca is reduced from four to two pairs in 2 out of 20 specimens collected (Table 2.6). Clearly, the parthenogens may have originated from sexual oligochaetes by a stroke of one or more mutations arresting spermatogenesis in parthenogenic oligochaetes but the components of male organs are reduced gradually with more and more mutations.

2.4.3  Ploidy Levels In oligochaete parthenogens, polyploidy is the most common phenomenon. An advantage of polyploidy in parthenogens is the increased scope for genetic diversity (Diaz-Cosin et al., 2011). Table 2.7 indicates that the number of polyploid parthenogenic species is more in lumbricids than enchytraeids. Diaz-Cosin et al. (2011) have listed the following advantages for polyploid oligochaetes, especially the earthworms: 1. High levels of heterozygosity and exceptionally fit genomes that are sustained and inherited by avoiding recombination and segregation. 2. Manifestation and maintenance of a single reproductive system facilitate relatively higher growth and reproductive rates. The questionable parthenogenic singletons of Octolasion cyaneum grow faster at the rate of 204 mg/g/d, in comparison to the twins generated by cross fertilizers (177 mg/g/d). With diploid and haploid eggs in them, cocoon production is 0.8 and 1.55 cocoon/w for the singletons and twins, respectively. The singletons generate also 32 mg weighing hatchling, in comparison to cross fertilizing twins producing 18 mg weighing hatchling (Lowe and Butt, 2008). In Aporrectodea trapezoides, parthenogenic singletons attain sexual maturity at the age of 153 days and 1 g size, and produce 2.03 cocoon per

Sexual Reproduction  69

w. 3. The no need to search for a mate and mating expedite colonizing ability and 4. Polyploid and polymorphic morphs are generated from selection at the level of genomes. The level of polyploids ranges from triploid, tetraploid, pentaploid, hexaploid and heptaploid in lumbricids and enchytraeids (Table 2.7). Of 319 parthenogens collected from Australia, Canada and Europe, 54.5, 6.0, 39.5 and 1.0% are diploids, triploids, tetraploids and pentaploids, respectively (Coates, 1995). Briefly, the incidence of polyploid parthenogens is more fecund in enchytraeids than in lumbricids (Table 2.8). Table 2.7 Polyploids in parthenogenic oligochaetes (compiled from Jaenike and Selander, 1979, Coates, 1995) Species

Ploidy

Chromosomes (no.)

Lumbricidae Aporrectodea trapezoides

2n 3n

36 54

Octolasion tyrtaeum

2n 3n 4n

38 54 72

A. rosea

3n 5n 6n

54 90 108

Dendrodrilus rubidus

2n 4n 6n

34 68 102

D. octaedra

6n 7n

108 124

Enchytraeidae Lumbricillus lineatus

2n 3n 4n 5n

26 39 52 65

Table 2.8 Polyploid distribution in Lumbricidae and Enchytraeidea (modified from Coates, 1995) Lumbricidae

Enchytraeidea

Species (no.)

56

79

2n species

37

47

Parthenogens (no.)

0

1

In + Polyploidy species (no.)

9

9

Polyploidy species (no.)

10

23

Parthenogens (no.)

8–9

4

70  Reproduction and Development in Annelida

2.5 Fecundity Fecundity is a decisively important factor in recruitment and sustaining a population. By providing space to accommodate the oocytes/eggs, body size becomes a critically important factor in determining fecundity. Sexuality and egg size are also important factors in deciding the level of fecundity. Beside these, food supply (e.g. Dinophilus gyrociliatus, diet quality, Prevedelli and Vandini, 1999), temperature (e.g. D. gyrociliatus, Akesson and Costlow, 1991), salinity (e.g. D. gyrociliatus, Akesson and Costlow, 1991, Neanthes limnicola, Fong and Pearse, 1992) and photoperiod (e.g. Streblospio benedicti, Chu and Levin, 1989) also play a role in deciding fecundity. The total number of oocytes contributing to fecundity is assured by waves of oogonial proliferation and subsequent oocyte recruitment (see Pandian, 2013). With regard to fecundity of annelids, the terms used by fishery biologists have to be introduced. Firstly, Batch Fecundity (BF) or a clutch is the number of eggs produced per spawning or brood. It is a function of body size (Length, L or Weight, W). BF is related to the volume of space available in the (intra-ovarian) ovary or (extra-ovarian) coelomic cavity. To accommodate maturing oocytes, geometry (F = aL/Wb) suggests that the length/weight exponent would be 3.0. Unlike broadcasting oviparous fishes, ~ 50% of polychaetes brood their eggs and the clitellates encapsulate them in cocoons. Though these features complicate BF relation to body size, BF increases with increasing body size in many annelids (Figs. 1.10D, E, F, 1.11A, B). In annelids, growth is expressed mostly in length, girth in some (e.g. Hediste japonica) and breadth in others (e.g. Hirudo medicinalis). With these morphological differences, the fecundity-body size relation may tilt the expected exponent of 3.0. However, no information is yet available on this aspect. Secondly, Potential Fecundity (PF) is the maximum number of oocytes commencing to differentiate and develop. However, due to one or other environmental factors like food supply, a fraction of these developing oocytes may not be developed or resorbed through atresia. For example, ~ 42% of the pre-vitellogenic oocytes released into the coelom in Spirorbis spirorbis are matured representing the Realized Fecundity (RF) (Fig. 2.4B). Life time Fecundity (LF) is a cumulative number of eggs produced in different batches/seasons. For example, S. spirorbis produces five batches of broods during its LF totaling to 633 eggs (Daly, 1978b). Relative Fecundity (RF) is the number of eggs/oocytes per unit weight of the body. For example, it is ~ 1.0 egg/µg dry body weight of S. spirorbis (Daly, 1978b). It provides a scope for analysis of the reproductive performance with reference to body size within a species, populations from different geographical areas and different years within a geographical area as well as between comparable species. Thirdly, a distinction must also be made between species with determinate and indeterminate fecundity. In the former, the PF is fixed prior to the

Sexual Reproduction  71

commencement of spawning or a spawning season. It depends on the stored energy reserves from the body and/or storage organs, as the chloragogue in oligochaetes and blood in leeches; hence the clitellates are all capital breeders (see Pandian, 2015). In them, development of oocytes is likely to be synchronized. In the indeterminates, the PF, however, is not fixed and the development of oocytes is asynchronized (e.g. Perkinsiana spp, Gambi et al., 2000). Their PF depends on incoming energy resource and are not capital income breeders (see Pandian, 2015). The determinate annelids adopt two different spawning strategies: (i) synchronous determinate total spawning (e.g. semelparous Nereis virens, Fig. 2.7A) and (ii) synchronous determinate iteroparous spawning (e.g. Marphysa sanguinea, Fig. 2.7C). On the other hand, consequent to low incoming energy, oocytes are asynchronously developed in those characterized by indeterminate fecundity, resulting in successive spawning during a spawning season (e.g. Nephelopsis obscura, Peterson, 1983). In them too, fecundity decreases with advancing time/age and increasing body weight. Though these fecundity traits are applicable to annelids, they remain to be tested in more number of species.

2.5.1 Sexuality Within hermaphroditic oligochaetes, incidence of parthenogenics is not uncommon. A fact that is known but not adequately recognized is that the establishment and maintenance of dual sex within a single individual cost resources. In them, resource allocation for reproduction is limited with the consequence of reduced fecundity. But the reduced fecundity is compensated by (i) internal fertilization ensuring a higher level of fertilization success and (ii) embryonic development within a hard cocoon affording fairly high level of protection. However, these two clitellate features may further reduce fecundity. Within the genus Ophryotrocha, there are gonochoric and hermaphroditic species, providing a rare but excellent opportunity to study the effect of sexuality on fecundity. Fecundity values range from 80 eggs in gonochoric O. macrovifera to 200 eggs in O. robusta but from 11 eggs in hermaphroditic O. gracilis to 50 eggs in O. maculata (Table 2.9). Hence, the mean fecundity is 9.4 eggs/d and 4.5 eggs/d in gonochoric and hermaphroditic Ophryotrocha spp, respectively, despite the interspawning period averaging to 13.6 days and 4.8 days in gonochorics and hermaphrodites, respectively. Not surprisingly, maintaining the dual sex within an individual hermaphrodite reduces the batch fecundity to half of that gonochorics. To reduce the cost of maintaining dual sex, many earthworms have ‘sacrificed’ one or more organs in the male reproductive system (Table 2.6). Consequently, some of these ‘female hermaphrodites’ switched to parthenogenesis (Diaz-Cosin et al., 2011). To enhance genetic diversity, ploidy level has been increased up to heptoploidy in some of these parthenogenic clones (Table 2.7). Hence, elevated ploidy is also expected to express in their eggs. For example, diploid and triploid eggs are produced by Aporrectodea

72  Reproduction and Development in Annelida

Table 2.9 Effect of sexuality on fecundity in gonochoric and hermaphroditic species in polychaete genus Ophryotrocha (from Premoli and Sella, 1995, modified) and hermaphroditic and parthenogenic earthworms (c = cocoon) Species

Fecundity (egg no./spawn)

Inter-spawning Interval (d)

Gonochoric polychaetes O. labronica

130

O. macrovifera O. notoglandulata O. robusta Mean

11.1

80

11.3

115

14.0

200

18.1

9.4 eggs/day Hermaphroditic polychaetes

O. diadema

25

2.9

O. gracilis

11

6.2

O. hartmanni

30

5.3

O. maculata

50

–

O. socialis

25

–

Mean

4.5 eggs/day Hermaphroditic earthworms

Eudrilus eugeniae (Mba, 1988)

10.27 c/w

Eisenia foetida (Siddique et al., 2005)

9.0 c/w

Apporrectodea longa (Lowe and Butt, 2005)

0.33 c/w

Allolobophora chlorotica (Lowe and Butt, 2005)

0.20 c/w

Mean

4.95 c /w Parthenogenic earthworms

Aporrectodea trapezoides (Fernandez et al., 2010)

2.03 c/w

Pontoscolex corethrurus (Bhattacharjee and Chaudhuri, 2002)

2.30 c/w

Drawida nepalensis (Bhattacharjee and Chaudhuri, 2002)

0.46 c/w

Aporrectodea caliginosa (Lowe and Butt, 2005)

0.61 c/w

Lumbricus terrestris (Lowe and Butt, 2005)

0.46 c/w

Mean

1.17 c / w

trapezoides (Table 2.7). Considering the relatively more frequent incidence of parthenogens in earthworms, available information on fecundity of hermaphrodites and parthenogens is summarized in Table 2.9. Expectedly, the mean cocoon production is 1.17/week (w) and 4.95/w in parthenogens and hermaphrodites, respectively. Hence, the switch to parthenogenesis and polyploidy may have facilitated the survival of these earthworms in ‘patchy’ habitats but certainly at the cost of fecundity. Briefly, both hermaphroditism and parthenogenesis have significantly reduced batch fecundity.

Sexual Reproduction  73

2.5.2  Oogenic Anlage Being a key factor, food quality and availability may affect the age at sexual maturity. In the sediment feeding Capitella sp I, the age is significantly increased from 28 days in those feeding sediment containing 3.0% Total Organic Matter (TOM) to 56 days in those feeding sediment containing only 0.25% TOM (Ramskov and Forbes, 2008). Oogenesis can be intraovarian, as in Streblospio spp and Branchipolynoe seepensis with 10–300 brooded eggs involving indirect development in the former and direct one in the latter (Table 2.11) or extraovarian, as in most polychaetes. In the dorvilleid Veneriserva pygoclava meridionalis, vitellogenesis involves progressive reduction in the number from about 100 pre-vitellogenic oocytes of 50 µm to 25 eggs each measuring > 150 µm (Fig. 2.4A). Presumably, oogenesis is intraovarian in V. pygoclava meridionalis, in which only 25% oocytes are matured. In the extraovarian oogenesis, about 50% oocytes are developed into mature eggs, as in the serpulid Spirorbis spirorbis (Fig. 2.4B). With comparatively far less space available in the intraovary, less number of pre-vitellogenic oocytes is expected to mature; hence, the anlage, in which oocytes undergo vitellogenesis (see p 31) and maturation, imposes a profound effect on fecundity.

Figure 2.4 Progressive decrease in the number of oocytes from pre-vitellogenic to vitellogenic/mature egg stage in the presumably intraovarian A. Veneriserva pygoclava meridionalis with increasing egg size and B. extraovarian Spirorbis spirorbis during succesive calendar months. Note the levels of variance indicated by thin vertical lines in Fig. 2.4B (modified and redrawn from Micaletto et al., 2002, Daly, 1978b).

2.5.3  Egg Size In polychaetes, egg size ranges from 30 µm (e.g. Protodrilus albicans, Westheide, 1984) to 600 µm (e.g. Thelepus cincinnatus, Garraffoni et al., 2014) and to

74  Reproduction and Development in Annelida

1,000 µm (e.g. Myxicola cf sulcata, Gambi et al., 2001). It has a profound effect on fecundity and recruitment. From his survey, Giangrande (1997) has grouped them into semelparous and iteroparous forms: the latter is divided into short- and long-living forms. The short living forms may produce two broods, the fecundity of the second brood being strongly influenced by resource availability (e.g. Harmathoe imbricata, see Cassai and Prevedelli, 1998a). But the long living forms generate about five broods (e.g. Spirorbis spirorbis, Daly, 1978b). Of 32 families (sabellids, spionids and syllids represented in two groups) considered by Giangrande, 7, 9 and 16 families comprise semelparous, short- and long-living iteroparous forms, respectively. Clearly, long living iteroparity is the most common reproductive strategy of polychaetes. On the basis of their reproductive mode, these polychaetes are considered in this analysis under 1. planktotrophic (PLK), 2. lecithotrophic (LEC) and 3. direct developers (DIR). The latter is considered in two subgroups, in which the egg size measures > 100 µm (DIR1) and < 100 µm (DIR2), which obviously draw nutrients from the mother. From Table 2.10, the following may be inferred: 1. expectedly, the mean egg size increases from 113 µm in PLK to 196 µm and 209 µm in LEC and DIR1, respectively but decreases to 83 µm in DIR2. The variations from the respective mean are very wide in the first three groups but the least in DIR2 subgroup, 2. within the families considered, the choice for reproductive strategy decreases in the following order: long living iteroparity > short living iteroparity > semelparity; for the reproductive mode, it is PLK > LEC > DIR1 > DIR2. Notably, the long living iteroparous species do not invest on viviparity. Within a species, egg size remains equal in worms of different body weights (e.g. Galeolaria caespitosa, Fig. 1.12A). However, contrasting trends become apparent, when fecundity-egg size relation is considered at interspecific level. Expectedly, fecundity is decreased with increasing egg size in semelparous and iteroparous polychaetes characterized by indirect life cycle with holoplanktonic or meroplanktonic (brooder) larva (Fig. 2.5A). With small oocyte size (112 µm), the semelparous broadcasters spawn Table 2.10 Egg size (µm) in planktotrophic (PLK), lecithotrophic (LEC) and directly (DIR) developing polychaetes. Values in brackets represent the number of families and square brackets the range of egg size (compiled from data reported by Giangrande, 1997) Developmental Mode

PLK

LEC

DIR1, egg size > 100 µm

DIR2, egg size < 100 µm

Semelparous Iteroparous short living long living Total Range

98 (2)

205 (2)

178 (2)

96 (1)

113 (4) 123 (8) 112 (14) [64–155]

175 (1) 196 (5) 196 (8) [122–250]

142 (1) 241 (4) 209 (7) [142–290]

178 (2) – 83 (3) [72–96]

Sexual Reproduction  75

Figure 2.5 A. Effect of oocyte/egg size on fecundity of (●) semelparous and (▲) iteroparous polychaetes (drawn using data reported for Nereis virens (Creaser and Clifford, 1982), Glycera dibranchiata (Creaser, 1973), Perinereis cultrifera (Cassai and Prevedelli, 1988a), Hediste japonica (Sato, 1999), Nicolea zostericola (Eckelbarger, 1973), Eupolymnia crescentis, Neoamphitrite robusta, Thelepus crispus and Lanice conchilega (McHugh, 1993). B. Effect of egg size on fecundity in dorvilleid polychaete embryos with (●) maternal, (■) biparental and (▲) communal care (drawn from the data reported by Sella and Ramella, 1999), C. Effect of body length/oocyte size on fecundity of interstitial polychaetes. The relationship in smaller interstitial worms is shown in a window (drawn using data reported by Westheide, 1984).

millions and thousands of eggs, while the iteroparous brooders generate a few thousands or hundreds of larger (196, 209 µm) eggs. Interestingly, a slightly negative but linear relation is also apparent between egg size and fecundity in Ophryotrocha spp that exhibit biparental care (Fig. 2.5B). Hence, egg size significantly imposes reduction in fecundity of broadcasters and others displaying parental care. In some subgroups of sabellid brooders a positive linear relation between log body size and log fecundity is apparent (Fig. 1.10E). However, this positive relation may be altered, when normal values are considered (cf Fig. 2.5A, B, Fig. 2.7H, I). On the other hand, increasing body size may also increase egg size, when the relation is considered at interspecies level. For example, the oocyte/egg size increases with increasing body length in broadcast spawning nereidids, dorvilleidid and dinophilid as well as extra- and intra-tubular brooding polychaetes (Fig. 1.10C, D, E, F), although the positive linear relation in Fig. 1.10E, F are based on log values. Unusually, planktotrophic duration increases with increasing egg size in spionids (Fig. 1.10A). In them, with increasing egg size, however, the chaetigers stage, at which they are released, also increases (Fig. 1.10B). At this point, a brief account on the interstitial polychaetes becomes necessary. Represented by as many as 16 families (Westheide, 1971), these polychaetes constitute one of the most species-rich and number-rich communities of the marine fauna. In an impressive concept, Westheide (1984) has characterized their life history traits. Inhabiting within the interstitial space of ~ 125 µm, (i) they are small (1–2 mm body length, the smallest

76  Reproduction and Development in Annelida

Diurodrilus sp measuring 300 µm), (ii) mostly hermaphroditic (e.g. Nerillidae, Microphthalmus), (iii) fertilization by pseudocopulation (Ophryotrocha gracilis)/hypodermic injection (e.g. Protodrilus albicans) of limited sperms (with considerable morphological alterations including the lack of flagellum and loss of mid-piece, Pisione remota) and (iv) brood/gestate their eggs (O. vivipara). In the interstitial polychaetes, body size is measured in number of times of the egg diameter (Fig. 2.5C). Fecundity also increases with increasing body length/oocyte diameter from 45 times in Hesionides arenaria with 70 oocytes to 400 times in P. albicans with 1,200 oocytes in fairly large worms. In smaller polychaetes, it is, however, from four times in Ikosipodus carolensis with one–two oocytes to 20 times in M. sczelkowii with 11–15 oocytes.

2.5.4  Body Size In a rare investigation, Daly (1978b) has made two important observations in the serpulid Spirorbis spirorbis. The extent of individual variations in the output of pre-vitellogenic oocytes by the 2.9 mm size class ranges between 25 and 35% (Fig. 2.4B). The oocytes mature into vitellogenic ones in the coelomic cavity. Of the pre-vitellogenic oocytes released from the ‘ovary’, ~ 50% of them mature into vitellogenic ones during the period from January to April. From his estimates on changes in the mean number of previtellogenic and vitellogenic oocytes during successive broods, Daly has estimated that the gonadal release of pre-vitellogenic oocytes compensates the number of oocytes entering vitellogenesis. Secondly, the oocyte output in the first brood is positively correlated with body size and increases from 24 in 2.1 mm coil diameter body size (= 75 mg dry body weight) to 59 but from ~ 16 to ~ 28 in 3.1 mm body sized (= 225 mg body weight) (Fig. 2.6A). However, these values decrease to lower levels from 64–25, 54–24 and 24–20 in the first, second and fifth broods, respectively (Fig. 2.6B). Hence, fecundity decreases with successive broods in relation to increasing body size. In other words, it decreases from 1.2 eggs/µg dry body weight in the smallest worm to 0.97 egg/µg dry body weight in the largest worm. In polychaetes, reproductive modes are greatly diverse. Not surprisingly, the patterns of spawning and fecundity are also equally diverse. Spawning is a critically important event to release the time-long gametic investment and polychaetes, as other animals, select a strategically important location and time for spawning to ensure higher fertilization success and potential offspring survival. Some of these reproductive modes are listed below: (i) Oogenesis can be synchronized in broadcasters involving epitoky and semelparity, as in Glycera dibranchiata with a few thousands/millions oocytes or (ii) iteroparous synchronized atokous spawner, as in Marphysa sanguinea with a few thousand oocytes or (iii) asynchronous broadcast spawning, as in Eupolymnia crescentis involving indirect development of 128,500 oocytes (Table 2.11). In iteroparous brooders, (iv) development can be indirect with

Sexual Reproduction  77

discrete synchronized spawning, as in Nicolea zostericola with 665 eggs/y, (v) asynchronized spawning over a period of 6 months, as in Thelepus crispus with 515,555 eggs, (vi) asynchronous oocyte maturation with continuous spawning of 44 eggs/brood throughout the year and brooding 11 sequential broods at a time, as in Ramex californiensis and (vii) direct development of self-fertilized ~ 150 gestated embryos parturited once in 6 months, as in viviparous Neanthes limnicola. In others, (viii) spawning is continuous and lasts almost throughout the year with 11 eggs/w and 3.5 egg masses/d, as in Dinophilus gyrociliatus and Ophryotrocha adherens, respectively. (ix) In oligochaetes, cocoons are generated at the rate of 2.25–3.14/w in earthworms, 0.15/w in Branchiura sowerbyi but 10 eggs per w in Tubifex tubifex. (x) In hirudineans, the inverted ‘U’-shaped spawning trends within a reproductive bout are presumably regulated by blood ingestion in sanguivorous Hirudo medicinalis and actually by temperature in carnivorous Nephelopsis obscura. The large H. medicinalis can gorge itself sucking ~ 10 g blood at a time (Davies and McLoughlin, 1996). Clearly, the reported values for the observed fecundity are so diverse that it is difficult to consider them comparable for following reasons: 1. The number of gametogenic (ovarian) segments varies from two in Spirorbis spirorbis (Daly, 1978a) to a large number in nereidids. Hence, the fecundity– body size relationship becomes not comparable with such wide variations in the number of ovarian segments. 2. The few eggs developed and matured intraovarially cannot be compared with iteroparous brooders having thousands of eggs and semelparous broadcasters having millions of oocytes

Figure 2.6 Fecundity in Spirorbis spirorbis: A. increasing fecundity with increasing body size, B. decreasing fecundity as function (of body size) in successive broods from 1 to 5 (drawn from data reported by Daly, 1978b).

F = 11, 4 and 5.5 eggs per ovary in PLK, LEC and hybrid morphs, respectively (Levin and Bridges, 1994).

Anterior gametic segments extend from 21 to 30 chaetigers. 10–34 eggs (Lardicci et al., 1997). F decreases from ~ 60 eggs on 60th d to 10 eggs on 161st d and 500 d in PLK and LEC (Bridges and Heppell, 1996).

S. benedicti

Thelepus crispus 515,555 eggs/brood spawned during 6 months (McHugh, 1993).

Nicolea zostericola

Male and female pair prior to spawning. Single discrete spawning of 665 eggs/y (Eckelbarger, 1973).

Indirect

100–300 eggs/♀ brooded in I (125 µm) and II (300–350 µm) cohorts. Direct development (Jollivet et al., 2000).

Branchiopolynoe seepensis

44 eggs/coc. Asynchronized oocyte maturation and sequentially laid 11 coc. Breeding throughout y (McHugh, 1993).

Ramex californiensis

Direct

~ 150 embryos/brood. Breeding twice/y (Fong and Pearse, 1992).

Neanthes limnicola

Viviparous

128,500 oocytes spawning lasts for 3 months. Indirect development (McHugh, 1993).

Eupolymnia crescentis

Iteroparous, Asynchronous

Asynchronous

F increases 8,500 to 24,300 oocytes in 0.7 and 3.0 g size, respectively. Indirect development (Cassai and Prevedelli, 1998b).

F increases from 0.7 to 9.5 million oocytes in 17 and 47 cm body size, respectively. Indirect development (Creaser, 1973).

Semelparous, synchronous

Marphysa sanguinea

Glycera dibranchiata

Iteroparous brooders

Iteroparous, synchronous atoky

Semelparous, synchronous epitoky

Extraovarian oogenesis: Broadcast spawners

S. benedicti

Streblospio shrubsolii

Intraovarian oogenesis

Fecundity (F) and modes of oogenesis, spawning and breeding by annelids, as reported by respective authors. coc = cocoon

Table 2.11

78  Reproduction and Development in Annelida

Nephelopsis obscura.  An inverted U trend for F within a reproductive bout (Holmstrand and Collins, 1985).

Hirudinea:  Hirudo medicinalis. F increases with increasing age, body weight and blood ingestion (Davies and McLoughlin et al., 1996) (Fig. 2.7F, G, H).

Tubificid:  Tubifex tubifex ~ 10 eggs/w. F, as coc no. decreases with increasing age (Pasteris et al., 1996) (Fig. 2.8D, E).

Naidid:  Branchiura sowerbyi 0.15 coc/w. F, as coc no. decreases with increasing age (Lobo and Alves, 2011) (Fig. 2.8B).

Eisenia foetida 2.6 coc/w. (Siddique et al., 2005).

Hyperiodrilus africanus 2.25 coc/w. in paired worms; 1.12 coc/w. in singletons (Tondoh and Lavelle, 1997).

Earthworms:  Lumbricus terrestris 3.14 coc/w. F, as coc no. decreases with increasing age (Lowe and Butt, 2005) (Fig. 2.8A).

Clitellates

Protandric Ophryotrocha adherens ~ 3.5 egg mass/d. F, as coc no. decreases with increasing age (Paavo et al., 2000).

Dinophilus gyrociliatus ~ 11 eggs/w (30‰ S, tetramin) F, as coc no. decreases with increasing age (Prevedelli and Simonini, 2000).

Continuous spawners

Sexual Reproduction  79

80  Reproduction and Development in Annelida

developed and matured in the coelom. In terms of space to accommodate eggs/oocytes, there can be a vast difference between the ovary and coelom. 3. The thousands-millions of oocytes spawned by semelparous epitokous broadcasters are considered as batch (BF) as well as lifetime fecundity, in contrast to the PF consisting of few thousand oocytes of iteroparous, atokous broadcasters. 4. Within iteroparous brooders, the counts for fecundity in synchronous and asynchronous spawners are not comparable. In iteroparous asynchronous spawners characterized by indirect development like T. crispus produce > 500,000 eggs, which may not be comparable to the 150 embryos gestated by self-fertilizing viviparous N. limnicola, a biannual breeder. 5. R. californiensis is not comparable with any of the above, as it simultaneously broods 11 sequentials broods, each consisting of 44 eggs. A second complication is that different morphological features have been considered to describe fecundity-body size relationship: (i) body length (e.g. Nereis virens, Fig. 2.7A, body length/oocyte length, Fig. 2.5C), (ii) body width (e.g. Hediste japonica, Fig. 2.7B), (iii) number of chaetigers (e.g. Ophryotrocha puerilis puerilis, Fig. 2.7D), (iv) number of segments (e.g. Enchytraeus variatus, Fig. 2.8C), (v) body weight (e.g. live body weight: Tubifex tubifex, Fig. 2.8E; dry body weight Spirorbis spirorbis, Fig. 2.6A, B), and (vi) age (e.g. T. tubifex, Fig. 2.8D). Notably, similar trends for the fecundity-body size relationship have been reported, when body size is considered in units of body weight

Figure 2.7 Fecundity in broadcast spawning polychaetes as function of A. body length in semelparous epitokous Nereis virens (Creaser and Clifford, 1982), B. body width in estuarine (large eggs) and marine (small eggs) forms of semelparous epitokous Hediste japonica (Sato, 1999), C. body weight in iteroparous Marphysa sanguinea (Cassai and Prevedelli, 1998b) and D. chaetiger number in protandrus continuous spawner Ophryotrocha puerilis puerilis (Berglund, 1991). Fecundity in sanguivorous hirudineans as function of body weight in E. the giant leech Haementeria ghilianii (Sawyer et al., 1981) and in Hirudo medicinalis as functions of F. age and G, H. ingestion of blood (Davies and McLoughlin, 1996).

Sexual Reproduction  81

Figure 2.8 Hermaphrodites: Fecundity as function of age in A. earthworm Lumbricus terrestris (for a period of 3 y, Lowe and Butt, 2005), B. in naidid Branchiura sowerbyi (Lobo and Alves, 2011), C. segment number in Enchytraeus variatus (Bouguenec and Giani, 1989), D. age and E. body weight in fast and slow growing morphs of Tubifex tubifex (Pasteris et al., 1996). Polychaetes: Fecundity as function of age in F. Ophryotrocha adherens (Paavo et al., 2002), G. digamic Dinophilus gyrociliatus fed on different diets (Prevedelli and Vandini, 1999), H. tube-dwelling, egg brooding planktotrophic and lecithotrophic morphs of Streblospio benedicti and I. self-fertilizing viviparous Neanthes limnicola (Fong and Pearse, 1992). All figures are modified and redrawn.

and age. Obviously, growth in body weight proceeds along with age in T. tubifex. But in others like S. spirorbis, age may significantly vary for a given body size. Some scientists (Britayev and Belov, 1994, Plyuscheva et al., 2004) have unsuccessfully attempted to correlate jaw length with age in polychaetes. In most hirudineans, a third complication is the dorso-ventrally flattened body to an almost rectangular shaped mid-body, in which a pair of ovary is accommodated. Contrastingly, the cylindrical body shape in all other annelids differs distinctly from that of the hirudineans. The flattened rectangular body provides 1.5–2.0 times larger surface area to nourish more number of eggs to undergo vitellogenesis than that of cylindrical body in typical annelids limiting the number of oocytes entering vitellogenesis. Of

82  Reproduction and Development in Annelida

course, there are exceptions to this generalization. For example, the body shape of continuously breeding polychaetes like Dinophilus gyrociliatus (Fig. 2.8G) and Ophryotrocha puerilis puerilis (Fig. 2.7D) tends to be rectangular and that of annually breeding hirudinean Nephelopsis obscura (Fig. 2.7E) is more of cylindrical than rectangular. Despite the diversity and complications, two major trends emerge for the fecundity body size relation. In the first one, the relation is positive and linear, and is observed in broadcasters and sanguivorous leeches. Irrespective of the morphological features (taken to represent body size) like body length in semelparous epitokous Nereis virens (Fig. 2.7A), body width in Hediste japonica (Fig. 2.7B), body weight in iteroparous Marphysa sanguinea (Fig. 2.7C) and number of chaetigers in the protandrous continuous breeders Ophryotrocha puerilis puerilis (Fig. 2.7D), the linear relationship remains, albeit at different levels. In the semelparous Perinereis cultrifera, the reported correlation per se is weak. About 26% of the variances of fecundity are influenced by body size and the remaining 74% by environmental factors like food supply (Rettob, 2012). However, BF decreases in the following order: semelparous epitokous spawners (e.g. Nereis virens, Fig. 2.7A) < iteroparous spawners (e.g. Marphysa sanguinea, Fig. 2.7C) < iteroparous brooders (e.g. Nicolea zostericola, Table 2.11). In them, the number of gametogenic segments also progressively decreases from almost all the abdominal segments in the epitokous spawners to a few in the iteroparous brooders. Apparently, semelparous species allocate majority of the available resources to gamete production, whereas iteroparous species invest the bulk of available resources on somatic growth (see Cassai and Prevedelli, 1998a). In broadcasters, the durations required to complete oogenesis are 10, 12 and 12 or 19 months in Glycera dibranchiata (Creaser, 1973), Neoamphitrite robusta (McHugh, 1993) and N. virens (Creaser and Clifford, 1982), respectively. With dorso-ventrally flattened body and ovaries in the sanguivorous leeches Hirudo medicinalis and Haementeria ghilianii, fecundity also increases linearly and positively with advancing age (Fig. 2.7F), increasing body weight (Fig. 2.7E) as well as the quantum of ingested blood (Fig. 2.7F, G). Apparently, more than age and body weight, the assured supply of nutritional source ensures increase in fecundity with increasing quantum of ingested blood. However, within a single reproductive season/bout, an inverted ‘U’-shaped trend for the fecundity-time relations has been reported for the glossiphoniid Helobdella californica (Kutschera, 1989) and Nephelopsis obscura, in which temperature regulates the spawning pulses (Peterson, 1983, Holmstrand and Collins, 1985). Notably, the trend is linear for the continuous spawner Ophryotrocha puerilis puerilis (Fig. 2.7D), in which eggs are spawned once a day. But it is an inverted ‘V’ trend in Dinophilus gyrociliatus (Fig. 2.8G), which spawns egg masses once a week. About 50% of polychaetes brood their eggs/egg sacs attaching them on suitable substratum (e.g. mucous nesting site in some sabellids, Gambi et al., 2001), on dorsal elytra (e.g. Harmothoe imbricata, Daly, 1972) or within

Sexual Reproduction  83

the branchial crown (e.g. Potamilla antarctica, Knight-Jones and Bowden, 1984). Encountering problems of safety and space, extra- or intra-tubular structures have been developed. Being cylindrical, these tubes also limit space with increasing body size. It must also be noted that only the number of brooded eggs are counted; however, none has reported the number of fertilized eggs that have been accommodated within the tube. In all the brooding polychaetes, an inverted ‘U’ or ‘V’ shaped trends is noted for the fecundity-body size relation. The trends are, however, linear and negative but are parallel with different levels in PLK and LEC morphs of Streblospio benedicti (Fig. 2.8H). Notable is the shorter lifespan of LEC morph. In viviparous Neanthes limnicola, no trend is apparent (Fig. 2.8I). Almost all the oligochaetes with cylindrical body also exhibit an inverted ‘V’ shaped trend, as in Enchytraeus variatus (Fig. 2.8C) or an inverted ‘U’ shape, as in Branchiura sowerbyi (Fig. 2.8B) and Tubifex tubifex (Fig. 2.8D, E). In the earthworm Lumbricus terrestris, fecundity also decreases linearly with advancing age (Fig. 2.8A). Arguably, almost all the oligochaetes and brooding polychaetes exhibit declining trends for fecundity with advancing age or increasing body size. In contrast, broadcast spawning polychaetes with more but variable number of gametogenic segments provide cumulatively a larger coelomic surface area to nourish more number of eggs to undergo vitellogenesis. Similarly, the flattened ovaries of the sanguivorous leeches provide relatively more surface area than that in the saccular ovaries of oligochaetes. Consequently, the broadcast spawning polychaetes and sanguivorous leeches do not limit the number of oocytes entering vitellogenesis. In them, fecundity increases with advancing age and/or increasing body size. However, it decreases per se in oligochaetes and brooding polychaetes, as their relatively smaller ovarian/ coelomic surface area in old worms limit the number of pre-vitellogenic oocytes entering vitellogenesis. In this context, the maintenance of Oogonial Stem Cells (OSCs) is relevant. The OSCs are responsible for the sustained production of oogonia (see Pandian, 2012). In an important publication, Daly (1978b) has shown that the number of pre-vitellogenic oocytes released from the ‘ovary’ for the ensuing brood is compensated by that entering vitellogenesis in the preceding brood of the serpulid Spirorbis spirorbis. This rare observation is likely to be confirmed in other annelids. Hence, it is likely that: 1. The compensatory release of oogonia indicates that the OSCs in annelids do not undergo ageing and senescence (however see Martinez and Levinton, 1992). 2. The senescence is rather imposed in oligochaetes and brooding polychaetes by the progressively decreasing ovarian/coelomic surface area in the old/ large worms by limiting the number of pre-vitellogenic oocytes entering vitellogenesis. As in annelids, broadcast spawning crustaceans also do not suffer reproductive senescence but the brooders suffer from it (Pandian, 2016).

84  Reproduction and Development in Annelida

2.6  Poecilogony and Dispersal Poecilogonics exhibit multiple-patterns of dichotomy in egg size/egg type and larval development. They alter the existence and efficiency of selection force for alternative and coexisting reproductive and developmental strategies (Fischer, 1999). It is a rare but interesting reproductive mode that occurs in half a dozen opsithobranch molluscs (Pandian, 2017). In polychaetes, it occurs in Hediste japonica (Sato, 1999) and in six spionid species: Streblospio benedicti (Levin and Huggertt, 1990), Boccardia acus, B. andrologyna, B. chilensis (Read, 1975), B. proboscidae (Gibson, 1997), B. semibranchiata (Guerin, 1991) and Pygospio elegans (Jenni, 2012). In H. japonica and S. benedicti, it includes planktotrophic (PLK) and lecithotrophic (LEC) morphs. The PLKs produce many small eggs that develop into feeding larvae with 2–3 weeks stay as plankton. But the LECs generate a few but larger eggs that develop into non-feeding larvae with or without planktonic phase. An advantage of the PLK larval development includes (i) greater fecundity, (ii) reduced parental investment, (iii) enhanced dispersal and gene flow, (iv) ability to colonize new habitats and (v) greater potential to delay metamorphosis to select a better habitat. On the other hand, the potential advantages of LEC larval development includes (i) higher larval survivorship, (ii) independence from external food supply, (iii) utilization of potential habitat and resources resulting in denser density and greater production (Levin and Huggett, 1990 see also Table 2.12). Available information on the cluster of life history traits of the poecilogonic polychaetes reveals dichotomy in S. benedicti and H. japonica but trichotomy in B. proboscidae (Table 2.13). Production of nurse eggs and adelpophagy occurs in B. proboscidae but not in S. benedicti and H. japonica. In the rivers of Aomori and Kagoshima (Japan), H. japonica PLK and LEC morphs co-occur but may not interbreed. The free spawning small sized (65 mm) PLK adult produces ten thousands to one million small (150 µm) eggs (Fig. 2.7B) and its neochaete larva with long chaete migrates to the sea and returns as juvenile. But the larger (73 mm) LEC adult spawns a few thousand eggs (230 µm) and its direct development is completed within the estuary (Sato, 1999). Electrophoretic studies have revealed the homogenous genetic structure in PLK morph but genetic differentiation in LEC morph. In the coast of North Carolina (USA), the brooding S. benedicti PLK and LEC morphs within a deme co-occur, interbreed and generate viable hybrids (Table 2.12). With available hybrids, the composition of the PLK within the deme consists of mostly PLK (64%) and LEC (10%) and hybrids (20%). Within the same deme also, it consists of predominantly 74% LEC but only 10% PLK and 16% hybrids. The PLK embryos seem to be more dependent

Sexual Reproduction  85

Table 2.12 Dichotomic poecilogonic developmental traits of Streblospio benedicti (compiled from Levin and Huggett, 1990, Levin and Bridges, 1994) Trait

PLK

LEC

Hybrid

Ova Size (µm)

65

152

74

Volume (µl × 10–3)

0.5

3.1

–

Carbon/embryo (µg)

0.11

0.85

–

Nitrogen/embryo (µg)

0.023

0.017

–

11

4

5.5

No./ovary

Larvae No./brood

200

40

80

No./pouch

10

3

5

Released chaetiger stage (no.)

3–5

9–12

–

Released larval length (µm)

250

600

–

Swimming setae

Present

Absent

Variable

Larval nutrition

Feeding

Non-feeding

Facultative

12–20

0–9

7–9

Larval duration (d)

Adults Life span (w)

38

30–75

–

9–10

13–14

–

6

6

–

PLK deme

64

10

20

LEC deme

10

74

16

Colonizing PLK (%)

82

20

–

Mean density (no./m2)

5,030

12,935

–

Production (g/m2)

2.57

3.68

–

Age at Ist spawning (w) Brood (no./life time) Composition (%)

on protein (0.023 µg N/embryo), whereas the LEC on carbohydrate (0.85 µg C/embryo). With relatively more number of broods and brood pouches, the PLKs release smaller (250 µm) feeding larvae at ~ 4 chaetiger stage lasting for 12–20 days, in comparison to the large (600 µm) non-feeding larvae released at 9–12th chaetiger stage with planktonic duration of 0–9 days. Though the colonizing ability of PLK is higher (82%), their density (5,030 no./m2) and production (2.6 g/m2) are lower than (12,935 no./m2, 3.7 g/m2) those of LEC. Briefly, the PLK is more an explorative morph, while LEC is more a productive morph in the favorable parental habitat. Reciprocal mating between PLK and LEC reveals a tendency for perpetuation of the dichotomy (Levin et al., 1991).

86  Reproduction and Development in Annelida

Table 2.13 Dichotomic poecilogonic developmental traits of Boccardia proboscidae (condensed from Gibson, 1997) Trait

Type 1

Type 2

Type 3

Egg size (µm)

94.5

92.7

108.8

Capsules/brood

41.7

46.8

42.9

Eggs/brood

2180

1637

2309

Larvae/capsule

52.4

35.0

53.8

Nurse eggs/capsule

0.6

6.7

34.4

Nurse egg/larva

0.01

0.014

8.7

6

6

11 Type 3A

Type 3B

Post-hatchling

PLK

PLK

PLK

Benthic

Larval size (µm)

206

251

209  

484

Adelpophagic

No

Some

Some  

Yes

Planktonic duration (d)

30

19

15  

0

Age at sexual maturity (mo)

4

4

4

3

Encapsulated period (d)

In contrast, the morphs of B. proboscidae are not recognizable as PLKs and LECs but are typed as Types 1, 2 and 3; Type 1 is characterized by the absence of or presence of a very few nurse eggs and is developed into planktotrophic dispersive larvae but Types 2 and 3 by the presence of abundant nurse eggs consumed by adelpophagous viable number of embryos (Table 2.13). In Types 2 and 3, the nurse eggs appear similar to the viable oocytes prior to cleavage but fail to cleave. Females (78%) dominate the population of Type 1, but they are much less abundant in Type 2 (6%) and Type 3 (16%). Type 1 females brood eggs in the absence or presence of 0.01 nurse egg/viable egg; the hatched offspring (200 µm) is planktotrophic for 30 days. Type 2 females are similar to that of Type 1 but for every viable larva, there is 0.14 nurse egg and the hatched (251 µm) larva is planktotrophic for 19 days only. Type 3 females are further subdivided into A and B groups. In Type 3A, adelpophagy may occur but its larva is planktonic for 15 days. In Type 3B, adelpophagy is most common and direct development results in the release of benthic juvenile. Remarkably, the poecilogonic patterns vary among broods produced by different females as well as within a single brood and even in an egg capsule. Type 1 and Type 3 adults can successfully be mated. All the offspring produce broods characterized by maternal breeding type, regardless of paternal origin. Apparently, its sex is determined by ZZ-ZW female heterogametic system. In Type 1, female produce no or very few nurse eggs and release 3-chaetiger larva on 6 days after spawning. Type 3 females spawn both viable and a large number of nurse eggs. In

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B. proboscidae, poecilogony represents a gradual shift from planktotrophy to adelpophagy.

2.7  Mating Systems Egg production is costlier than that of sperm (Charnov, 1982). Consequently, reproductive success of the female is limited by access to resources, whereas that of a male is limited by availability of females (Bateman, 1948). The sex allocation theory (Charnov, 1987) explains the differences in resource allocation to male versus female reproduction. Fishes have proved as an excellent animal system to test a number of hypotheses on sex allocation theory (Fischer and Petersen, 1987). For the following reasons, annelids, especially polychaetes as an ancient group can also serve as an excellent invertebrate model: 1. The presence of different forms of sexuality ranging from gonochorism to sequential, serial and simultaneous hermaphroditism (SH). 2. About 50% of polychaetes have free swimming larval stages(s), while their adults are sessile. Hence, they allow a study on the effects of a range of population structure and selection pressure on mating system. 3. The ease with which they can be reared in the laboratory as well as culture system (see Chapter 8) allows precise estimates on individual reproductive success (Premoli and Sella, 1995). Ophryotrocha sp collected from 1,500 m depth is readily amenable for culture in laboratory, where it has completed three successive generations within 7 years of the experimental study (Mercier et al., 2014). Annelids display the three recognized mating systems. However, monogamy is limited to very few (e.g. Nereis acuminata, Weinberg et al., 1990; enforced monogamy, Ophryotrocha puerilis puerilis, Berglund, 1986, O. diadema, Cannarsa et al., 2015). A microsatellite marker study has revealed the presence of a near monogamy in Homogaster elisae, in which all the four spermatheca stores sperm from the same male (see Diaz-Cosin et al., 2011). However, H. elisae individuals select partners with similar body size (Novo et al., 2010). In-breeding and out-breeding pairs of Eisenia andrei produce 30 and 19%, respectively fewer cocoons than intrapopulation mating pairs (Velando et al., 2006). From their observations, Dominguez et al. (2003) have reported that 88 and 10% of matings in E. foetida are reciprocal and unilateral, respectively. Body shape and locations of male and female pores minimize the scope for self-fertilization in earthworms, though rare such events are recorded in some species. For experimental study on mating systems, a few species belonging to Ophryotrocha have served as an ideal model. They are small marine polychaetes and amenable to rearing. In them, some are SH (e.g. O. diadema),

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others are sequentials (e.g. O. puerilis puerilis) and yet others are gonochoric (O. labronica). O. diadema commences its sexual life very early with a progametic male phase lasting for 21 days and sperms are produced from the 3rd and 4th segments in a worm with six segmented body. When body size reaches a length 15 chaetigers, it commences to generate oocytes from the last five segments onward. On becoming SH, the mating pairs regularly take turns to assume either the male or female role. Assuming the female role, it allows only a smaller number of oocytes within a parcel to mature at a time and limits the parcel to hold 29 eggs/cocoon and lay parcels more frequently during the 30–40 d-SH phase (Sella, 1988), in comparison to gonochores and sequential hermaphroditic Ophryotrocha species that lay hundred eggs/ cocoon.

2.7.1  Simultaneous Hermaphrodites As it is cheaper to produce sperms, male assuming SH tends to cheat the other partner. To guard against the non-reciprocating mating partners, three mechanisms have been developed: 1. The female acting worms can distinguish whether ‘her’ partner is an adolescent male on the verge of sex change or a hermaphrodite or an young/small male. The female-acting SH can regulate the clutch size by releasing more or less number of oocytes to mature and subsequently spawn. Accordingly, successive spawnings are regulated at intervals of 3.0, 5.2 and 5.4 days, with the partner being a young/small male, adolescent male and hermaphrodite, respectively (Sella, 1988). 2. The female acting SH parcels 29 eggs per cocoon (Akesson, 1973). This parceling strategy enables her to regulate the number of parcels, according to her mate being a young male, adolescent male or SH. 3. A mating partner from a bigger group invests more on sperm production to escape from male-male competition than a male acting partner from a small group. Hence, the chances for the male acting SH from a larger group to escape from reciprocation are less, in comparison to the male acting SH from a smaller group (Premoli and Sella, 1995). In another SH O. gracilis, mating occurs in pairs of ovigerous hermaphrodites, which sequentially alternate sex roles more than once. No male competition occurs. Protandrous males are not involved in pair formation. Hence reciprocal insemination is safeguarded (Sella et al., 1997). Female acting SH can assess the group size through a water-borne lipidic pheromone and adjust the reproductive output appropriately (Schleicherova et al., 2010). On the other hand, males can also regulate the quantum of sperm donation. For example, Eisenia andrei, an epigeic earthworm, detects the virgin status of the mating partner and appropriately adjusts the quantum of donated sperms (Velando et al., 2008). Not only sex allocation is adjusted by SH according to social conditions, but can also adjust the body size of sex change. In O. diadema, protandrous males from isolated and intermediate groups attain sexual maturity with significantly larger number

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of segments than those from a larger group. In O. diadema, the female acting monogamaous SH spawns more number of eggs than the SH exposed to promiscuity (Cannarsa et al., 2015). Characterized by strongly female-based allocation, the ovarian biomass of O. diadema, O. gracilis and O. hartmanni is ~ 80% of the gonadal biomass. Hence, the reciprocating hermaphrodites of Ophryotrocha produce eggs equivalent to 0.8 times of the eggs produced by a hypothetical mutant pure female. With reciprocation of equal amount by the partner, the reproductive success of SH shall be 0.8 × 2 = 1.6 times more than that of a pure female (Sella and Ramella, 1999).

2.7.2  Sequential Hermaphrodites The majority of sequential hermaphrodites are protandrics. Schroeder and Hermans (1975) recorded 32 out of 67 hermaphroditic species as protandrics. In them, male ratio decreases from 0.80 in young worm to 0.25 in older ones (e.g. Hydroides elegans, Qiu and Qian, 1997). The size advantage hypothesis of Ghiselin (1969) and others predicts that reproductive success will increase less with increasing body size for males than for females. In Ophryotrocha puerilis puerilis, an individual commences sperm production at the nineth segment size and changes sex, when it attains a body length with 19 chaetigers. From a series of experiments, in which the potential mating pairs consisted of all possible combinations of different body sizes of males and females of O. puerilis puerilis in mate choice experiment, Berglund (1990) has brought experimental proof for the size advantage hypothesis. He has observed that the pair consisting of a small male and a large female produces the highest daily egg output, a measure of reproductive success. In an another experiment, Berglund (1991) has assessed the fecundity in isolated malefemale pairs of protandric O. puerilis puerilis and gonochoric O. labronica and concluded that the former are not benefited from large body size. Hence, they will be better off by changing sex, as females prefer to mate with small males. However, the larger O. labronica males have greater access to more females. Hence, they gain nothing by changing sex. In O. puerilis puerilis, the time cost for sex change is only ~ 6 days. Interestingly, Monahan (1988) has shown that within 48 hours, individuals in male phase can commence oocyte production. Amazingly, the mating partners in a pair are able to simultaneously change sex several times in their life time (Premoli and Sella, 1995). Trypanosyllis zebra and Syllis amica are also reported to undergo this type of serial sex change (Policansky, 1982). Berglund (1986) has recorded that 4 out of 14 pairs in O. puerilis puerilis have become simultaneous hermaphrodites, after they have reproduced for a month by performing simultaneous sex change. Subsequently, these pairs require assuming an alternate male and female role alone. Indeed, that has saved much of the resource and time required to change sex from male to female or female to male. Berglund explained this phenomenon in the light

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of sex allocation theory. As eggs are costlier to produce than sperm, a female is quickly exhausted all of its resources. The sex changed to male phase provides adequate time to recover quickly and store adequate resource to act as female.

2.7.3  Labile Gonochorics In a few gonochorics, either the sex determination or differentiation process is labile. For example, the obligate need for the presence of male pheromone for the mature female to spawn is demonstrated in Brania clavata. In the absence of the male pheromones, the female resorbs the oocyte and changes sex to male (Hauenschild, 1953). The protandric Typosyllis prolifera is a stolonizing broadcast spawner with a planktonic larval phase and a sedentary adult phase. A small or larger portion of females in a population undergo irreversible change to male sex at one of its subsequent reproductive cycles, indicating the labile sex differentiation (Franke, 1986b). Consequently, resource allocation for reproduction is altered in these sex changing males. In Capitella sp I, the processes of sex determination and differentiation remain stable in heterogametic (ZW) female but labile in homogametic (ZZ) male. When the worm is reared in isolation or in a small group, some males become ZZ hermaphrodites and function as either sex. The development of hermaphroditism is regarded as an ‘emergency adaptation’ to low density. Not only in the absence of females but also excess food is required to induce hermaphroditism. Isolated males require only 22 days to develop ova. Interestingly, these hermaphrodites do not self-fertilize. But their ability to mate as a male or female depends on density and frequency of hermaphrodites. However, the hermaphrodite is unsuccessful as a male. On crossing with a normal ZZ male, the female-acting ZZ hermaphrodite produces only males (Petraitis, 1985a, b). Estimates on the fertility of these female-acting hermaphrodites range from 0.1 to 0.3, assuming that of a true female as one (Petraitis, 1988). On the other hand, the male heterogametic Dinophilus gyrociliatus is gonochoric, dimorphic and reproduces iteroparously. Females lay transparent egg capsules and in its life time; she may spawn a minimum of 40–50 eggs and a maximum of 120–130 eggs. In it, a single ovary simultaneously generates one small (XO) male egg of 40 µm size and many larger female eggs each measuring about 80 µm in a single batch. Consequently, a female egg (by its larger volume) receives ~ 8-times more resources than the male egg with its smaller volume. With sex ratio of 3 ♀ : 1 ♂, the resource allocation is ~ 24-times more to daughters, in comparison to the sons (Minetti et al., 2013). In D. gyrociliatus, the unique mechanism, that allows the mothers to overcome the chromosomal mechanism of sex determination, is the selective

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fertilization of larger eggs by X barring sperms and of small egg by sperms without sex chromosome. The dwarf male arising from a small egg possesses no digestive system, inoculates sperms into his many immature sisters and dies prior to hatching (Traut, 1969a, b, 1970). When reared under stressful temperature (Simonini and Prevedelli, 2003, Akesson and Costlow, 1991) and/or salinity (Akesson and Costlow, 1991) or diet (Prevedelli and Vandini, 1999), more number of small eggs are produced resulting in more number of dwarf males.

3 Regeneration

Introduction Regeneration is the potency to repair and replace the voluntary loss of cells, tissues and organs of an animal to escape from (i) sub-lethal predation, (ii) self-inflicted spontaneous autotomy and (iii) intolerable shock (e.g. encounter frequency in Diopatra aciculata, Safarik et al., 2006, hypo-osmotic stress in Marenzellaria viridis, David and Williams, 2016). In 1898, Morgan has classified regeneration into two types: 1. Morphallaxis involving the remodeling of existing tissues into missing ones without extensive cell proliferation (e.g. regeneration of mid-body segments in the oligochaete Lumbriculus variegatus, Martinez-Acosta and Zoran, 2015). 2. Epimorphosis involving massive proliferation of undifferentiated cells/stem cells and formation blastema (e.g. head and tail regeneration in L. variegatus). Morphallaxis may be less energy-intensive than epimorphosis, as it involves remodeling some pre-existing structure, whereas epimorphosis requires de novo formation of missing body parts from blastema. Like echinoderms (Pandian, 2018), annelids represent another phylum with exceptional prodigius potency for regeneration. The head of L. variegatus can be regenerated as many as 21 times and the tail 42 times and both together 20 times (Muller, 1908). However, unlike the radially symmetrical echinoderms, the bilaterally symmetrical body of annelid is composed of metameric segments, each consisting of the same organs and so on. Hence, amputation at any axial position along the body of annelids results primarily in the removal of different quantum of the same organ system rather than the removal of a different organ system with a unique structure of echinoderms. Once a small or larger body part is lost, the individual suffers from locomotor function and feeding activity (e.g. Dipolydora quadrilobata, Lindsay et al., 2007), growth (e.g. Eisenia foetida, Xiao et al., 2011) and longevity (e.g. D. commensalis, Dualan and Williams, 2011), reproduction (e.g. Polydora ligni, P. cornuta, Zajac, 1985, 1995) and behavior (e.g. Pseudopolydora kempi japonica, Lindsay and Woodin, 1995). Not

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surprisingly, only a few polychaete and oligochaete species (~ 250 species, Zattara and Bely, 2016) can regenerate the lost body parts. Further, the regenerative potency varies greatly from species that can regenerate every part of the body from even a single isolated segment (e.g. polychaetes: Chaetopterus variopedatus, Berrill, 1928; oligochaetes: L. variegatus, Morgulis, 1907) to those that cannot regenerate even a single lost segment (e.g. all hirudineans).

3.1  Regeneration and Reproduction Embryogenesis, regeneration and clonal reproduction are similar but not identical developmental processes. For example, wound healing and blastema formation are not part of embryogenesis (Myohara, 2004). No blastema is formed during clonal reproduction (see Martinez-Acosta and Zoran, 2015). In Amphipolydora vestalis, for example, morphogenetic regeneration requires only 8 days but that of embryogenesis as long as 22 days (Gibson and Harvey, 2000). Table 3.1 represents a comparative summary of major events in regeneration, and architomic and paratomic reproduction. The differences in many of these events are explained in the ensuing sections and Chapter 4, as well. On the other hand, regeneration and clonal reproduction share extensive similarities, especially in development of peripheral nervous system and thereby provides a strong support for the hypothesis that clonal reproduction is derived from regeneration (Zattara and Bely, 2011). From a more detailed analysis, Zattara and Bely (2016) have noted that of 87 species with potency for anterior regeneration, 32% of species alone are capable of clonal reproduction (however, see Table 3.8). But only 7% of (67) species with potency for posterior regeneration can reproduce clonally. Hence, clonal reproduction may have evolved from those with potency for anterior regeneration. Still, there are differences between these two processes. For example, four anterior segments of the naidid Paranais litoralis are generated in each clonal reproductive cycle but regeneration fails to produce these four anterior segments (Bely, 1999). In Lumbriculus variegatus, these two processes share common cellular and molecular (Lan 3-2 epitopic expression) mechanisms during temporal and spatial developmental events. However, the Lan 3-2 epitopic upregulation is confined to regenerative segments and occur prior to clonal reproduction (Martinez et al., 2005). Differences between the trajectory of regeneration and clonal reproduction occur throughout these developmental processes suggesting that the divergence has occurred all along the developmental course of these trajectories (Zattara and Bely, 2016).

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Table 3.1 Comparison of major events during regeneration and two types of clonal reproduction: architomy and paratomy (compiled from Kostyuchenko et al., 2016) Event

Regeneration

Architomy

Paratomy

Wound/Fission

Occurs before

Occurs after

Terminates the process

Wound healing

Commences after

Poorly apparent

Not apparent till end

Cell migration

Commences after

Commences after

Precedes at fission zone

Blastema formation

Occurs after cell proliferation

Precedes fission

Precedes but continues until segmentation

Segmentation

Hypomorphic. Accompanied by morphallaxis

Hypomorphic, morphallactic, old segments reorganized in new anteroposterior (A–P) axis in polychaetes (Boilly et al., 2017) and oligochaetes (Myohara, 2004)

Limited morphallactic old segments precede and persist through the process

Muscular system

Circular muscle fibers formed de nova

Changes in muscles facilitate fission

Changes in muscles facilitate fission of new fragments

Digestive system

Transient reorganization. Stomach formed by morphallaxis

Morphallactic reorganization of digestive system along A–P axis

Emergence of morphallactic digestive system at late stage of fission

Nervous system

Following rupture, new fibers grow out from old ventral nerve cord (VNC) to form cereberal commissures and VNC ganglia

New nerves, cerebral commissures and VNC ganglia developed after fission

New nerves, cereberal commissures and VNC ganglia are formed prior to fission. VNC is ruptured at fission (see also Muller, 2004)

Molecular changes

Commences immediately after an injury/amputation

Precedes fission from 1–7 day

Precedes since formation of fission zone and persists through growth and differentiation of new zooids

In the light of the definition for clonal reproduction, the term of regeneration may have to be extended. Clonal reproduction is bi- (or multi-)directional, when an animal (genet) divides to produce two or more fully functional progenies (ramets). But it is unidirectional, when only a single (half of the ramets) progeny is developed (Rychel and Swalla, 2009, see also Bely, 1999), as in enteropneusts and solitary ascidians (Pandian, 2018). In principle, reproduction, whether sexual (including self-fertilizing hermaphrodite involving a single parent) or asexual (clonal), is expected to generate more

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than one offspring. Hence, a fission or amputation that generates a single ramet, as a product of unidirectional cloning, can be considered more as regeneration than reproduction. For some reason, many authors/reviewers (e.g. Berrill, 1952, Zattara and Bely, 2011, 2016) have not taken this into consideration. This chapter includes annelid species that are characterized by unidirectional cloning as regeneration. The following examples may explain it. The siboglinid vestimentiferan polychaetes live in chitinous tubes and inhabit primarily in hydrothermal vents. They lack a mouth, an anus and a digestive tract but harbor chaemoautotrophic symbiotic bacteria in the trophosome and derive metabolic needs from the bacteria (see also Chapter 1.5). The body of Lamellibrachia satsuma is divisible into (i) tentacular, (ii) vestimental, (iii) trunk and (iv) ophisthosoma regions (Table 3.2). The vestimental region contains the brain, heart, excretory organs and reproductive systems. The longest trunk region harbors the trophosome with symbiotic bacteria and reproductive organs. The ophisthosoma is the only segmented region of the body. From the description and Fig. 1 of Miyamoto et al. (2014), the following inferences can be made: 1. Amputated into two, Table 3.2 Stages of regeneration in two series of experiments in Lamellibrachia satsuma (compiled from information reported by Miyamoto et al., 2014; freehand drawing of L. satsuma is included, ten = tentacular, ves = vestimental, tr = trunk and op = ophisthosoma regions) Stage

Post-amputation (d)

External Features

Internal Features

Amputation between anterior and posterior trunk 1

0–10

Wound healing

Trophosome evaginated

2

14

Wound sealed

Bacteriocytes beneath the healed epidermis

3

20

White/pink blastema formed

Mesodermal cells aggregated

4

30

No sign of segmentation

Septa formed

5

40

Blastema projection Continuous nerve cord and blood vessels formed

Ophisthosoma segmented

Amputation between anterior and posterior ophisthosoma Anterior

Posterior

80% of ophisthosoma Single row of setae

20% of ophisthosoma No setae

21 days after amputation, trophosomal structures appeared 40 days after amputation, regeneration completed

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the anterior fragment containing the tentacular, vestimental and an anterior part of the trunk regenerates the entire posterior region. 2. However, neither an anterior fragment containing only the tentacular and vestimental regions nor the posterior fragment containing the posterior fraction of the trunk and ophisthosoma is able to regenerate the missing body regions. The stages of regeneration in two series of the experiment are summarized in Table 3.2. Clearly, (i) neither the so called ‘anterior’ nor ‘posterior’ fragment of L. satsuma is able to regenerate, (ii) the stem cells responsible for regeneration remain in the mid-body, namely the anterior trunk and (iii) potency for regeneration limited to the mid-body and represents an example for regeneration of unidirectional cloning polychaetes. In oligochaetes, earthworms are a good model for the study of regeneration because of their availability and economic importance. However, their regeneration is complicated by the chloragogue, position and size of clitellum (Fig. 3.1) as well as aestivation/hibernation (e.g. aestivation in the tubificid Rhyacodrilus hiemalis, Narita, 2006, hibernation in Lumbricus terrestris, Liebmann, 1942). In earthworms, the amputed head and tail are regenerated by the mid-body; however, the head and tail do not have the potency to regenerate the mid-body. Amputation at different positions along the mid-body length of Eisenia foetida is followed by regeneration toward anterior or posterior direction in immature worms. In fact, the middle amputee containing 30–66 segments in immature E. foetida and midbody segments of Perionyx excavatus (Fig. 3.1) regenerate bidirectionally. But regeneration is limited to the head (six segments) alone in sexually mature E. foetida (Fig. 3.1). In sexually mature E. foetida and L. terrestris, regeneration completely ceases (Liebmann, 1942). As the amputated fragment dies but the amputee regenerates in many earthworms, it is arguably clear that amputation at any level results in regeneration and not in clonal reproduction. Understandably, despite low survival (see Xiao et al., 2011), regenerative potency for anterior regeneration is common among these investigated earthworms, as they emerge with the head first from the burrows. Notably, the potency for the head is retained in both immature and mature worms. Recently, S. Sudhakar and his team (pers. comm.) have found that any fragment consisting of six segments in the post-clitellar region of P. excavatus has the potency to clonally reproduce, a feature experimentally shown for the first time (Fig. 3.1). While neoblasts may have been retained in almost all the post-clitellar segments, the chloragogue to provide adequate nutrients for clonal reproduction may have to originate from the minimum of six segments. It is not clear whether neoblasts are also present in other earthworms and if so, the need for research is obvious to know the overriding role played by chloragogue on regeneration and clonal reproduction.

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Figure 3.1 Death (D) and regeneration (R) of different fragments following amputation at pre-clitellum, clitellum and post-clitellum positions in selected earthworms. Long vertical arrows indicate the position of amputation. Short horizontal arrows indicate the direction of regeneration. Rd = regenerated worm. Note the experimental clonal reproduction in Perionyx excavatus (compiled from different sources including Sudhakar et al., unpublished).

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3.2  Incidence and Prevalence Incidence: Among oligochaetes, terrestrial species emerge the head first from their burrow. In contrast, most aquatic oligochaetes live with their heads burrowed into substrate but their tail is extended up into the water (Martinez-Acosta and Zoran, 2015). Hence, it is likely that the former suffer more frequently from the loss of anterior head (e.g. earthworms) but the latter from the posterior tail tip. Due to the loss of brain ganglia and reproductive structures in the anterior region, regeneration of anterior region can be costlier than the posterior one (Zattara and Bely, 2013). Understandably, polychaete species belonging to Arenicolidae and Opheliidae are incapable of anterior regeneration (see Weidhase et al., 2014). Sub-lethal predation has been a selective force responsible for the sustenance of regeneration of missing body parts (Bely, 2010). Not surprisingly, many authors believe that “Almost all polychaetes can regenerate appendages such as palps, tentacles, cirri and parapodia, and most are capable of regenerating the posterior end of the body” (Pires et al., 2012). However, an intense search has revealed that there are reports for ~ 40 species only (Table 3.3). Description in these reports is also limited to palps, head, a few anterior and/or posterior segments including the tail. This is also true for the > 2,000 speciose aquatic oligochaetes (Table 1.3), as reports available on structural regeneration are limited to ~ 10 species (Table 3.3). On this aspect, more information is provided later. The need is obvious for reports in species with feeding tentacles. Incidentally, a few polychaetes, which are unable to regenerate one or another body part, undertake clonal reproduction, whose products namely the ramets regain them (e.g. Paranais litoralis, Bely, 1999). Prevalence: On prevalence of regenerated structures, limited information is available from field collected species. Prevalence for the palp(s) ranges from 7–8% in Rhynchospio glutaeus to 20–50% in Pygospio elegans (Table 3.4). The values for the tail range from 29% (head + tail) in Lumbriculus variegatus to as high as 90% in Arenicola marina. Notably, a larger fraction of population undergoes posterior regeneration than anterior regeneration (e.g. Euclymene oerstedi, Clavier, 1984). Clavier (1984) has also estimated that the biomass production from regeneration of E. oerstedi is in the range of 2 g/m2/y. Notably, there are more losses of the tail (45%) than the head (28%) in earthworm Perionyx excavatus. Yet, that the field collected 2.6 and 1.1% of the worms can regenerate the tail and head, respectively indicate that the worm, albeit in very smaller proportion, has retained the bidirectional potential to regenerate the tail and head (see also Fig. 3.1). Regeneration: Our understanding of regeneration of body parts is based mostly from experimental studies. In spionids, the deciduous palps serve as food gathering organs (Lindsay and Woodin, 1992). But they (each measuring

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Table 3.3 List of annelid species reported to undergo regeneration (compiled from many sources) Organ and segmental regeneration Nereididae:  Nereis virens, Perinereis nuntia, Platynereis dumerilii Phyllodocidae:  Eulalia spp Amphinomidae:  Eurythoe complanata Nephtyidae:  Nephtys caeca Dorvilleidae:  Dorvillea bermudensis Chaetopteridae:  Chaetopterus variopedatus, Spiochaetopterus oculatus Cirratulidae:  Cirratulus cirratus Capitellidae:  Capitella sp I Arenicolidae:  Abarenicola, Arenicola marina Maldanidae:  Clymenella torquata, Euclymene oerstedi Oweniidae:  Owenia fusiformis Spionidae:  Boccardia syria, Dipolydora commensalis, D. quadrilobata, Marenzellaria viridis, Pseudopolydora kempi japonica, Polydora caulleryi, P. ciliata, P. cornuta, P. flava, P. ligni, Pygospio elegans, Rhynchospio glutaeus, Scolelepis squamata, Spio benedicti, S. setosa Sabellidae:  Branchiomma luctuosum, B. nigromaculata, Myxicola aesthetica, Sabella pavonina, S. melanostigma, S. spallazanii Serpulidae:  Hydroides dianthus, Kirkegaardia Onuphidae:  Diopatra amboinensis, D. dexiognatha, D. neapolitana Lumbriculidae:  Lumbriculus variegatus Lumbricidae:  Allolobophora molleri, Eisenia foetida, Eophila Enchytraeidae:  Dero, Limnodrilus hoffmeisteri, Pristina leidyi, Paranais litoralis, Tubifex tubifex Hirudinidae:  Hirudo medicinalis

0.9 mm in length) are difficult to be removed in Dipolydora commensalis, which receives food particles from its host hermit crab, on which it lives within the shell. On being reared in isolation from its host, the palp grows to a length 1.4 mm to enlarge the food gathering structure with cirri (Dualan and Williams, 2011). Ablation at different positions of the gill-bearing anterior part of the tubiculous onuphid Diopatra neapolitana indicates that the ablated first few (15) segments die and are not regenerated from the anterior. However, the ablation of anterior fragments from the 10th and those from the 25th to 45/55th chaetigers (except that of 20 chaetigers) are all regenerated from the cut ends of the remaining anterior zone (Fig. 3.2A). Notably, the ablated/lost anterior segments are regenerated by the remaining anterior segments. However, the posterior is not able to regenerate the anterior (Pires et al., 2012). The regenerative growth of the errant Eurythoe complanata is estimated as 0.14 segments per day. And, it is completed within 105 and

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270 days for the 16-segmented anterior and 37-segmented posterior, respectively (Kudenov, 1974). In polychaetes, segmental regeneration ranges from simple extension of the last abdominal segment by Arenicola marina (De Vlas, 1979) to complete formation of segments, as in many species. The regenerative process in Pygospio elegans, for example, proceeds as follows: on the 3rd day after ablation, the wound is sealed and blastema is formed, on the 6th day segments without setae are formed, on the 9th d prostomium, nuchal organs, mouth, short palps and setae are formed and on the 16th day the head is formed, chaetiger number is increased and palp length is increased (Lindsay et al., 2007). However, survival and normal growth of polychaetes decrease with increasing number of chaetigers ablated. For example, ablation of > 30% chaetigers is followed by 25% mortality and 51% abnormal regeneration in Marenzellaria viridis. But that of < 20% chaetigers results in 13% mortality and 9% abnormal regeneration (Whitford and Williams, 2016). In Polydora caulleryi too, only 10 chaetigers are regenerated, when ablation exceeds 14 chaetigers. A climax seems to be Spio setosa (83–126 chaetigers), in which regeneration ceases, when > 32 chaetigers are ablated. The response of Autolytus pictus is a shade different. In response to ablation of 1 to 5 and 5 to 13 segments, a head + 3 or 4 chaetigerous segments are regenerated but for that of 13 to 42 segments, a head alone is regenerated and for that involving > 42 segments, only a stump is developed (Okada, 1929). However, amputation, a half of the frontal segment (a half of prostomium and another half of prostomium in Ophryotrocha notoglandulata is followed by an increase of 0.7 (female) and 1.0 (male) segment (Pfannenstiel, 1974). Whereas only 14 anterior segments are regenerated in the field, the number of regenerated segments in Eurythoe complanata progressively decreases from 40–24 for the loss of ~ 80 segments. Notably, the anterior fragment regenerates a constant number of 14 segments in this worm with 130 segments (Kudenov, 1974). The oligochaetes also display similar reductions with increasing ablated number of segments. In Lumbriculus variegatus, ablation upto eight segments virtually from any position of the body are all replaced. However, the maximum of only eight segments are replaced, when ablation exceeds eight segments (Von Haffner, 1928). In Eisenia foetida too, regeneration ceases with ablation of 20 segments, although that for four to eight segments, three or four segments are regenerated (Moment, 1950). Hence, the degree of anterior regeneration has an upper limit, as in M. viridis and is reduced with increasing number of segments lost, as in Eisenia foetida (Fig. 3.2B). Conversely, the speed of regeneration is accelerated with increasing number of segments lost, especially in gill-bearing chaetigers, as in Pygospio elegans (Fig. 3.2C). An important factor that controls regenerative potency is the cerebral hormone in errant polychaetes. In Nereis diversicolor, sexual maturation stages are assessed by the oocyte size and their synchronous development.

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Figure 3.2 A. Survival of Diopatra neapolitana following amputation at different positions in the anterior region. The anterior amputee dies following ablation. The remaining anterior fragment survives and regenerates the missing parts, but the posterior is unable to regenerate the anterior (modified and redrawn from Pires et al., 2012). B. The number of segments regenerated as a function of the number of ablated segments in Marenzellaria viridis (drawn using data reported by Whitford and Williams, 2016) and Eisenia foetida (drawn using data reported by Moment, 1950). C. Number of regenerated segments as a function of time in Pygospio elegans, in which 5, 18 (50% gill bearing-) and 33 (100% gill bearing-) segments have been ablated (simplified and redrawn from Lindsay et al., 2007).

Figure 3.3 A. Effect of declining juvenile hormone level on oocyte size and number of regenerated segments in Nereis diversicolor S = submature, M = mature, P = postspawing, C = control (compiled, modified and redrawn from Golding, 1983). B. Schematic representation of declining level of ootrophic hormone in females and males and its effect on first and second stolonization in mature Typosyllis pulchra (modified schematic representation from Heacox and Schroeder, 1982).

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Females with occyte measuring < 100 µm, those with asynchronous population of oocytes measuring ~ 180 µm and those with synchronously developing oocytes of > 200 µm are considered as immature, submature and mature, respectively. Golding (1983) has estimated the number of regenerated segments in a 3 weeks period after amputation of 30 segments in decerebrated immature worms implanted with the ‘brain’ from a matured one. Figure 3.3A shows progressively decreasing number of regenerated segments with increasing oocyte size, a marker of sexual maturity. The decerebrated recipient control without the ‘brain’ implantation has almost lost the regenerative potency. The figure also indicates the reported decreasing trend for the cerebral Juvenile Hormone (JH) as a function of oocyte size (see Chapter 7). In the errant syllids, which reproduce through stolon, regeneration is also eliminated with the commencement of stolonization (e.g. Typosyllis pulchra, Heacox and Schroeder, 1982). Figure 3.3B also shows the reported declining trend for Ootrophic Hormone (OH) (see Chapter 7) with the commencement of stolonization. With the progressive decrease in JH/ OH, reproduction is commenced but regeneration is ceased. Understandably, these two processes competing for resource allocation mutually eliminate each other. However, sedentary polychaetes, which are mostly suspension feeder with continuous food supply, may undertake regeneration even after sexual maturity. Incidentally, regeneration in errant polychaetes depends on cerebral neurosecretion, whereas the sedentary polychaetes on posterior segmental ganglia (see p 111). Polychaete vs oligochaete reproduction: Limited information is available for the loss and regeneration of body parts on growth and/or reproduction. Nevertheless, the available information clearly indicates the adoption of contrasting strategies by oligochaetes with the chloragogue and sedentary polychaetes without it to meet the cost of regeneration. Whereas the former mutually eliminates regenerative and reproductive processes, the latter simultaneously undertake both of them but at a cost of reproductive output. In oligochaetes, the chloragogue functions as an organ equivalent to the liver in vertebrates. Derived from the peritoneum, it is located around the intestine and serves as the major center for synthesis and storage of glycogen and fats. It is colored due to the inclusion of green-yellow colored lipids. On release into the coelom, the chloragocytes become eleocytes (Barnes, 1974). Abundance of the chloragocytes and eleocytes depends on the amount of food consumed. The processes of regeneration and reproduction require eleocytes for their functioning. In E. foetida, the occurrence of one of these processes retards or arrests the other (Table 3.5), as the quantum of eleocytes is evidently not adequate to meet the simultaneous costs of these two resource-demanding processes (Liebmann, 1942). This is also true of Lumbricus terrestris and Pheretima (indica ?) and Tubifex tubifex (Liebmann, 1942). In the naidid Stylaria lacustris, Harper (1904) reports the mutual elimination between regeneration on one hand, and sexual and asexual reproduction on the other.

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Table 3.4 Prevalence of the loss of body parts in some annelids Species, Reference

Reported Observations Sedentary polychaetes

Arenicola marina (de Vlas, 1979)

90% worms have lost the tail; regeneration is limited to extension of the last segment but no new segment is added

Rhynchospio glutaeus, Pseudopolydora kempi japonica (Lindsay and Woodin, 1992)

7–8 and 10% of worms have lost I and II palps, respectively

Pygospio elegans, Polydora cornuta (Woodin, 1982, Miller and Jumars, 1986)

50–20% worms have lost palps

Marenzellaria viridis (Whitford and Williams, 2016)

7% worms have lost and are regenerating anterior segments

Euclymene oerstedi (Clavier, 1984)

8 and 44% worms are regenerating anterior and posterior segments, respectively

Clymenella torquata (Sayles, 1936)

0.1 and 3.9% of worms are regenerating anterior and posterior segments, respectively Errant polychaetes

Eurythoe complanata (Kudenov, 1974)

More headless fragments than tailless ones. 35–80% (mean ~ 50%) regenerating fragments have been recorded Oligochaetes

Lumbriculus variegatus (Morgulis, 1907)

Perionyx excavatus (Gates, 1927)

Loss of palps + head, head + tail and posterior segments is 19, 29 and 41%, respectively Of 533 worms in 4 collections, 77% have lost tail or head and regenerating it. Of them, 45 and 28% of the worms have lost tail and head, respectively. About 2.6 and 1.1% of the worms are regenerating tail and head, respectively

Clonal reproduction and regeneration intensely compete with sexual reproduction for resource allocation. Not surprisingly, both these reproductive modes are temporally separated. In most asexually reproducing annelids, clonal reproduction occurs prior to sexual maturity but sexual reproduction after sexual maturity. However, these two modes may also alternate, as in the sedentary Polydorella spp (Radashevsky, 1996). In other sedentary polychaetes like the spionid Amphipolydora vestalis, architomic fragmentation is the dominant form of reproduction but occurs concurrently with sexual reproduction. About 46% of asexual propagules or ramets contain gametes in the parental abdominal chaetigers (Gibson and Paterson, 2003). The other ramets gain the germline by the migrating piwi-positive ventral cells through the ventral nerve cord (Ozpolat and Bely, 2015).

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Table 3.5 Effect of regeneration of body parts on reproduction in some annelids Species, Reference

Reported Observations

Regeneration and reproduction: Sedentary polychaetes Polydora ligni (Zajac, 1985)

Palp regeneration in 8% worms reduces fecundity by 10–29%. Regeneration of 71% posterior segment reduces 49–80% fecundity

Polydora cornuta (Zajac, 1995)

In 30-, 60- and 70-segmented worm, regeneration of gametic segments is 13.8, 23.4 and 17.2% of the worms. Consequent reduction in reproductive performance is 27, 32 and 28% for gametogenic segments, egg capsules and cumulative fecundity, respectively

Regeneration and reproduction: Oligochaetes Eisenia foetida, Lumbricus terrestris, Pheretima (indica?), Tubifex tubifex (Liebmann, 1942)

Regeneration, asexual and sexual reproduction processes are mutually excluded, as they depend on reserve tissue, chloragogue. Regeneration of head is not affected by reproduction. During hibernation, not even posterior regeneration occurs, as eleocytes/ chloragogue are exhausted

Stylaria lacustris (Harper, 1904)

Regeneration, asexual and sexual reproduction processes are mutually excluded

Enchytraeus variatus (Bouguenec and Giani, 1989)

Simultaneously occur for a brief period at the cost of cocoon production

In general, oral structures like palps are more vulnerable in epifaunal polychaetes than the infaunal ones. Fecundity may be decreased with a loss of one or two oral palps, which are important oral structures for food collection and thereby decrease fecundity. But the loss of gametogenic or posterior segments profoundly reduces fecundity. Zajac (1985, 1995) has provided valuable information to show that regeneration of palps and segments occurs at the cost of reproduction. From Fig. 3.4, the following may be inferred: 1. The loss of one or two palps, followed their regeneration within 15 days, does not affect the number of gametogenic segments. 2. However, fecundity is significantly reduced, when gametogenic segments or posterior segments are lost. The loss of fecundity in the sedentary Polydora ligni ranges between 10 and 29% for palp(s) regeneration as well as 49 and 80% for regeneration of posterior segments (Table 3.5). In P. cornuta, the reduction in reproductive performance is 27, 32 and 28% for the loss of gametogenic segments, egg capsule and cumulative fecundity, respectively. Incidentally, from both field and experimental observations, Zajac (1986) has also reported that not only regeneration but also limited food availability and increased intra-specific density also slows growth, postpones sexual maturity, prolongs inter-brood and reduces fecundity.

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Figure 3.4 Effect of loss of one and two palps, as well as gametogenic segments (G segments) or posterior segments (P segments) on fecundity of the spionid polychaete Polydora ligni (compiled and simplified from Zajac, 1985).

3.3  Regenerative Process In annelids, regeneration has been studied for well over a century and in an array of representative species across the phylum (e.g. Randolph, 1892, Hyman, 1940, Hill, 1970, Bely, 2006). Earlier studies have focused on cellular and tissue-level dynamics of regeneration. They have laid the foundation for the present molecular level studies. This account has also consulted Bely’s (2014) review, which has considered a range of publications on histological and descriptive studies, experimental surgeries and graftings, and proliferation assays. The collated information is sequenced under the following headings: 1. wound-healing, 2. blastema formation and differentiation, 3. segmental reorganization, 4. growth and elongation (Weidhase et al., 2014). Briefly, regeneration is a fairly complex but an orderly process. In annelids, it ranges from species that are completely incapable of any regeneration, as in leeches and species that has no ability to regenerate even a single anterior segment,

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as in Streblospio benedicti (Lindsay et al., 2007) to species that are capable of regenerating an entire animal from a single mid-body (14th) segment in Chaetopterus variopedatus (Berrill, 1928). Not surprisingly, it is a highly variable trait.

3.3.1 Wound-healing In annelids, wound-healing includes (i) muscle contraction, (ii) autolysis, (iii) cell migration and (iv) re-epithelialization. Immediately following an amputation, most annelids seal the wound effectively by rapid longitudinal muscular contraction (e.g. Hyman, 1940, see also Samuel et al., 2012), stemming the loss of body fluids and possibly preventing infection. The acanthobdellid leeches are exception to it. With the presence of an outer stiff cuticle, they fail to seal the wound (Bely, 2006). Other leeches, however, respond to surgical lesion with a same sequence of events, as in vertebrates. In Hirudo medicinalis, for example, the newly synthesized collagen granulates the tissue and remodels the scar tissue (Tettamandi et al., 2005). In other annelids, wound-healing triggers early regenerative signaling events (King and Newmark, 2012). In Paranais frici, P. litoralis, Chaetogaster diaphanus, C. diastrophus as well as Amphichaeta raptisae B and C, the wound healing presumably fails to trigger the signaling. Hence, the sealed wound neither forms a blastema nor replaces any missing structures (Bely and Sikes, 2010). In Enchytraeus japonensis (Kawamoto et al., 2005) and Lumbriculus variegatus (Lesuik and Drewes, 1999), muscular contraction precedes autotomy. Within hours of an inflicted injury, autolysis is evident both in ectoderm and mesoderm (Cornec et al., 1987). Migration of an array of cells to the wound site is a general feature of regenerating annelids. The migrant cells phagocytize the damaged cells and tissue debris and form a tissue plug, while others serve as a source for the formation of new cells (Bely, 2014). The phagocytes are of different types and are called coelomocytes and splanchnopleural cells in Limnodrilus hoffmeisteri, chloragocytes/eleocytes and amoebocytes type I in L. variegatus as well as hyalocytes and macrophages in Eisenia foetida. Usually, the severed epidermal edges fuse with each other and the same occurs between the gut epithelia, as well (e.g. Hill, 1970). Hence, re-epithelialization is completed prior to cell proliferation at the wound.

3.3.2  Cell Migration An injury on any part of the body elicits an extensive migration of different cell types toward the wound. The origin of new cells and tissues is one of the fundamental problems in regeneration of annelids. Three hypotheses have been postulated concerning the origin of new tissue (Paulus and Muller, 2006). They are: 1. the epidermis proliferate new tissues including new epidermis or mesoderm (e.g. Stone, 1933), 2. the new tissue is generated by specific pluripotent mesodermal cells (e.g. Randolph, 1892) and 3. each

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germ layer retains its identity and proliferates material for the respective tissues (Hill, 1970). In a shade different from these hypotheses, Goss (1974) has postulated that the blastema is a mixture of differentiated cells and stem cells that migrate to the wound site. In a seminal publication, Randolph (1891) has discovered the ‘quiescent specialized embryonic cells called “neoblasts” from the posterior surface of the septa adjacent to the ventral nerve cord of Lumbriculus variegatus. Subsequently, her discovery is extended not only to some oligochaetes like Tubifex tubifex (Stone, 1932), Nais (Herlant-Meewis, 1947) and Limnodrilus hoffmeisteri (Cornec et al., 1987) but also to polychaetes like Aricia, Chaetopterus and Diopatra (see Berrill, 1952). Counting the number of neoblasts following posterior transection in the naidid Ophidonais serpentina, Bilello and Postwald (1974) have brought evidence for migration of neoblasts toward the wound site. Activated by an injury, these cells do migrate to the wound along the ventral nerve cord in oligochaetes (e.g. Enchytraeus japonensis, Tadokoro et al., 2006). Incidentally, the neoblasts of annelids are multipotent stem cells. Because of their pluripotency, the term neoblasts are more appropriate to planarians than to annelids (Randolph, 1892, Paulus and Muller, 2006). However, the term neoblasts (in multipotent sensu) here is continued to be used. Besides the mesodermal neoblasts, the injury-activated dedifferentiated stem cells are considered to be present in all the three germ layers (Jamieson, 1981). From their BrdU pulse-chase experiment in the amputated (at the 7th and others at every five segment up to the pygidium) Parougia bermudensis, Paulus and Muller (2006) have inferred the following: 1. Sealing of the wound is a pre-requisite for regeneration (see also Kato et al., 1999). 2. Following amputation, the amputees lose the gut contents (cf Subramanian et al., 2017). The everted gut epithelium forms an outer layer and closes the wound at both ends. The muscle fibers adjacent to the wound contract and constrict the tissue around the wound to ~ 20% of its size within 2 hours post-operation. Within a day, the entire wound is covered by the epithelium. 3. The BrdUlabeled cells appear 16 hours post-operation. Incidentally, this observation also confirms that of Hill (1970), who has reported that tritiated thymidine [3H] T-labeled cells of Nereis sp, Nephtys sp and Sabella melanostigma appear at the wound site 24 hours post-operation. 4. The delayed appearance of the epidermal stem cells indicates the time required for dedifferentiation of already existing epidermal cells. 5. The descendants of epidermal cells proliferate centrally but those of the gut peripherally and 6. The anterior region of the blastema transforms into the head, while the posterior forms the pygidium and persists as proliferation zones. 7a. Proliferation of the BrdU-labeled epidermal cells fills ~ 67% of the blastema within 250 hours of regenerative period. 7b. These BrdU-labeled epidermal cells appear in nearly almost all the areas of the body. 7c. But most of them are detectable at the posterior end and their number gradually decreases anteriorly. 7d. More

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importantly, the proliferation ceases earlier in the ‘older’ anterior segments than in the ‘young’ posterior segments. Incidentally, this may explain why less number of species regenerates anterior. Briefly, the interesting investigation by Paulus and Muller suggests that activated by the injury, the epidermal cells are dedifferentiated to become ectodermal stem cells. Incidentally, employing histochemical techniques, dedifferentiated mesodermal cells are shown to arise from the muscle cell tissues in the syllid Autolytus pictus (Okada, 1929) and Owenia (Probst, 1931). In this context, Prof. S. Sudhakar and his team at Manonmaniam Sundaranar University, Thirunelveli, India has brought a series of publications in recent years. Samuel et al. (2012) have traced the origin of mesodermal stem cells and blastema formation in Eudrilus eugeniae. The earthworm was amputated between the 11th and 12th segments passing through the clitellum. The wound was sealed 24–48 hours after amputation and soft transparent blastema was formed between 48 and 72 hours following amputation. A section through the worm 48 hours after amputation indicated reduction in the thickness of circular muscle layer from 170 to 98 µm, and an increase in the longitudinal layer from 100 to 188 µm, when the blastema was being formed. BrdU-labeling retention assay in the blastema revealed the presence of the BrdU-positive longitudinal cells in the blastema, presumably arising from the muscles. The immunochemical-BrdU antibody staining of the 2-day old blastema also confirmed the presence of BrdU-positive cells in the blastema. In another interesting study, Kalidas et al. (2015) demonstrated that Lamin A, an intermediate filament protein, is absent in the BrdU-positive stem cells of E. eugeniae. In the blastema of E. eugeniae, the cytoplasm but not the nuclei of the BrdU-positive cells is autofluorescent (Fig. 3.5A) due to the accumulation of riboflavin. In the amputated worm, functional mouth and anus are formed 6 days after amputation. The adaptive starvation during the first 5 days Table 3.6 Riboflavin content (µg/g) in different organs in intact and regenerating Eudrilus eugeniae. Right panel A and B shows the milky white and yellow riboflavin synthesizing Bacillus endophyticus, respectively (modified from Subramanian et al., 2017) Organ

Intact Worm

Regeneration on 3rd day

6th day

Intestine

305

966

321

CML + LCL

203

640

277

Coelomic fluid

368

386

415

Prostate gland

395

279

216

Setae

284

112

103

CML = Circular muscle layer, LCL = Longitudinal cell layer

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Figure 3.5 Eudrilus eugeniae: Upper panel shows A. BrdU-positive cells marked by arrows and B. shows autofluorescence of BrdU-positive cells under red filter. Lower panel showing regeneration on 3rd, 5th, and 7th d in intact worm (lower panel I), riboflavin supplemented worm (lower panel II) and antibiotic injected worm (lower panel III). Note the transparent regenerative blastema in these worms (modified from protocol record of Dr. S.C.J.R. Samuel).

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triggers the gut microflora to rapidly multiply and synthesize riboflavin. The presence of an efficient riboflavin synthesizing Bacillus endophyticus has been confirmed by 16Sr RNA sequencing. The microbial synthesis of riboflavin has also been confirmed as the colony of B. endophyticus changes its color from milky white to yellow on the second day. Table 3.6 shows that effectively employing the symbiotic microflora, E. eugeniae draws riboflavin for its regeneration between the 3rd and 5th day following the amputation. The intestine is the major location, in which much of riboflavin is synthesized by the riboflavin synthesizing symbiotic microbes. Other organs like prostate gland and setae do neither harbor the symbiotic microbes nor synthesize riboflavin. The presence of riboflavin in the circular muscle layer and longitudinal cell layer clearly indicates the stem cell rich tissues have abundant riboflavin in these layers to augment regeneration (Subramanian et al., 2017). The riboflavin supplementation accelerates blastemal length (Fig. 3.5C), a marker for the blastemal growth and possibly differentiation. But its suppression by injection of antibiotic reduces the blastemal growth (Samuel et al., 2012). Yet it remains to be known whether the mesodermal stem cells differentiate into mesoderm and its derivatives alone or they can also differentiate into epidermal and gut cells. Thanks are due to the Japanese scientists for establishing the source and sequence of mesodermal regeneration in Enchytraeus japonensis, a small (10–15 mm), almost transparent worm with the life span of < 2 weeks (Yoshida-Noro and Tochinai, 2010). Analyzing ~ 400 clones, Takeo et al. (2010) have isolated five genes, whose expression level is altered during the regenerative process. RT-PCR studies have shown that three of them Ejpsmd, EjTuba and grimp are upregulated, while the two others horu and minor are downregulated. Of the three, grimp is transiently but specifically expressed only in mesodermal cells just beneath epidermis at the blastema tip and entire amputees from 3 to 12 hours. But it is never expressed in the epidermis and digestive tract. The suppression of its expression inhibits mesodermal proliferation and anterior differentiation. Cross section analysis has shown that grimp is expressed in the mesodermal cells and neoblasts, characterized by large nuclei and nucleolei, and a large nucleo-cytoplasmic ratio. It must, however, be noted that Myohara (2012) has shown that the neoblasts are dispensable for regeneration in annelids. The annelid ‘neoblasts’ are peritoneal stem cells and may not be equal to those of planarians (Bilello and Postwald, 1974). Hence, activated by the injury, the dedifferentiated mesodermal stem cells are responsible for regeneration of mesoderm and its derivatives. Regarding regeneration of endoderm and its derivatives, there are only passing remarks (e.g. Paulus and Muller, 2006, Takeo et al., 2008). Briefly, available information lends support to Hill’s (1970) hypothesis that each germ layer has retained its identity. Activated by an injury, the differentiated stem cells from the respective germ layers regenerate their respective organs

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and systems. This view is further supported by the fact that annelids undergo mosaic development. In Tubifex rivulorum, blastomere deletion at four cell stage has revealed the loss of totipotency by all the three blastomeres (A, B, C) except the D blastomere (Penners, 1922). Subsequent to the formation of germ layers in spiralian embryos of annelids, each of the differentiated germ layer retains its identity but has already lost the potency to revert back (Needham, 1990).

3.3.3  Blastema and Differentiation Shortly after wound healing, a blastema is developed with the contribution from all three germ layers (e.g. ectoderm: Paulus and Muller, 2006, mesoderm: Randolph, 1892, endoderm: Takeo et al., 2008). Cellular contribution to the blastemal tissue arises from dedifferentiated cells beneath the wound site (Balavione, 2014). Differentiation: A large number of publications (Clark and Bonney, 1960, Clark et al., 1962, Hauenschild and Fischer, 1962, Nayar, 1966) have experimentally shown that posterior regeneration is induced by neurosecretion arising by the suboesophageal ganglia, the brain in many errant polychaetes like Nereis diversicolor, Platynereis dumerilii and Nephtys. Recently, Muller et al. (2003) have also found that the cephalic ganglia are essential for regeneration in the errant amphinomid Eurythoe complanata. However, the ventral ganglia in each posterior segment of the sedentary polychaetes are capable of posterior/caudal regeneration in the complete absence of a brain (e.g. Branchiomma nigromaculata, Chaetopterus variopedatus, Hill, 1972). Correlated with the reduction in sense organs, the brain of sedentary tube-dwelling polychaetes is quite simple, in comparison to that of the errant polychaetes (Bullock and Horridge, 1965). As the projected head of the sedentary worms is often subjected to sublethal predation, it may be advantageous for them to have the neurosecretory ability distributed along the nerve cord rather than localized in the cephalic ganglia. Not surprisingly, barring Arenicola marina, all the investigated sedentary tubiculous worms are capable of regeneration. Investigations on the Antero-Posterior Axis (APA) of the errant polychaete Nereis latescens have found that the Ventral Nerve Cord (VNC) assigns the APA. In the absence of VNC, the regenerate fails to develop APA especially the parapodia. But in the sedentary sabellids Dasychone infrata and Branchiomma nigromaculata, parapodial inversion occurs along the APA in the absence of VNC and it is correlated with the position of nerve cord (Boilly et al., 2017, see also Table 3.1). In the blastema, the system induces self-proliferation initially and regeneration subsequently. In Eurythoe complanata, extirpation of a single ganglion retards regeneration and that from more ganglia inhibits regeneration completely, unless the affected regenerate is autotomized (Muller et al., 2003). Understandably, the nervous system develops remarkably earlier than

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Figure 3.6 Self assemblage of Eudrilus eugeniae, when kept in water. A. Intact worm, B. Worm in which the brain harboring cephalic region up to 4th segment has been amputated, C. Worm, in which regions beyond 70th segment has been amputated (modified from protocol record of Dr. G. Daisy).

musculature in Cirratulus cf cirrus (Weidhase et al., 2014). Using phalloidinlabeling, anti-body staining and confocal laser scanning microscopy, Weidhase et al. (2014) have described the differentiation processes of nerves and musculature in the blastema of C. cirrus. In blastema, the nervous system is a typically a rope-ladder like arrangement and circumpharyngeal connectives exhibit two separate roots leading to the brain. As a general pattern of annelids, the regeneration of circular and longitudinal musculature originates from different group of cells. During regeneration, the longitudinal musculature commences with diffuse growth and subsequent structuring into the blastema. In contrast, circular musculature develops independently inside the blastema. In many oligochaetes, self assemblage is a behavioral response to stress conditions. For example, an exposure to hypoxic water (1.5 mg O2/l), the self assemblage of Tubifex tubifex forms a ball enabling the harvest of the worm in culture system (Marian and Pandian, 1984). In earthworm Eudrilus eugeniae, stressed by immersion into water, Daisy et al. (2016) have observed that the self assemblage behavior remarkably changes in the presence and absence of the brain. Even in the absence of the brain, the worm, in which the first four segments with brain are amputed can survive but is unable to self assemblage (Fig. 3.6B) and requires the intact brain for the selfassemblage within an area of 2.3 cm2 per worm. However, in the presence of the brain, the worm amputated at the 70th segment is able to self assemblage (Fig. 3.6C) and requires an area of just 0.9 cm2 per worm.

3.3.4  Segmentation and Reorganization In annelids, the number of segment ranges from a few (e.g. ~ eight segments, 400 µm long Apodotrocha progenerans, see Westheide, 1984) to as many as 300–700 in Eunice siciliensis (Hofmann, 1974). The segments are almost uniform throughout the body. Yet, based on the parapodial structure and

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function, they can be grouped into cephalic, trunk and caudal regions. In annelids, segments are added successively forward from the growth zone of the structurally well developed pygidium, although the number is limited in a few exceptional polychaetes (e.g. Clymenella torquata, Sayles, 1932) and oligochaetes (e.g. Eisenia foetida, Moment, 1946, 1950). Following amputation at the position of 50th or 80th segment in E. foetida, new segments are added posteriorly by regeneration until the original total numbers of 100 segments are restored. Considering only the publications by Moment, Berrill (1952) has wrongly concluded that in all oligochaetes, the addition of segments is ceased early and is precociously attained. In oligochaetes, the segment number varies widely from 80 to 100 in Eudrilus eugeniae (shodhganga.inflibnet. ac.in), 135 to 150 in Lumbricus terrestris (The Robinson Library) and 145 to 195 in Perionyx excavatus (shodhganga.inflibnet.ac.in) as well as 34 to 120 in Tubifex tubifex (http://eol.org/pages/620440/overview). Barring the exceptional polychaetes and oligochaetes, the segment number of annelids is a labile trait and is regulated mostly by food availability and temperature (Van Cleave, 1937) as well as photoperiod (Kharin et al., 2006). In the segmentation process, polychaetes and oligochaetes display following contrasting differences: 1. In oligochaetes, the processes of segmental regeneration and reproduction, irrespective of sexual or clonal, are competitive to draw resources from the chloragogue. Consequently, they mutually eliminate each other (Liebmann, 1942, Okrzesik et al., 2013). 2. In majority of oligochaetes (e.g. enchytraeids), the neoblasts are obligately required for asexual reproduction but not for segmental regeneration. In polychaetes, the presence of neoblasts is reported only from Chaetopterus variopedatus, in which the posterior (without the 14th segment) is not capable of anterior regeneration (Berrill, 1928) and Diopatra amboinensis (see Berrill, 1952), which is not known to regenerate. These neoblasts are not known to obligately require for asexual reproduction. 3. In polychaetes, expression of the genes hedgehog (hh) in the segment-addition zone (SAZ) of Perinereis nuntia (Niwa et al., 2013) and Hox genes in the central nervous system in regenerating Nereis virens (Novikova et al., 2013) is reported to manifest segmentation. But in oligochaetes, grimp expression in the neoblasts and mesodermal cells is reported to manifest segmentation. 4. More than 50% polychaetes are characterized by indirect life cycle involving external fertilization (see p 44) and one or more larval stages lasting for some hours to several months (Giangrande, 1997). But the oligochaetes display direct life cycle involving internal fertilization and completion of entire embryogenesis within the relatively safer cocoon. Consequent to these contrasting features, regeneration in polychaetes and oligochaetes markedly differ from each other. Though this has been overlooked by many reviewers (e.g. Berrill, 1952, Zattara and Bely, 2016), this account considers them separately. Polychaetes: Experimental amputation studies have indicated that the minimum number of segment(s) required to regenerate segments both

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anteriorly and posteriorly is only the 10th in the maldanid Clymenella torquata and 14th in Chaetopetrus variopedatus (Fig. 3.7). In the syllid Procerastea halleziana too, it is limited to the 17th and 18th segments, i.e. the seminal segments, harboring stem cells (a newly coined term) responsible for regeneration, are positioned between the cephalic and anterior region of the trunk (cf Lamellibrachia satsuma, p 96). In these worms, anterior regeneration commences with formation of the head, from which new segments are added posteriorly. The posterior regeneration begins with differentiation of the pygidium, from which new segments are added anteriorly (see Licciano et al., 2012). Remarkably, these seminal segments retain the same respective position in the newly regenerated worms indicating that they direct the reorganization of newly added segments. Whereas the seminal 10th and 14th segments can regenerate anteriorly and posteriorly, their posterior (in the absence of the seminal segment) is unable to regenerate anterior in C. torquata (Moment, 1951) and C. variopedatus (Berrill, 1928) and are not capable of

Figure 3.7 Regenerative potency of representative polychaetes. Seminal segments are boxed. Arabic numerals indicate the segment number. Note the heads are decorated with food acquiring structures in polychaetes. Arrows indicate the directions of regeneration. Continuous lines indicate regenerative segmental regions.

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clonal reproduction. However, P. halleziana, in which the posterior containing 17th and 18th seminal segments regenerate the anterior, is capable of clonal reproduction by architomy (see Table 4.2). The intact cirratulid Dodecaceria concharum is also able to naturally regenerate both anterior and posterior from the 13th seminal segment and is also capable of clonal reproduction. But it is difficult to comprehend why the syllids like Autolytus edwardsii, Haplosyllis spongicola and Trypanosyllis zebra do require (7 + 11) 18 to (7 + 18) = 25, anterior segments to regenerate the posterior, as the head consists of only seven anterior segments in syllids (Fig. 3.8). In S. gracilis, a fragment consisting of 6, i.e. the 19th to 24th segments regenerate all the 18 anterior segments.

Figure 3.8 Regenerative potency of representative oligochaetes. Straight lines with arrow heads indicate the number of body segments and directions of regeneration. The straight lines above the arrow headed lines indicate the number of segments harboring neoblasts. The neoblasts are present in all the segments beyond the boxed number. The dotted lines on either side of the Arabic number indicates the seminal segment, on either side of which the neoblasts harboring segments progressively decrease. Note the narrower head in the earthworms.

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These observations on syllids indicate that the positioned seminal segments between 18th and 34th harbors stem cells. The presence of 17 seminal segments in the mid-body markedly differs from the presence of one or two seminal segments alone in the identified species listed above. Further, any fragment containing ~ 75 segments from any part of the body in the 600-segmented Sabella pavonina is reported to add regenerative segments both anteriorly and posteriorly (Murray et al., 2013). In the spionids too, the fragment consisting of a few segments in Pygospio elegans (Gibson and Harvey, 2000) and 2–3 chaetigers in Amphipolydora vestalis (Gibson and Paterson, 2003) drawn from any part of the body is able to clonally reproduce. This is also true of Potamila torelli (Watson, 1906). These observations imply the presence of regenerative stem cells almost throughout the body. Hence, the polychaetes seem to fall in one group including maldanid, chaetopterid and cirratulid possessing one or two seminal segments and the others including syllids, spionids and sabellids harboring many regenerative seminal segments positioned in almost all the body segments. Although the identified seminal segment(s) in the first group of polychaetes are capable of adding segments both anteriorly and posteriorly, their posteriors (without the seminal segments) are unable to regenerate and add segments anteriorly (e.g. C. variopedatus). Hence, in the first group the position of seminal segment(s) harboring stem cells is positioned in the anterior part of the trunk region immediately behind the cephalic region and the second group by almost all the segments are seminal. This may imply that the regenerative stem cells harbored in all the seminal segments are lost from the second group and are progressively limited to one or a few segments in the first group and thereby confirm the postulation by Bely (2006, 2010). Oligochaetes: In general, the ‘head’ in an annelid is recognizable from the rest of the trunk. Using Methyl Green-Pyronin (MGP) staining technique, Myohara (2012) recognized the presence of a pair of neoblasts located in all the trunk segments but not in the first seven ‘head’ segments and the 8th, i.e. the 1st trunk segment in a 50 to 70 segmented oligochaete Enchytraeus japonensis. Thereby the presence of a head and trunk in this worm can be distinguished. This finding seems to hold true for a range of oligochaetes from the small E. japonensis to a large 120 segmented Lumbriculus variegatus. With reference to segmentation, Myohara (2004) has reported an important observation that in the 7th segmented head of oligochaetes, segmentation occurs in sequence from anterior to posterior in embryogenesis but simultaneously in regeneration. Hence, segmentation and its reorganization during regeneration in annelids assume a great importance. A prelude on the segmentation process in E. japonensis is required for a better understanding of segmentation in different taxonomic groups. Molecular level studies have shown that regeneration occurs through the epimorphic regeneration of the head and tail, and morphallactic transformation of old segments into the appropriate middle segments.

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In these studies, different combinations of techniques are employed; for example, MGP staining and BrdU-labeling and detection (Myohara, 2004) or simple hematoxylin and eosin staining, sequence analysis of cDNA clones and expression of marker genes (Takeo et al., 2008). Confirming the finding of Randolph (1892), Myohara (2004) has shown that each segment from the 9th to the last 34th contains neoblasts in asexually reproducing E. japonensis and is capable of regenerating a complete head and a complete tail, irrespective of its position in the body, from which it has been derived. Therefore, no antero-posterior gradient of regeneration potential exists in the trunk. Yet, the number of regenerated segments is dependent on the position of amputation along the body axis, when amputation is made within the head segments of 1 to 7, where no neoblast is present. Conversely, E. buchholzi, which reproduces sexually alone and possesses no neoblasts in any of its segment, does not regenerate a complete head but displays an antero-posterior gradient, when subjected to amputation. Hence, the presence of neoblasts correlates with an absence of antero-posterior gradient of regeneration ability along the body axis. The presence of neoblasts is obligatory for asexual reproduction but not for regeneration of missing body parts. This observation also distinctly delineates regeneration from asexual reproduction. Incidentally, of 27 aeolosomatid clonal species, five alone alternate clonal with sexual reproduction (Falconi et al., 2015). A few lumbriculid species (e.g. Lumbriculus variegatus, Morgulis, 1907) are capable of regeneration, besides clonal reproduction. In enchytraeids, eight species (Christensen, 1959, Collado et al., 2011) out of 670 (Schmelz and Collado, 2012) are capable of either regeneration or clonal reproduction at a time. In naidids, the majority of them are capable of clonal reproduction. However, five species (e.g. Chaetogaster) can clonally reproduce but are not capable of regeneration (Bely and Sikes, 2010). On the other hand, all the investigated lumbricid and megascolecid earthworms (except Perionyx excavatus) do not clonally reproduce but are capable of regeneration. Hence, regeneration and clonal reproduction are quite independent processes in oligochaetes. In E. japonensis, Takeo et al. (2008) have isolated three region-specific genes namely EjTuba, mino and horu. In the normally growing worm, EjTuba and mino are expressed in the head and trunk, respectively. The expression areas for EjTuba and horu in the trunk are proportionate to the total number of segments. In a regenerating worm, the expression of these genes is suppressed but subsequently restored on the 7th day, when regeneration is completed. Using these molecular markers, Takeo et al. have found that the epimorphic regeneration is completed on day 4. Subsequently, morphallactic regeneration occurs between the 4th and 7th day. As a result, the epimorphically regenerated head reorganizes the subsequent morphallactic segmental organization. The morphallactic regulation arises from the balance of molecular gradient derived from the head and tail. Briefly, the morphallactic regeneration regulates the segmentation process but the epimorphic regeneration of

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head and tail regulates segmental reorganization through the newly formed antero-posterior axis. Using immunohistochemistry and immunoblotting with antibodies against these proteins, Martinez et al. (2005) have shown the existence of a gradient distribution of mannose-rich neural glycoproteins with varying molecular masses along the antero-posterior body axis and overlapping in the architomic fission zone of Lumbriculus variegatus. It is known that the neoblasts are firmly adhered to the posterior surface of septa of E. japonensis. Sugio et al. (2012) have discovered a second type of smaller neoblast-like cells designated as N-cells. The N-cells are located dorsal to the neoblasts on the septa. A BrdU labeled pulse chase study has shown that neoblasts are slow cycling in the growing intact worm and possess stem cells characteristics, as evidenced by the expression of Ej-vlg2 and by the telomerase activity during regeneration. Both neoblasts and N-cells of are mesodermal origin, and actively proliferate and migrate toward the autotomized site to form the mesodermal region of blastema. In Lumbriculus variegatus too, any segment beyond the 8th is capable of regenerating a complete head and a complete tail. However, the regenerative potential progressively decreases posteriorly, indicating the increasing loss of neoblasts from the posterior end (Morgulis, 1907). In the naidid Stylaria fossularis, the distribution is limited between the 8th and 22nd segments with the maximum on the 18th segment and progressive decreases on either direction (Fig. 3.8). In Nais commensalis too, the maximum is on the 17th segment with progressive decreases from 16th to 12th segment anteriorly and 18th to 23rd posteriorly (Kharin et al., 2006). Investigations on the naidids Dero limosa (Hyman, 1916), Pristina longiseta (Van Cleave, 1937) and Nais paraguayensis (O’Brien, 1946) suggest that the distribution of neoblasts is a labile feature. It shifts toward posterior, with favorable food availability and temperature but anterior during unfavorable winter temperature conditions. In Tubifex tubifex, the amputations upto the 12th anterior segment results in regeneration of the hypomorphic 3-segmented head but the posterior beyond the 15th segment is unable to regenerate the anterior. Evidently, the number of segments harboring neoblasts is limited between the 13th and 14th segments alone. However, the tubificids, which are unable to clonally reproduce, have opted to parthenogeneic mode of reproduction (Table 2.3). The trend for the loss of neoblasts from posterior is almost total in the megascolecid (except in P. excavatus) and lumbricid earthworms, as they are unable to reproduce clonally. The following descriptive surgical studies on the easily available earthworms have formed the basis for the segmentation process. In Eisenia foetida, Gates (1950) has noted that regeneration potency decreases with increasing number of amputated segments. In 1943, Moment also recorded that the rate of regenerative growth is faster, when the position of amputation is closer to anterior segment. In a key publication, Liebmann (1946) reported that the rate of head regeneration in E. foetida is progressively decreased with amputation increasing from the 9th segment toward

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posterior segment. Incidentally, his observation confirms that the first 8th anterior segments do not possess neoblasts (see Randolph, 1892, Myohara, 2004). Xiao et al. (2011) have made amputation at selected positions (head, up to 8th segments, pre-clitellar 9th to 25th segment, clitellar from 26th to 32nd segments and post-clitellar segments from the 33rd segment onwards and have observed survival and regeneration of E. foetida. Their observations may briefly be summarized: 1. Regeneration commences with blastema formation at the amputated end, followed by epimorphic cellular proliferation and differentiation of the head and tail bud. At 4 hours following amputation, the wound is sealed, 5–7 days after, a conical, unpigmented tail bud is developed, 7–11 days after, segmental delineation begins and 12–16 days after, segments are formed followed by increasing segmental dimension but not segmental number. 2. Survival of amputated worms is linearly and significantly increased with the number of remaining segments but not with position of amputation. 3. However, the anteriorly regenerated body length is correlated with the position of amputation but not with the remaining segments. 4. The posteriorly regenerated body length is correlated neither with a number of remaining segments nor with a position of amputation. Clearly, there are remarkable differences between the formation of anterior head and posterior tail. Mostly, anterior regeneration is achieved, albeit at low survival. In another earthworm Allolobophora caliginosa, the anterior is generated by a fragment containing the 16th segment, indicating the presence of regenerative stem cells in the 16th segment. Briefly, the presence and distribution of the stem cells in oligochaetes and seminal segments or its equivalent stem cells in polychaetes vary from family to family and in families like the syllids from species to species. Clearly, regeneration in annelids is a labile process and has independently originated and/or lost multiple numbers of times (see Bely, 2006, 2010).

3.4  Anterior vs Posterior A large volume of literature concerning anterior and/or posterior regeneration in polychaetes and oligochaetes has been accumulated. Surprisingly, no author has defined the anterior and posterior fragments. A reason for it seems to be the fact that despite the uniformity of segments throughout the body, different functional regions with varying number of segments have been recognized. For example, the polychaete body is considered by different authors as divisible into (i) two regions, the anterior with 1–8 segments and posterior with 8–12 segments in the maldanid Euclymene oerstedi (Clavier, 1984), (ii) three regions, the head, thorax (12 segments) and abdomen (21) in the sabellid Bispira brunnea (Davila-Jimenez et al., 2017) and (iii) into four regions, head, thorax (10–12 segments), abdomen (20–35) and tail (6–12) in

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Table 3.7 Ability of annelids to regenerate anterior or posterior segments (compiled from Bely, 2006, also added from other sources) Anterior regeneration occurs Maldanidae:  Clymenella torquata, Euclymene oerstedi, Petaloproctus socialis Phyllodocidae:  Eulalia viridis Syllidae:  Autolytus pictus, Procerastea halleziana, Syllis spp, Streptosyllis websteri, Typosyllis prolifera, T. pulchra, Trypanosyllis asterobia Amphinomidae:  Eurythoe complanata Dorvilleidae:  Dorvillea bermudensis Eunicidae:  Lysidice sp, Nematonereis unicornis Onuphidae:  Diopatra spp Oweniidae:  Owenia fusiformis Sabellidae:  Branchiomma nigromaculata, Myxicola aesthetica, Sabella spp Serpulidae:  Hydroides dianthus Spionidae:  Dipolydora quadrilobata, Pygospio elegans Criodrillidae:  Criodrilus lacuum Enchytraeidae:  Enchytraeus dudichi, E. fragmentosus, E. japonensis Lumbricidae:  Eisenia foetida, Lumbricus terrestris Lumbriculidae:  Lumbriculus lineatus, Rhynchelmis vagensis Megascolecidae:  Metaphire peguana, Perionyx excavatus, Pheretima sp Naididae:  Allonais paraguayensis, Dero digitata, Limnodrilus claparedianus, Nais elinguis, Stylaria spp Tubificidae:  Tubifex tubifex Posterior regeneration occurs Eunicidae:  Eunice fucata, E. siciliensis, E. viridis (Hofmann, 1974) Syllidae:  Autolytus cornuta, Brania pusilla, Eusyllis blomstrandi, Exogone naidina, Grubeosyllis clavata, Haplosyllis spongicola, Haplosyllides floridana, Myrianida pachycera (Franke, 1999), Odontosyllis enopla, Pionosyllis lamelligera, P. procera, P. pulligera, Odontosyllis prolifera, O. polycera (Fischer and Fischer, 1995), O. phosphorea (Tsuji and Hill, 1983) Anterior regeneration does not occur Arenicolidae:  Arenicola marina Capitellidae:  Capitella sp I, Capitella sp II, Mediomastus sp Opheliidae:  Polyphthalmus pictus Nereididae:  Platynereis dumerilii Polynoidae:  Harmothoe imbricata Dorvilleididae:  Ophryotrocha notoglandulata, O. puerilis puerilis Eunicidae:  Eunice afra, E. schizobranchia, E. siciliensis, E. torquata, E. viridis Spionidae:  Streblospio benedicti Species/taxa, in which posterior regeneration does not occur Dinophillidae:  Dinophilus gardnieri Arenicolidae:  Arenicola marina Opheliidae:  Polyophthalmus pictus Hirudinea:  Almost all species

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the spionid Pygospio elegans (Gibson and Harvey, 2000). In the siboglinid Lamellibrachia satsuma, it is divisible into the three non-segmented tentacular, vestimental and trunk regions and the fourth segmented opisthosoma region (see p 95). Considering the head alone, this consists of seven segments in oligochaetes and can be recognized by the absence of chloragogue, metanephridia (e.g. Nais communis, Kharin et al., 2006) and neoblasts (e.g. Enchytraeus japonensis, Myohara, 2012). Regeneration potency of the head ranges widely from Paranais litoralis, which is unable to regenerate the head to Lumbriculus variegatus, which stereotypically regenerate a restricted but precise number of 7–8 head segments, regardless of the position of amputation (see Martinez-Acosta and Zoran, 2015). Nevertheless, it is possible to distinguish the anterior from posterior in natural fission from field and experimental observations. In the cirratulid Dodecaceria concharum, natural fission occurs between the 1–13 anterior segments and 13– 34 posterior segments (Martin, 1933). In the amphinomid Eurythoe complanata, a field study has shown that the anterior consists of 16 segments and posterior 40 segments (Kudenov, 1974). Experimental studies have identified the 10th, 14th and 17–18th seminal segments in Clymenella torquata, Dodecaceria concharum, Chaetopterus variopedatus, Procerastea halleziana in polychaetes (Fig. 3.7) and Stylaria fossularis in oligochaetes (Fig. 3.8). All the segments in front of the respective seminal segments constitute the anterior and those behind them the posterior. Similarly, all the segments ahead of the 20-segmented mid-body constitute the anterior and those behind it posterior in Perionyx excavatus (Fig. 3.1). Pigmentation has been used as a marker of pre-fragmentation stage in Dodecaceria pulchra (Gibson, 1977) and post-pigmentation stage in Bispira brunnea (Davila-Jimenez et al., 2017). In Typosyllis antoni, anterior can be identified by its distinct red lines on the dorsal and posterior by the presence of bendantate chaetae and a tiny thin acicula (Aguado et al., 2011). Besides them, the identified seminal segments or the mid-body enable the identification of anterior and posterior fragments. However, the fact that any segment beyond the 8th can regenerate the entire body in 34-segmented E. japonensis and 120-segmented Lumbriculus variegatus in oligochaetes and a few (2–3 chaetigers) segments in the spionids Amphipolydora vestalis (Gibson and Paterson, 2003), P. elegans (Gibson and Harvey, 2000) and in the 600-segmented sabellids Sabella pavonina (Murray et al., 2013) in polychaetes complicate the identification of anterior and posterior. Despite these constraints, it is still possible to understand the differences between anterior and posterior fragments and their distribution across the phylogenetic groups of polychaetes and oligochaetes. Thankfully, Zattara and Bely (2016) have accomplished an onerous task of assembling the relevant information on regeneration in 247 species belonging to 28 families of polychaetes and 129 species belonging to 7 families of oligochaetes. Zattara (2012) must be complimented for accomplishing this job earlier and promptly sending the appendix of Zattara and Bely (2016) on

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email request ([email protected]). For immediate reference, Table 3.7 lists some of these annelids with potency for anterior, posterior and/or anterior cum posterior regeneration. The said appendix provides an opportunity for further analysis. In view of the contrasting features, regeneration and clonal reproduction in polychaetes and oligochaetes are separately considered. From Table 3.7, the following inferences can be made: 1. The number of species capable of anterior, posterior and anterior cum posterior regeneration is 149, 206 and 143, respectively (Table 3.8). A calculation of the said numbers as fractions of 16,931 annelid species (Table 1.2) indicates that only 0.88, 1.22 and 0.85% of annelids are capable of anterior, posterior and anterior cum posterior regeneration, respectively. Comparing even the 1.22% incidence for the posterior regenerative potency of the bilaterally symmetrical annelids with 2.95% of 7,000 speciose radially symmetrical echinoderms (Pandian, 2018), the potency of annelids is far less than that of echinoderms. It is not clear, whether radial symmetry facilitates a greater regenerative potency. Comparative studies on the incidence frequency of regeneration between the bilaterally symmetrical non-segmented, acoelomate Turbellaria and the segmented coelomate annelids as well as bilaterally symmetrical hydrozoan cnidarians and radially symmetrical schiphozoan cnidarians may clarify it. 2. In annelids, posterior regeneration occurs (incidence frequency 1.2%) more frequently than the anterior (0.85%) indicating that the former costs less than that of the latter. The reasons for it are listed: (i) in most polychaetes and oligochaetes, the anterior consists of the brain, heart and metanephridia as well as reproductive organs in oligochaetes. (ii) regenerative potency diminishes faster in the ‘old’ anterior segments than in the ‘young’ posterior ones (see Paulus and Muller, 2006), (iii) in oligochaetes, the loss of neoblasts progresses from the posterior end (e.g. Lumbriculus variegatus) toward the mid-body (e.g. Stylaria fossularis) and (iv) amputation causes a strong shift Table 3.8 Regeneration in annelids. Percentage values in brackets. For details see text (estimated from Appendix of Zattara and Bely, 2016) Regenerating Fragment(s)

Total (no.)

Polychaeta (no.)

Oligochaeta (no.)

Anterior only

149 (0.88)

104 (0.80)

45 (1.57)

Posterior only

206 (1.22)

146 (1.12)

60 (1.89)

Anterior + posterior

143 (0.85)

98 (0.75)

45 (1.42)

3

2

1

61 (43)

32 (33)

29 (64)

Anterior + posterior but no cloning

34 (23.8)

31 (31.6)

3 (6.7)

Anterior without cloning

19 (13.2)

18 (17.3)

1 (2.2)

Posterior without cloning

15 (10.5)

10 (6.9)

5 (11.2)

6

0

6

No anterior + posterior Anterior + posterior and cloning

No anterior + posterior but cloning

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in resource allocation from growth to regeneration and the shift is strong and persists longer during anterior regeneration than the posterior (Zattara and Bely, 2013). The values of 149 for anterior and 143 for anterior cum posterior indicate that the former is usually accompanied by the posterior (Table 3.8). In the absence of anterior, the posterior regeneration occurs only in the naidids Amphichaeta raptisae, Chaetogaster diaphanus, C. diastrophus, Paranais frici and P. litoralis (see Bely and Sikes, 2010). 3. When the number of anterior, posterior and anterior cum posterior regeneration is considered as a fraction of 13,012 polychaete species and 3,175 oligochaete species (Table 1.2), the percentage values (1.57, 1.80, 1.42) obtained indicates that the prevalence of regenerative potency is 1.5 times greater in oligochaetes than the respective ones (0.76, 1.15, 0.76) of polychaetes. 4. Only 61 species (out of 143, i.e. 42.6%) possess clonal potency; of them, 33% (i.e. 32 species out of 98) polychaetes and 64% (i.e. 29 species out of 45) oligochaetes are characterized by anterior cum posterior regeneration along with clonal reproduction. Surprisingly, all the 32 polychaete species per se undergo clonal reproduction by architomy. Of 29 oligochaete species per se, 19 and 21 species undergo clonal reproduction by paratomy and architomy, respectively. When these 32 and 29 species numbers are related to 98 polychaete species and 45 oligochaete species characterized by per se, it is only 32% of the polychaetes undergo architomic clonal reproduction (with a couple of exceptions), in comparison to 64% of oligochaetes. 5. As indicated earlier, regeneration and clonal reproduction are quite independent processes. Hence, it is not correct to consider that the latter is derived from the former (Zattara and Bely, 2016). 5a. In clonal oligochaetes, the presence of neoblasts is obligatorily required to manifest the clonal potency. In the absence or loss of the neoblasts, almost all the lumbricid and megascolecid earthworms (except Perionyx excavatus, see p 96, Fig. 3.1), some enchytraeids (e.g. Lumbricillus lineatus, Enchytraeus buchholzi [Myohara, 2012]) and naidids (e.g. Limnodrilus claparedianus, Tubifex rivulorum, T. tubifex) are unable to clonally reproduce. Even in the absence of anterior cum posterior regenerative ability clonal reproduction does occur in Chaetogaster diastrophus. Clearly, the clonal potency of oligochaetes arises from the neoblasts and not from anterior cum posterior regenerative potency. 5b. In polychaetes, however, 60% species characterized by anterior (18.3%), posterior (10.2%) and anterior cum posterior (31.6%) regeneration are unable to clonally reproduce. Only 33% of them characterized by anterior cum posterior regenerative ability are able to clonally reproduce. Researches in polychaetes to identify the factor responsible for clonal reproduction are urgently required. 6. Despite the absence of anterior cum posterior regeneration, C. diaphanus and C. diastrophus are unique being capable of clonal reproduction by paratomy. 7. Both anterior and posterior regeneration is absent Dinophilus gardnieri, Polyophthalmus pictus (Opheliidae) and Arenicola marina in polychaetes as well as C. diaphanus and C. diastrophus in oligochaetes. For immediate reference Table 3.9 lists the families, in which anterior, posterior and anterior cum posterior regeneration occurs. Whereas the

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Table 3.9 Families in which anterior, posterior or anterior cum posterior regeneration occurs. Continuous underline indicates the incidence of regeneration in almost all the investigated species of the family. Dotted underline indicates the incidence in a few of investigated species of the family (compiled from Appendix of Zattara and Bely, 2016) Anterior Polychaetes:  Oweniidae, Chaetopteridae, Amphinomidae, Saccocirridae, Dorvilleidae, Eunicidae, Onuphidae, Syllidae (exception: Brania pusilla), Polygordiidae, Nephtyidae, Hesionidae, Phyllodocidae, Orbiniidae, Siboglinidae, Cirratulidae, Ctenodrilidae, Sabellidae, Serpulidae, Spionidae (exception: Streblospio benedicti), Terebellidae, Maldanidae Oligochaetes:  Aeolosomatidae, Naididae, Enchytraeidae, Glossoscolecidae, Lumbricidae, Megascolecidae, Lumbriculidae Posterior Polychaetes:  Oweniidae, Chaetopteridae, Amphinomidae, Saccocirridae, Dorvilleidae, Eunicidae, Onuphidae, Syllidae, Polynoidae, Polygordiidae, Nephtyidae, Hesionidae, Nereidae, Phyllodocidae, Tomopteridae, Orbiniidae, Siboglinidae, Cirratulidae, Ctenodrilidae, Sabellidae, Sabellaridae, Serpulidae, Spionidae, Captellidae, Terebellidae, Maldanidae Oligochaetes:  Aeolosomatidae, Naididae, Enchytraeidae, Glossoscolecidae, Lumbricidae, Megascolecidae, Lumbriculidae Anterior cum posterior Polychaetes:  Oweniidae, Chaetopteridae, Amphinomidae, Saccocirridae, Dorvilleidae, Eunicidae (only in Lysidice collaris, L. ninetta, Nematonereis unicornis), Onuphidae, Syllidae, Polygordiidae, Orbiniidae (only in Proscoloplos cygnochaetus, Scoloplos armiger), Cirratulidae, Sabellidae, Serpulidae, Spionidae (exception: Myxicola aesthetica), Maldanidae Oligochaetes:  Aeolosomatidae, Naididae, Enchytraeidae, Lumbricidae, Megascolecidae, Lumbriculidae

incidence of posterior regeneration is spread over as many as 33 families, that of anterior and anterior cum posterior is limited to 28 and 15 families, respectively.

4 Asexual Reproduction

Introduction Besides the amazing potency of regenerating the missing body parts from a small feeding structure to larger fragment of the entire anterior and/ or posterior body, some annelids are capable of agametically clone and reproduce asexually. As indicated earlier, regeneration and reproduction are quite independent processes. In oligochaetes, the presence of ‘multipotent’ neoblasts is obligately required to manifest clonal reproduction. Sex is costly, and demands time and resource but clonal reproduction saves them and avoids the risks involved in sexual reproduction. It also provides a mechanism for potential rapid amplification of a genotype known for its fitness. However, it involves no gametogenesis and recombination. In the absence of recombination and fusion of gametes in clonally reproducing polychaetes and oligochaetes, adaptation can be impeded and deleterious mutations may be accumulated, due to Muller’s ratchet (Engelstadter, 2008). Understandably, the incidence of clonal reproduction is limited to ~ 190+ species, i.e. 1.1% of annelids; it is also limited to a few polychaete and oligochaete species alone. In them, the clonal reproduction can broadly be grouped into architomy and paratomy. In architomy, fission is followed after completion of regeneration and formation of progenies (ramets). But, it occurs even before the ramets are fully formed in paratomy. Further, a fission zone(s) is formed, prior to fragmentation in paratomics but not in architomics.

4.1  Obligate Cloners? About a dozen polychaete and oligochaete species are reported to survive and flourish by clonal reproduction alone for periods from 3 years (Aeolosoma

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hemprichii) to 60 years in Zeppelina monostyla and Pristina leidyi (Table 4.1). In Polydorella kamakamai, sexual reproduction is rare or transient in the field and covers only 0.3% of the population (Williams, 2004). These observations may arguably question the general conclusion that no animal species exclusively reproduce asexually (see Pandian, 2016). While the institutions at Bologna, Table 4.1 Obligately asexually reproducing polychaetes and oligochaetes Species/Reference

Reported Observations Polychaetes

Cirratulidae: Zeppelina monostyla (see Akesson and Rice, 1992)

No sexual reproduction observed for 60 years in an Aquarium Freiburg, Germany

Dorvilleidae: Parougia albomaculatus P. bermudensis (Akesson and Rice, 1992)

On breeding in the laboratory (University of Goldberg, Sweden), no indication of sexual reproduction observed over 15–17 years

Spionidae: Pygospio elegans (Anger, 1984) Polydorella kamakamai (Williams, 2004)

A single population entirely relies on asexual reproduction

Sabellidae: Perkinsiana milae (Gambi et al., 2000)

In the field, sexual reproduction is rare or transient and covers only 0.3% population In the field collected specimens, no gametes in the coelom. Indications for fission within a single tube Oligochaetes

Enchytraeidae: Cognettia sphagnetorum (Christensen, 1959) Enchytraeus higentius (Christensen, 1984) Aeolosomatidae: Aeolosoma hemprichi (Stolc, 1903) A. viride (Falconi et al., 2015) Naididae: Nais communis Potamothrix bedoti P. vejdoskyyaneum (Timm, 1984) Pristina leidyi (Ozpolat and Bely, 2015) Stylaria lacustris (see Schierwater and Hauenschild, 1990)

Sexually mature worm is very rare at any season in Denmark. Sexual eggs never hatch Asexual reproduction totally suppressed only at high density Survive in laboratory for 3 years by only paratomic fission Reproduce only by paratomic fission for > 20 years in the laboratory of Bologna University, Italy Survive in laboratory (Estonia) for 6 years only by paratomic fission A clone flourishes for 8 years without any sign of senescence (cf Martinez and Levinton, 1992) Clonal reproduction by only paratomic fission for 20 years (University of College Park, USA) and earlier for 40 years (Carolina Biological Supply Co, USA) At LD = 16 hours and T = 20ºC, the naidid reproduced asexually alone for > 6 years

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Freiburg, Goteburg and College Park deserve admiration for accomplishing the arduous task of maintaining clonally reproducing annelids over long years, it must be noted that these clonally reproducing worms have been maintained at optimal rearing conditions enabling them to reproduce clonally alone. The absence of gonads and gametes (see Table 4.8) and failure of spawning in some seasons/years (e.g. Streblospio benedicti, Nephtys hombergii) may not indicate that the species concerned is asexual. For, the failure of spawning may be a consequence of poor nutritional conditions (see Kolbasova et al., 2013). When stressed/induced, these worms can switch over to sexual reproduction. Indeed, P. leidyi is capable of becoming sexual after ~ 1,000–3,000 agametic rounds of cloning over a prolonged period of 60 y (Ozpolat and Bely, 2015).

4.2  Incidence and Prevalence Table 4.2 summarizes the incidence of clonal reproduction in polychaetes and oligochaetes. Some species like the oweniid Myriochele heeri is not included by Zattara and Bely (2016). However, Oliver (1984) reported 30% prevalence of clonal reproduction in it. Publications by Lohlein (1999) and Naidu (2005) have shown the need for inclusion of another 35 species. Hence, this list includes all of them. It has formed the base for further analysis and lists many new findings. 1. Of 100 and odd annelid families, clonal reproduction is limited to 12 polychaete families and five oligochaete families (Table 4.3). This estimate on clonal incidence reveals that the incidence is limited to 79 polychaete and 111 oligochaete species. Together, they make 190 species, i.e. only 1.1% of annelids are capable of clonal reproduction, which may be compared with 1.90% for echinoderms (Pandian, 2018). 2. When the values are related to the respective number of polychaetes (13,012) and oligochaete (3,175) species, the incidence of clonal reproduction in oligochaetes is ~ 4.6-times greater (3.18%) than that (0.68%) in polychaetes. 3. Further analysis of the incidences of architomy and paratomy has revealed the following: Among polychaetes, the incidence of architomy is widespread over 12 families, but paratomy is limited to four families alone (Table 4.3). Architomy occurs exclusively in eight polychaete families but paratomy exclusively in Aeolosomatidae in oligochaetes and Dinophilidae in polychaetes. The incidence within a family ranges from < 2% (13 out of 700 species, for species number see Franke, 1999) in Spionidae to 54% (69 out of 175 species, for species number see Ferraguti et al., 1999) in Naididae. Secondly, in terms of species number, the incidence is higher (75.6%, 60 species out of 79) for architomy in polychaetes but 77% (81 species out of 100) for paratomy in

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Table 4.2 Annelid species reported to asexually reproduce; species in bold letters were not included in Zattara and Bely, 2016 1. Polychaetes Architomy (i) Oweniidae: Myriochele heeri (prevalence: 30%, Oliver, 1984) (ii) Chaetopteridae: Phyllochaetopterus prolifica (see Purscke, 2006), P. socialis (see Kudenov, 1974), Spiochaetopterus costarum costarum, Spiochaetopterus solitaries (Zattara and Bely, 2016) (iii) Amphinomidae: Eurythoe complanata (see Kudenov, 1974), Linopherus canariensis (Zattara and Bely, 2016) (iv) Dorvilleidae: Parougia albomaculatus, P. bermudensis (see Akesson and Rice, 1992) (v) Syllidae: Odontosyllis gibba (see Kudenov, 1974), O. ctenostoma (Zattara and Bely, 2016), Procerastea halleziana (Allen, 1923), Syllis gracilis (see Franke, 1999) (vi) Orbiniidae: Proscoloplos cygnochaetus (Zattara and Bely, 2016) (vii) Cirratulidae: Caulleriella viridis, Cirratulus cirratus (Zattara and Bely, 2016), Dodecaceria berkeleyi (see Kudenov, 1974), D. concharum (Gibson, 1977), D. coralii (Gibson, 1978), D. fistulicola, (see Kudenov, 1974), D. fewkesi, D. fimbriata (Berkeley and Berkeley, 1954), D. pulchra, Timarete filigera, T. punctata (Zattara and Bely, 2016), Protocirrineris chrysoderma (Purscke, 2006), P. antarctica (Zattara and Bely, 2016) (xiva) Ctenodrillidae: Raphidrilus nemasoma, Raricirrus beryli, Zeppelina monostyla (see Akesson and Rice, 1992), Raricirrus maculatus, R. arcticus (A?) (Buzhinskaja and Smirov, 2017) (viii) Sabellidae: Bispira brunnea (Davila-Jimenez et al., 2017), Branchiomma bairdi (Arias et al., 2013), B. curtum (see Tovar-Hernandez and Knight-Jones, 2006), Megalomma cinctum (Yuan, 1992), Myxicola aesthetica (Knight-Jones and Bowden, 1984), Perkinsiana milae (Gambi et al., 2000), P. rubra, Potamilla torelli, Pseudobranchiomma emersoni, P. perkinsi (Knight-Jones and Giangrande, 2003), P. punctata, P. minima (Nogueira and Kinght-Jones, 2002), P. schizogenica (see Davila-Jimenez et al., 2017), Pseudopotamilla reniformis (Kolbasova et al., 2013), Sabella discifera (Rioja, 1929), S. pavonina, Sabellastarte sp (Murray et al., 2013) (xva) Serpulidae: Josephella sp, Rhodopsis simplex (Zattara and Bely, 2016) (xvia) Spionidae: Amphipolydora abranchiata (Blake, 1983), A. vestalis (Gibson and Paterson, 2003), Dipolydora armata (Radashevsky and Nogueira, 2003), Dipolydora caulleryi, D. socialis (Allen, 1921), Polydora colonia, P. elegantissima (Allen, 1921), Pygospio californica, P. elegans (Gibson and Harvey, 2000) Paratomy (xii) Dinophilidae: Dinophilus rostratus (Zattara and Bely, 2016) (xivb) Ctenodrilidae: Ctenodrilus serratus (Gibson, 1977), Kirkegaardia (Purscke, 2006) (xvb) Serpulidae: Filogranella elatensis, F. gracilis, Josephella marenzelleri, Rhodopsis pusilla, Spiraserpula snelli (see Halt et al., 2006), F. implexa, Salmacina amphidentata, S. australis (Zattara and Bely, 2016), S. incrustans (Schroeder and Hermans, 1975), S. dysteri (Nishi and Nishihara, 1994) (xvib) Spionidae: Polydorella dawydoffi, P. kamakamai, P. tetrabranchia (Gibson and Harvey, 2000), P. smurovi, Polydorella prolifera (Tzetlin and Britayev, 1985), P. stolonifera (Zattara and Bely, 2016) Table 4.2 contd. ...

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...Table 4.2 contd. 2. Oligochaetes Architomy (ix) Enchytareidae: Buchholzia appendiculata, Cognettia sphagnetorum (Christensen, 1959), Enchytraeus variatus (Bouguenec and Giani, 1989), E. dudichi, E. fragmentosus, E. japonensis (Collado et al., 2011), Cognettia glandulosa, Enchytraeus bigeminus, Marionina sp (Zattara and Bely, 2016) (x) Lumbriculidae: Lumbriculus variegatus (Martinez et al., 2006) (xi) Megascolecidae: Perionyx excavatus (S. Sudhakar, pers. comm.) (xviia) Naididae: Allonais paraguayensis (Bely and Sikes, 2010), A. inaequalis, A. lairdi, A. pectinata, Autodrilus japonicus, A. pluriseta, A. sp, Bothrioneurum righii, B. vedjioskyanum, Branchiodrilus menoni, B. semperi, Bratislavia unidentata, Dero bauchiensis, D. borelii, D. lutzi, D. malayana, Pedonais crassifaucis, Slavina evelinae, S. sawayai (Zattara and Bely, 2016) Paratomy (xiii) Aeolosomatidae: Aeolosoma travancorense (Aiyer, 1926), A. hemprichi, A. quaternarum, A. singutare, A. titorale (see Falconi et al., 2015), A. kashyapi, A. niveum, A. spp, A. viride (Zattara and Bely, 2016), A. beddardi, A. headleyi, A. ternarium (Naidu, 2005) (xviib) Naididae: Arcteonais lomondi, Dero furcata, Dero sp I, Nais communis, N. elinguis, Piguetiella michiganensis, Pristina aequiseta, P. leidyi, Ripistes parasita, Slavina appendiculata, Specaria josinae, Stylaria lacustris (Bely and Sikes, 2010), Dero digitata, Pristina longiseta, Stylaria fossularis (Bely, 1999), Paranais litoralis (Nilsson et al., 1997), Amphichaeta raptisae, A. sannio, Branchiodrilus hortensis, Chaetogaster diaphanus, C. diastrophus, C. limnaei, Cruistipellis tribranchiata, Dero carteri, D. flabelliger, D. gravelyi, D. huaronensis, D. superterrenus, D. tonkinensis, D. vaga, Nais bretscheri, N. stolci, Ophidonais serpentina, Paranais frici, Stephenosoniana sp, Uncinais uncinata, Vejdovskyella sp (Zattara and Bely, 2016), Chaetogaster langi, Nais sp, N. barbata, N. pseudobtusa (Lohlein, 1999), Allonais rayalaseemensis, Aulophorus carteri, A. flagellum, A. furcatus, A. gravelyi, A. hymanae, A. indicus, A. michaelseni, A. moghei, A. tonkinensis, Chaetogaster cristallinus, C. limnae bengalensis, Nais andina, N. andhrensis, N. pardalis, N. simplex, N. variabilis, Pristina breviseta, P. evelinae, P. macrochaeta, P. proboscidae, P. sperberae, P. synchites, Pristinella acuminata, P. jenkinae, P. menoni, P. minuta, Stephensoniana trivandrana (Naidu, 2005)

oligochaetes. Thirdly, 63 out of 79 clonal polychaete species are sedentary/ tubiculous, indicating that clonal reproduction is more frequent in sedentary than errant polychaetes. Fourthly, all the members of 10 families (i) Oweniidae, (ii) Chaetopteridae, (iii) Amphinomidae, (iv) Dorvilleidae, (v) Syllidae, (vi) Orbiniidae, (vii) Cirratulidae, (viii) Sabellidae, (ix) Enchytraeidae and (x) Lumbriculidae are architomic, but (xi) Dinophilidae and (xii)Aeolosomatidae are paratomic. Only, (xiii) Ctenodrillidae, (xiv) Serpulidae, (xv) Spionidae and (xvi) Naididae have representations for architomy and paratomy (Table 4.3). It is likely that each of the first 10 architomic family members underwent mutation for the clonal reproduction perhaps simultaneously at a time. It may also be true for the two paratomic families. However, it is not clear why different members of the last four families undergo architomy or paratomy.

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Table 4.3 Incidence of clonal reproduction in polychaetes and oligochaetes. Estimations are based on Table 4.2 Family

Architomy Species (no.)

Paratomy Species (no.)

Polychaetes Sedentary polychaetes (i) Oweniidae

1

0

(ii) Chaetopteridae

4

0

(vi) Orbinidae

1

0

(vii) Cirratulidae

13

0

(viii) Sabellidae

17

0

(xv) Serpulidae

2

10

(xvi) Spionidae

9

6

Errant polychaetes (iii) Amphinomidae

2

0

(iv) Dorvilleidae

2

0

(v) Syllidae

4

0

(xii) Dinophilidae

0

1

(xiv) Ctenodrillidae

5

2

Subtotal

60

19

Oligochaetes (ix) Enchytraeidae

9

0 0

(x) Lumbriculidae

1

(xi) Megascolecidae

1

0

(xiii) Aeolosomatidae

0

12

(xvi) Naididae

19

69

Subtotal

30

81

Total

90

100

4.3  Observations and Characteristics (i) Publications: (a) There are more publications on clonal reproduction in polychaetes than in oligochaetes. (b) On these annelids, there are more publications on experimental observations than field ones. Not surprisingly, there are limited publications on prevalence of clonal reproduction. (c) Only a few authors have reported observations from both field and experimental study (e.g. Pseudopotamilla reniformis, Kolbasova et al., 2013). Others have reported the findings from either experimental (e.g. Parougia bermudensis, Akesson and Rice, 1992) or field (e.g. Pseudobranchiomma schizogenica, TovarHernandez and Dean, 2014) observation.

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(ii) Phylogeny: (a) The oligochaetes, which have recolonized aquatic habitats, have retained a direct life cycle, although aquatic habitats can sustain haloor mero-planktonic larvae, as in polychaetes. (b) For reasons not yet known, they are unable to manifest a sedentary/tubiculous mode of life, albeit a few inhabit within mucous or gelatinous tube (see Table 4.7). (c) Of 79 clonal polychaetes, 63 (architomic 47 species + paratomic 16 species), i.e. ~ 80% of them are sedentary and/or tubiculous. Of 290 tubiculous sabellids (McEuen et al., 1983), only 17 species, i.e. 5.9% of them are architomic cloners. Hence, it is difficult to consider that an intense sub-lethal predation alone have enforced clonal reproduction in the sedentary/tubiculous polychaetes (cf Oliver, 1984). (iii)  Sexuality: (a) In echinoderms, clonal reproduction occurs only in gonochoric oviparous species (Pandian, 2018). But the clonal polychaetes include protandric (e.g. Salmacina australis) and self-fertilizing simultaneous hermaphrodites (e.g. Bispira brunnea, Davila-Jimenez et al., 2017) as well as brooders. For example, the architomic cloning spionids Dipolydora caulleryi, D. socialis, Pygospio californica and P. elegans are all brooders (see Blake and Arnofsky, 1999). Similarly, the paratomic clonal serpulids Salmacina amphidentata and S. dysteri are brooders (Kupriyanova et al., 2001). (iv) Life stages: (a) Barring crinoids, all other classes of echinoderms have representative species, in which one or other larva reproduces clonally (Pandian, 2018). In annelids, clonal reproduction is limited to immature and mature stages alone. Larval cloning is not so far reported for any annelid. (v) Reproduction: Clonal species alter sex ratio and/or eliminate or reduce the number of gametes. For example, males are unknown in Dodecaceria pulchra (Gibson, 1977) and females in 12 clonal naidids (Table 4.8). Female ratio is also reduced to 0.03 in Polydorella kamakamai (Williams, 2004) and 0.09 in the oweniid Myriochele heeri (Oliver, 1984). Sperms are not observed in Pseudobranchiomma schizogenica (Tovar-Hernandez and Dean, 2014). Egg strings of Amphipolydora vestalis hold 69 eggs + 329 nurse eggs, i.e. 4.7 nurse eggs/egg (Gibson and Paterson, 2003). In P. kamakamai, the 13th, 14th and 15th gametogenic segments generate 39, 30 and 18 eggs, respectively (Williams, 2004). (vi) The trigger: In polychaetes, food availability (e.g. Parougia bermudensis, Akesson and Rice, 1992) and density (e.g. Pygospio elegans, Wilson, 1985) as well as temperature (e.g. P. elegans, Rasmussen, 1953) trigger the initiation and proportion of architomic clonal reproduction. For example, increasing temperature during spring and consequent food availability initiates clonal reproduction in P. elegans. Increasing density may limit food availability and thereby suppress clonal reproduction. Contrastingly, none of these factors control clonal reproduction in naidids. It is photoperiod of < 12 hL that switches on sexual reproduction in Stylaria lacustris (Schierwater and Hauenschild, 1990). This (< 16 hL) is also true of other

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naidids Nais communis and Pristina longiseta (Kharin et al., 2006). However, clonal reproduction, at high densities above 300 and 400 worms, is suppressed in the architomic Enchytraeus japonensis (Myohara et al., 1999) and E. bigeminus (Christensen, 1973), respectively. Dependance on phytoplankton for food has limited the vertical distribution of naidids to euphotic zone (e.g. Martin et al., 1999, Hirabayshi et al., 2014). Understandably, decreasing photoperiod and the consequent diminishing phytoplankton in temperate and sub-arctic freshwater habitats initiate sexual reproduction and production of diapausing cocoons to overwinter. This may be an adaptive strategy to engage photoperiod as a trigger to sexual reproduction. Researches are required to know whether the photoperiod also serve to trigger sexual reproduction in the paratomic polychaetes and tropical naidids, to whom phytoplankton and/or detritus/sediment is available around the year. (vii) Clonal vs Sexual Reproduction: These two reproductive processes intensely compete for resources. Understandably, they may temporally be separated in clonal polychaetes and oligochaetes. In the exceptional sabellid Pseudopotamilla reniformis from the sub-arctic White Sea, clonal reproduction dominates but lasts for (6 months) during autumn and winter. In it, sexual reproduction may rarely occur during spring and summer. Conversely, clonal reproduction dominantly occurs during the favorable spring and summer in the subtropical Pygospio elegans, over periods of 10 months in a year (Fig. 4.1). In them, sexual reproduction occurs only during the winter months. Sexual reproduction is limited to austral summer in the antarctic oweniid Myriochele heeri; but clonal reproduction occurs from austral autumn to spring. In the temperate naidid Stylaria lacustris too, clonal reproduction dominates over a period of 7 months during spring and summer. With the irreversible switch, sexual reproduction is limited to September only. In the enchytraeids, in which clonal reproduction dominates and sexual reproduction is suppressed until the critical densities are attained in the culture system. Nevertheless, both clonal and sexual reproduction concurrently occurs in a few polychaetes and in enchytraeids. In the cirratulid Dodecaceria pulchra (South Africa), clonal reproduction dominates and occurs round the year with parthenogenic eggs being spawned only during late autumn. In it, sexual reproduction is virtually suppressed. Conversely, sexual reproduction dominates and occurs round the year in the sabellid Bispira brunnea (Mexico) with clonal reproduction concurrently occurring almost throughout the year but with a single ramet/fragmentation. In it, clonal reproduction does not also occur during spawning. Experimental study on the spionid Amphipolydora vestalis (New Zealand) indicates the concurrent occurrence of clonal and sexual reproduction. In fact, 46% of the ramets carry gametes and a fragment holding three chaetigers is adequate to accomplish bidirectional clonal reproduction. This is also true of Lumbriculus variegatus (Morgulis, 1907). It is not clear whether sexual and clonal reproduction in A. vestalis can

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Figure 4.1 Temporally separated or concurrently synchronized clonal (– – –) and sexual (–—) reproduction in architomic polychaetes and oligochaetes. Symbols marked by indicate parthenogenic reproduction. Thick lines indicate dominant frequency and thin line poor frequency. Lines marked by the symbols indicate clonal and    sexual reproduction at different densities in culture system. Lines marked by the symbols ð ð ð represent the period, during which cocoons diapause.

concurrently occur in the fields also. The concurrent occurrence of clonal and sexual reproduction is hinted in Enchytraeus bigeminus (Christensen, 1973) and E. variatus (Bouguenec and Giani, 1989). With a life span of ~ 85 days, clonal reproduction dominates initially in the ‘young’ worms but is replaced subsequently by sexual reproduction in the ‘old’ worms (Fig. 4.1).

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Prevalence: Available information is limited to 11 architomic and one paratomic species. Among architomics, the values range from 4 to 13% for sedentary sabellid Megalomma cinctum to 90% in errant Lumbriculus variegatus and up to 100% in paratomic Polydorella kamakamai (Table 4.4). Of 297 tubiculous sabellid species, only 17 are cloners. Hence, the sublethal predation and the consequent prevalence of regenerates may be more related to the predatory pressure rather than errant/sedentary habit or architomic/ paratomic fragmentation. Table 4.4 Prevalence of clonal reproduction in field populations of some polychaetes and oligochaetes Family/Species/Reference

Reported Observations Errant polychaetes

Amphinomidae Eurythoe complanata (Kudenov, 1974)

Architomy prevalent in 30% of population during spring from April to July

E. oerstedi (Clavier, 1984)

Architomy prevalent in 40 and 32% of anterior and posterior fragments, respectively

Oweniidae Myriochele heeri (Oliver, 1984)

Architomy prevalent in 30% of population

Cirratulidae Dodecaceria pulchra (Gibson, 1977)

Architomy prevalent in 100% of population round the year. Thirteen percent are at prefragmentary stage. 5, 21 and 54% are clonally regenerating at anterior, mid-body and posterior zones, respectively

Sedentary/Tubiculous polychaetes Sabellidae Megalomma cinctum (Yuan, 1992) Bispira brunnea (Davila-Jimenez et al., 2017)

Architomy prevalent in 4–13% of population

Pseudobranchiomma schizogenica (Tovar-Hernandez and Dean, 2014)

Architomy prevalent in 82% of population

Pseudopotamilla reniformis (Kolbasova et al., 2013)

Architomy prevalent in 95% of population from October to March Architomy prevalent in 80% of population

Spionidae Pygospio elegans (Wilson, 1985) Polydora colonia (David and Williams, 2011)

Architomy prevalent in 23 and 13% of population during autumn and winter, respectively

Architomy prevalent in 16–33% of population from September to December Paratomy prevalent in 99.7% of population Paratomy prevalent in 83% of population

Polydorella kamakamai P. stolinifera (Williams, 2004)

Errant oligochaete Lumbriculidae Lumbriculus variegatus (Morgulis, 1907)

Architomy prevalent in > 90% (19% anterior, 42% posterior and 29% anterior + posterior of population) during observation period of July– September

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4.4 Architomy In general, clonal reproduction is faster than that of sexual reproduction. For example, the former involves a fortnight but the latter a month in Enchytraeus japonensis (Yoshida-Noro and Tochinai, 2010). From field and experimental observations, duration from the commencement to the completion of clonal reproduction is reported for a few species. Irrespective of errant and sedentary habit, it requires a short duration of 3–8 days to avoid predation during this sensitive period (Table 4.5). A couple of natural cloners are indicated to require long duration of > 180 days. Understandably, inhabiting a highly unstable habitat, the White Sea with a low average yearly temperature and photoperiod, and consequent low phytoplankton productivity, Pseudopotamilla reniformis accumulates adequate resources throughout spring-summer and clonally reproduces 2–4 progenies during autumnwinter period of 180 days. However, the worm ensures 100% survival of all the two–four progenies. Besides, 70% lengths of its elastic tube strengthened by encrusting fine sand are spread horizontally over the substratum. Within this posterior end of the horizontally spreaded tube, the newly arising clones are maintained (Fig. 4.9). The newly emerged young worm, surrounded by ascidians and others, stands upto a height, at which its visibility is minimal (Kolbasova et al., 2013). The so called schizoparity or schizometamery occurs in a number of polychaetes like Dodecaceria pulchra, Dipolydora armata. In it, the fission involves three fragments including mid-body breaking off singly and cloning a complete progeny (Petersen, 1999). Following schizoparitic fission in Pygospio elegans, 9, 64 and 24% of progenies arising from anterior, mid-body and posterior fragments survive, respectively (Gibson and Harvey, 2000). In Sabellastarte sp, survival of eight fragments is 10, 20, 50 and 75% for the cephalic, mid 5th, mid 6th and posterior (8th) fragments, respectively. All the remaining 2nd to mid 4th fragments fail to survive (Murray et al., 2013). The sequence of natural (e.g. P. reniformis) and artificial (e.g. Pygospio elegans) clonal reproduction is briefly summarized in Table 4.6. Notably, clonal reproduction involves blastema formation in naturally (e.g. Polydora colonia, David and Williams, 2011) and experimentally (e.g. Pygospio elegans, Gibson and Harvey, 2000, Amphipolydora vestalis, Gibson and Paterson, 2003) cloning spionids. Wound healing is noted in many architomics but blastema formation is not reported in any non-spionid species (cf MartinezAcosta and Zoran, 2015). Interestingly, the minimum number of segments required for successful architomic cloning ranges from one seminal segment in Dodecaceria concharum (Martin, 1933) to two–three chaetigers in A. vestalis (Gibson and Paterson, 2003) and to 3.5 segments in Lumbriculus variegatus (Morgulis, 1907). Short Life Span (LS) and high clonal frequency are typical features of some enchytraeids and dorvilleids. Elucidation of their clonal features becomes

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Table 4.5 Progeny production and regeneration duration in clonally reproducing architomic polychaetes and oligochaetes Family

Progeny (no.)

Regeneration (d)

Species, Reference

Errant polychaetes Oweniidae

2

–

Myriochele heeri (Antarctica) (Oliver, 1984) (F)

Chaetopteridae

6

–

Pseudochaetopterus prolifica (see Purscke, 2006)

Amphinomidae

2

240†

Eurythoe complanata (USA) (Kudenov, 1974) (F)

Dorvilleidae

6–8

3

Parougia bermudensis (Akesson and Rice, 1992) (E)

Cirratulidae

2

–

Dodecaceria concharum (USA) (Martin, 1933) (F)

–

D. pulchra (S. Africa) (Gibson, 1977) (F)

Sedentary/tubiculous polychaetes Sabellidae

Spionidae

1

8

Bispira brunnea (Mexico) (Davila-Jimenez et al., 2017) (F)

2–4

180

Pseudopotamilla reniformis (White Sea) (Kolbasova et al., 2013) (F)

3

–

Potamilla torelli (UK) (Watson, 1906) (E)

6–8

–

Pseudobranchiomma schizogenica (Gulf of California) (David and Williams, 2011)

8

> 28

2–3

–

Dipolydora armata (Brazil) (Radashevsky and Nogueria, 2003) (F)

Sabella pavonina, Sabellastarte sp (Murray et al., 2013) (E)

–

8

Polydora colonia (David and Williams, 2011)

2–8

8

Pygospio elegans (Florida/Canada) (Watson, 1985/Gibson and Harvey, 2000) (F/E)

3–6

8

Amphipolydora vestalis (New Zealand) (Gibson and Paterson, 2003) (E) Errant oligochaetes

3–6

6

Enchytraeus variatus (Bouguenec and Giani, 1989) (E)

6–13

5

E. japonensis (Myohara et al., 1999) (E)

7

–

E. bigeminus (Christensen, 1973) (E)

Lumbriculidae

51

–

Lumbriculus variegatus (Morgulis, 1907) (F)

Naididae

6

Enchytraeidae

Allonais paraguayensis (Hyman, 1916) (E)

† for posterior, 90 days for anterior. F and E indicate field and experimental observations, respectively

important from culture point of view (cf Chapter 10). Parougia bermudensis grows to a length of 9.5 mm and consists of ~ 40 segments. A 20-day feeding and accumulation of reserves in coelomocytes is an obligate requirement to initiate schizoparity, yielding three fragments once every third day. Clonal regeneration involves anterior head with < eight segments, mid-body

Asexual Reproduction  137

Table 4.6 Morphogenesis during clonal reproduction in Amphipolydora vestalis (modified from Gibson and Paterson, 2003) and Pseudopotamilla reniformis (modified from Kolbasova et al., 2013) Structure

Morphogenesis on 1st day

2nd day

3rd day

4th day

5th day

6th day

7th day

8th day

9th day

A. vestalis: Anterior blastema Head Palps Mouth Nuchal organ Thorax Gut Chaetigers Neuro chaetigers A. vestalis: Posterior blastema Tail Pygidium New chaetigers Gut Duration (d)

Structure

Figure

Pseudopotamilla reniformis 1–4

Wound healing

6–8

Formation of 2 anterior segments and pygidium

11–30

Development of crown pinnules on radioles and fecal grove

40–60

Further development of crown pinnules on radioles and fecal grove as well as reorganization of parapodia

100

Regeneration completed

fragment with ~ five or ~ nine segments and posterior with a few segments. Those with ~ nine mid-body segments regenerate faster than those with ~ five segments. Further, fragmentation in the mid-body and posterior zone can occur at any segment. However, based on the position and number of segments, the fragments are grouped into five classes. Due to continuous schizoparity, body segments are relocated to a progeny, which may itself produce additional fragments. As a result, some of these segments may be passed from generation to generation with little change and are potentially

138  Reproduction and Development in Annelida

immortal. Akesson and Rice (1992) have carried out a series of experiments and their findings from three experiments are summarized: 1. In all the experiments, the trends run almost parallel to each other in those that are continuously fed and fed following a brief period of starvation. 2. In all the size classes, the number of segments/fragment is decreased totally beyond the 16th and 32nd segments in the mid-body and tail fragments, respectively. This clearly indicates that the fragment consisting of the first 15th to 31st segments determines the number and pattern of segmentation. 3. With advancing age from 0 to 15 months, the fragments holding 13 to 28 segments doubles the number of segments in the cephalic lineages but retains the same number in the non-cephalic lineages. Briefly, posterior fragments undergo senescence but not the mid-body segments and the fragmentation potency increases from the mid-body to the cephalic zone. In the marine paratomic oligochaete Paranais litoralis too, Martinez and Levinton (1992) have found age specific senescence. From the field study, Akesson and Rice (1992) have also noted that in the absence of adequate resource, Parougia bermudensis acquires the dispersal morphism. This migration strategy is more eloborately described in P. litoralis by Nilsson et al. (2000). On depletion of resource, the swimming morph foregoes clonal reproduction and grows by adding 40% more segments and also become thinner. With these morphogenetic changes, the migrant swims faster than the non-migrants. Table 4.5 shows the clonally reproducing enchytraeids, lumbriculid and naidid oligochaetes also exhibit clonal features displayed by P. bermudensis. Of eight architomic clonal Enchytraeus species, more information is available for E. bigeminus and E. variatus. Life span of E. variatus is 81 days, grows to a length of 12 mm and consists of ~ 28 segments. Rearing clonal fragments of E. variatus under optimal conditions, Bouguenec and Giani (1989) have carried out three series of experiments. As they had not identified the fragments/worms, which underwent clonal and/or sexual reproduction, the results reported by them are confusing. An attempt is made to simplify and generalize their findings hereunder: when its fragments are reared under optimal condition in two series of experiments, some unidentified fragments generate immature and mature worms at the rate of 1.4 and 0.7 worms/d through sexual reproduction. Hence, clonally generated progenies are capable of restoring sexual reproduction (cf Ozpolat and Bely, 2015). Remarkably, other fragments have continued clonal reproduction and produced 5.1 fragments per day. The third series of experiment has shown that (a) clonal and asexual reproduction overlaps only for a short duration (Fig. 4.1) and (b) the number of fragments produced decreases from ~ 1.25/day during the initial 20-day period to 0.3/day during the period between 21st and 50th day, indicating the exhaustion of chloragogue to sustain clonal reproduction beyond 20 days. However, a sexually mature worm, which has not earlier undergone clonal reproduction, is able to produce ~ 0.13 cocoon or 1.5 eggs/d. Briefly, clonally produced progeny can restore sexual reproduction

Asexual Reproduction  139

and clonal reproduction; clonal and sexual reproduction may overlap for a brief period (Fig. 4.1) and occurs at the cost of sexual reproduction. E. bigeminus grows to a body length of up to 20 mm and consists of ~ 65 segments. At the age of 7 days, the worm obligately begins to clonally reproduce by dividing into seven fragments. Sexual maturity is attained by 2 and 3 weeks in anterior and other fragments, respectively. The worm is a polyploid. Hence, it is an obligate outbreeder. During the warmer season, the worm usually undergoes repeated clonal reproduction and attains high densities, when clonal reproduction is suppressed. As a result, sexual reproduction occurs mostly during colder months. On introduction of 21 fragments in a culture, the cumulative production of fragments by a single fragment is 276 immature worms. No sexual worm is produced, as clonal reproduction is suppressed only beyond the density of 400 worms in a culture. Interestingly, sexual reproduction is reduced in non-clonal E. albidus and E. irregularis, when they are cultured in combination with E. variatus (Christensen, 1973). Naidids: Of 88 naidid species, in which clonal types are known, 19 and 69 are architomic and paratomic, respectively (Table 4.3). For tropical naidids, publications by Aiyer (1924, 1929) are mostly devoted to taxanomy. Through a series of publications, Hyman (1916, 1938) narrated the factors controlling clonal reproduction in architomic Paranais paraguayensis. Her major findings are listed below: 1. The worm grows to a length of > 35 mm and has > 50 segments. 2. The head fragment holds more number of segments than the trunk and caudal fragments, which consist of 10–25 large and 30–50 smaller segments, respectively. 3. Prior to fragmentation, the worm repeatedly twists its body and after the formation of a constriction, the daughter fragments pull apart. Irrespective of illumination or darkness, it fragments only during night. 4. It can fragment only at temperatures between 15ºC and 27ºC as well as between pH 5 and 8. Carbonate level plays a decisive role in controlling fragmentation. Fragmentation is retarded by potassium cyanide and 1/25,000 mol inhibits it. The fragments hold 15 and 18 segments at 26ºC and 16ºC, respectively. Clearly, elevation in temperature increases the fragmentation frequency and results in smaller daughter fragments. 5. Unlike the enchytraeids, in which smaller immature worms fragment more frequently, the naidid does not fragment until it attains a minimum size of 13 mm. At 15–20 mm and 30–35 mm sizes, it divides into two and three– eight fragments, respectively. 6. At low density, it fragments more frequently and produces smaller daughter fragments. With increasing density, clonal reproduction diminishes, as is the case in enchytraeids. 7. In fresh culture medium, the fragment length is shorter than that in old medium, indicating the inhibitory role of the accumulated excretory and putrefactive products. Earthworms: Of ~ 500 species of earthworms (see Table 1.2), it is experimentally shown that Perionyx excavatus is capable of clonal reproduction (see Fig. 3.1, p 97).

140  Reproduction and Development in Annelida

Figure 4.2 Salmacina dysteri (shown in the window): A. Shows the proportion (%) of total worms as function of position in the pseudocolony. B. Proportion of (%) sexually mature and asexually reproducing worms as function of the duration of observation period. Note growth of the colony is also shown as function of the duration of observation period. C. Proportion of (%) sexually and asexually reproducing worms as function of colony size. D. Proportion of (%) immature and asexually reproducing worms as well as percentage of males and hermaphrodites as function of colony size (compiled and modified from Nishi and Nishihira, 1994).

Pseudocolony: Clonal reproduction by budding, as a distinct form of architomy or fragmentation, occurs in serpulids (Faulkner, 1930) and syllids (Franke, 1999). In Salmacina, buds are always single, terminal and remain attached to the parent stock until all development, apart from size, is completed. Architomy leads to the formation of pseudocolony, in which the calcareous tubes of adjoining worms are attached to each other but without functional connection between them. From the report by Nishi and Nishihira (1994) on the florescent serpulid S. dysteri on the Okinawan coral reef, the following are summarized: 1. The worm grows up to 3 mm length and 0.2 mm width. 2. It consists of a head, three–eight segmented thorax and 10–25 segmented

Asexual Reproduction  141

abdomen. 3. An immature worm is recognized by the presence of a bud at the posterior end of the abdomen and sexually mature one by eggs/embryos in the tube. 4. The approximate sex ratio is 0.3 for females, 0.1 for males and 0.6 for hermaphrodites, which seem to arise from males by addition of the ovarian component. 5. The experimental bisectioning of a pseudocolony reduces but not significantly the proportion of sexually reproducing worms. 6. Sexually reproducing worms occupy the basal and lower middle zones of the colony, while asexually reproducing worms are distributed in the upper middle and peripheral zones (Fig. 4.2A). 7. In a colony with a maximum of 2,000 members, the reverse trends are obtained for sexually and asexually reproducing worms with increasing colony size (Fig. 4.2C) and age (Fig. 4.2B). 8. With decreasing proportion of asexually reproducing worms, that of hermaphrodites increases (Fig. 4.2D). 9. The growth of the colony doubles and redoubles to a maximum of 9 cm (diameter) size during the observation period of 6 months (Fig. 4.2B).

4.5  Naidu’s Monograph In 1986, Brinkhurst had published a monograph on the North American tubificids, which are not known to reproduce asexually (Zattara and Bely, 2016). Thanks to the Zoological Survey of India, Naidu (2005) has brought out a monograph on Aquatic Oligochaetes in the Fauna of India series. Apart from taxonomy and distribution of these oligochaetes in the Indian waters, he has included very valuable information on asexual reproduction of eight aeolosomatid and 58 naidid species. His may prove to be the only literature source on asexual reproduction in the tropical aquatic oligochaetes. As an expert, Naidu has added to the paratomics 3 aeolosomatid and 32 naidid species to the list summarized for the oligochaetes by Zattara and Bely (2016) (Table 4.2). Assembling the scattered information from Naidu (2005), Table 4.7 summarizes clonal reproduction under two groups. According to him, clonal and sexual reproductions occurs concurrently in 18 naidid species and in the remaining 39 species either clonal or sexual reproduction occurs at a given body size or time. Naidu has noted the absence of ovary in all the 39 species but the presence of testes in a dozen species. Group 1. Clonal reproduction alone is reported to occur in seven aeolosomatid and 39 naidid species. In the absence of sexual reproduction and production of the (diapausing) cocoon, clonal reproduction in naidids can be sustained only in such aquatic systems, where water is available round the year. India is characterized by the southwest monsoon sweeping the west coast, and West Bengal and adjoining the northeastern seven sister states. In Kerala, for example, the precipitation lasts for > 150 days and amounts to 400–600 cm/y. Conversely, a larger portion of Andhra Pradesh and Tamil Nadu receive

142  Reproduction and Development in Annelida

Table 4.7 Clonal reproduction in Aeolosomatidae and Naididae, as reported by Naidu (2005). All species undergo paratomy, except a few architomic species, indicated by arch, sex = sexual, clo = clonal, con = concurrent, bz = budding zone, fr = fragmentation, † indicates switching from clonal to sexual, * = gonads disappear, FW = freshwater, BW = brackish water, Mar = marine, Tin = tube inhabiting worm. Species in bold letters and bold letter1 are already known for clonal reproduction from Zattara and Bely (2016) and Lohlein (1999), respectively Species

Reproduction

Occurrence

Aeolosomatidae: Gonads absent (GA) Aeolosoma beddardi A. headleyi A. hemprichi A. niveum A. ternarium A. travancorense A. viride

GA GA, 1–4 bz GA, 2–4 bz Clonal, 3–4 bz GA GA, bz GA, bz

Kerala, Vizianagar, AP Vehar lake, MH Kerala, Kashmir Kerala, Bihar lake, MH Stagnant waters, Sri Lanka Kerala, Nagarkoil (TN) Chandigarh lake

Naididae: Gonads absent (GA) Allonais inaequalis A. paraguayensis A. rayalaseemensis Aulophorus flabelliger A. gravelyi A. gwaliorensis A. hymanae A. indicus A. michaelseni A. moghei A. tonkinensis* Branchiodrilus hortensis B. semperi Chaetogaster limnae bengalensis Chaetogaster limnae limnae Dero cooperi D. indica D. nivea D. palmata D. pectinata D. plumosa D. ravinensis D. sawayai D. zeylandica Nais andhrensis N. pardalis N. variabilis

♀ A, ♂ +, arch GA, fr, arch GA, fr GA, bz GA, fr GA, ♂ +, fr GA, ♂ +, clo, fr GA, fr, bz GA, bz 1 + 1 GA Clonal GA, ♂ +, bz 1 GA GA, bz GA, ♂ +, fr GA, ♂ +, bz 1 GA, ♂ +, bz 1 + 1 GA, bz GA, bz 2 GA, bz GA, bz GA, bz GA, bz GA, ♂ +, bz 1 GA, bz 1 GA, ♂ +, clo, bz GA, bz 1

Kerala, West Bengal Kerala Kerala, TN, AP AP Ennur (TN) Kerala, Tanjavur (TN) Mucous Tin, TN, AP AP ponds Tin, Kerala Nagpur ponds, MP Tin, Kerala, Bhim Tal, Varanasi Kerala, West Bengal, MP In burrows, cosmopolitan West Bengal, Burma Kashmir, Naini Tal Mucous Tin, Kerala, Punjab Kerala, AP Kerala, TN Kerala Kerala Gelatinous Tin, AP Ganga River belt Tin, AP, Nagpur, MP Kerala Ooty (TN), permanent lakes Afghanistan Kerala, Yercaud (TN) Table 4.7 contd. ...

Asexual Reproduction  143

...Table 4.7 contd. Species

Reproduction

Occurrence

Naididae: Gonads absent (GA) Pristinella acuminata P. jenkinae P. menoni P. minuta Pristina aequiseta P. breviseta P. evelinae P. foreli P. proboscidea P. sperberae P. synchites Slavina appendiculata

GA GA, bz GA, bz GA, bz GA, ♂ +, bz GA, ♂ +, bz GA, ♂ +, bz GA, bz GA, bz 1 GA, bz 2 GA, bz 1 GA, ♂ +

Aeolosoma hyalinum Allonais pectinata Aulophorus carteri A. furcatus Chaetogaster cristallinus C. diaphanus† C. diastrophus C. langi1 Haemonais waldvozeli Nais andina† N. barbata N. bretscheri N. communis N. elinguis N. pseudobtusa N. simplex Pristina longiseta longiseta P. macrochaeta Stephensoniana trivandrana Stylaria fossularis†

Sex, fr, 3 zooids Sex, clo, arch Sex, clo, bz Sex, clo, bz Sex, clo, bz, con Sex, clo, bz, con Sex, clo, fr, con Sex, clo, bz, con Sex, clo, bz Sex, clo, con Sex, clo, bz, con Sex, clo, fr, con Sex, clo, fr, con Sex, clo, fr, con Sex, clo, bz Sex, clo Sex, clo Sex, clo, bz Sex, clo, con Sex, clo, bz

North Indian ponds FW, AP Kerala, Yercaud (TN) AP, Rabi river In sponges, Kerala Kerala, Chennai ponds FW, Kerala, Ooty (TN) FW, Dhaka In sponges, coasts of Kerala, W. Bengal FW, Kolkata, AP ponds FW, AP lakes Ganga, AP

Sexual and clonal Vehar lake (MH) In sponge, Kerala, eggs don’t develop Tin, Kerala Mucous Tin, burrowers, Ooty (TN) Kerala, Ooty (TN), West Bengal UP, MP, BW in Sri Lanka AP, Ganga Cosmopolitan in India Kerala, West Bengal MP Chandigarh, Lucknow (UP), W. Bengal Afghanistan Cosmopolitan In sponges, Kerala, West Bengal Afghanistan North Indian ponds Mar, in sponges, Kerala FW, Afghanistan Kerala Kerala, Ooty (TN)

AP = Andhra Pradesh, MH = Maharashtra, MP = Madhya Pradesh, UP = Uttar Pradesh, TN = Tamil Nadu

northwest monsoon of < 100 cm lasting for 30–40 d/y. Not surprisingly, Kerala happens to be the home for > 20 naidid and four aeolosomatid species that reproduce asexually alone. The others inhabit more or less permanent lakes in Kashmir and montane lakes like in Ooty and Yercaud of Tamil Nadu and aquatic bodies associated with live rivers like the Ganga. Notably,

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Branchiodrilus semperi, as a burrowing worm, can survive within the moist soil, when water is not available. Group 2. Concurrent clonal and sexual reproduction: Of 18 species in this group, nine species alone are recorded to undertake clonal and sexual reproduction concurrently. Even among these nine species, asexual reproduction is suspended with a commencement of sexual reproduction in Chaetogaster diaphanus and Nais barbata. For the remaining seven species, no information is provided. Typically, life span of tropical animals is shorter than their counterparts in temperate and arctic zones (e.g. crustaceans, Pandian, 2016). The life span of the tropical aquatic oligochaetes may not be an exception to this dictum. Hence, the life history strategy of the paratomic 18 naidid and one aeolosomatid (Aeolosoma hyalinum) species may fall into one of the following categories: 1. Within the short life span, a relatively longer duration of asexual reproduction is switched to a short duration of sexual reproduction with production of cocoons, which may hatch immediately, when water is present or switch to diapause, when water is not available. These worms may be characterized by semelparity (cf Stylaria lacustris, Schierwater and Hauenschild, 1990). 2. The third and more likely option is to have a very short period, during which clonal reproduction progressively diminishes and sexual reproduction gradually begins to dominate, as in Enchytraeus variatus (see Fig. 4.1). Research inputs are required to identify the option of 19 naidid and one aeolosomatid species, in which clonal and sexual reproduction are reported to co-occur. According to Naidu (2005), India is endowed with seven clonal aeolosomatid species (out of 27) and 58 clonal naidid species (out of 175), although all the three aquatic enchytraeid species listed by him are not cloners. Indeed, these clonal worms are an important resource. Research input in some of these worms can promote aquaculture of these worms and provide gainful employment for the large number of Indians.

4.6 Paratomy The incidence of paratomy is limited to two errant and two sedentary polychaete families as well as two oligochaete families (Tables 4.2, 4.3). In the errant syllids like Procerastea, Autolytus, Myrianida, Pionosyllis and Trypanosyllis, fission and budding are related to epitoky and are discussed in Chapter 5, although many authors have considered them with paratomy. It may be misleading to consider stolon formation as asexual reproduction. Stolonization is intimately associated with sexual reproduction and “stolons are little more than locomotive vessels for gametes” (Franke, 1999). Table 4.8 lists the number of progenies produced by some paratomic species. However, these numbers may be correct for the tubiculous spionids, in which space

Asexual Reproduction  145

Table 4.8 Progeny production and clonally reproducing paratomic polychaetes and oligochaetes Family Spionidae Naididae

Progeny (no.) 1–5

Species, Reference Polydorella kamakamai (Williams, 2004)

2

P. tetrabranchia (Campbell, 1955)

2

Nais communis (Kharin et al., 2006)

2

Stylaria lacustris (Schierwater and Hauenschild, 1990)

3

Pristina longiseta (Kharin et al., 2006)

7

Dero digitata (Drewes and Fourtner, 1991)

perhaps limits the formation of a chain, but may not be correct for the errant oligochaetes, which form chains of daughter and grand-daughter progenies. Secondly, these values have not taken into consideration for the frequency of paratomic fission in them. In Enchytraeus japonensis, each fragment regenerates into a small but complete worm in 4 days, which grows rapidly to divide again in another 10 days (Myohara et al., 1999). Stylaria lacustris splits once every fifth day from April to August/September, i.e. an individual may split > 30 times during this period. A theoretical estimate indicates that a single worm may give rise to a population of 3.4 billion worms (Schierwater and Hauenschild, 1990). Some polychaetes like Polydorella tetrabranchia produce only two offspring/fission once every fortnight (Campbell, 1955). Paratomy involves the division of the body into two distinctive halves followed by regeneration of the missing body parts. Typical of paratomy, the anterior stock and posterior stolon remain attached together, the secondary and tertiary divisions proceed, resulting in the formation of a chain of zooids. Available information indicates the occurrence of division in the stock of sedentary tubiculous spionids but in the stolon of errant oligochaetes like aeolosomatids. In the spionid, Polydorella dawydoffi, which grows to 2 mm length and consists of > 15 segments, the first split occurs on the 13th segment. Hence, the stock inherits the head + 12 segments, while the stolon the pygidium + 4 parental segments (Fig. 4.3A). Preparing for the ensuing second split, the stock has regenerated 12 small segments. The second split occurs on the same 13th segment and the process is repeated during the tertiary split too. Contrastingly, the ambiguously named ‘pygidial budding’ (Herlant-Meewis, 1951) occurs in the errant oligochaete Aeolosoma viride. Falconi et al. (2015) have described the process correctly by stating that the secondary zooids (i.e. the daughter fragments of the first split) in A. viride are positioned posteriorly in inverse order with respect to their age and growth level. The older and more advanced ones are at the posterior end, while the younger and less developed are located immediately behind the parental zooid (Fig. 4.3B). Both of these paratomic types occur in other errant oligochaete taxa, the naidids. In the naidian type, the second fission occurs at exactly on the same

146  Reproduction and Development in Annelida

Figure 4.3 Freehand drawings to show the process of paratomic clonal reproduction in A. Polydorella kamakamai. A2 development of stolon 2 (St2) growth zone, A2 development of first 12 segments of stolon 2 and stolon 2 (St2) growth zone; secondary stolon is also shown by an arrow, A3 paratomic chain of 3 developing individuals prior to paratomic division between stolon 2 and stolon 1. Sites of paratomic divisions are indicated by darker arrows (modified from Williams, 2004). B. Schematic diagram to show paratomic clonal reproduction in Aeolosoma viride. Z3 shows the growth of secondary zooids and its origin in the budding area (filled square) of the main zooid (ZP). The Z3 grows forming new chaetigers produced in anterior direction by the growth area and moves the chain in the posterior direction by the inter position of new secondary zooids. Z3 zooids complete its morphogenesis and separate from the parental chain. ZP is the main zooid; Z1, Z2, Z3, Z4, Z5, Z6, Z7 are all secondary zooids. Z1.P + Z1.1, Z2.P + Z2.1 and Z3.P + Z3.1 represent the first, second and third filial chains, respectively (freehand drawing from Falconi et al., 2015).

segment number, as the first one has been. But, it occurs one segment behind the first one, designated as n-1 in the stylarian type. As a result, successive fissions are named as n-2, n-3 and so on, and each new individual contains one segment of the original parent. In Stylaria, the limit is n-7, after that the anterior stock elongates, as in normal posterior growth until the total number of segments exceeds 40, when a new fission zone is intercalated in the middle region (Berrill, 1952). In Pristina, the limit is n-12 (Hempelmann, 1923). Nervous system and/or neurosecretion are known to play a dominant role in regeneration of clonal reproduction. The publications by Drewes and Fourtner (1990, 1991) on reorganization of segments and transformation of the giant nerve fiber sensory fields in the context of escape reflexes indicate a series of changes in the segment potential of Lumbriculus variegatus and Dero digitata. Following fission(s), a small and constant number of head segments are regenerated by the body segments, irrespective of their original axial origin. Thus, the original segment immediately behind the head becomes re-specified to match their altered position. The following morphallactic

Asexual Reproduction  147

Figure 4.4 Clonal reproduction in Aeolosoma viride. Cumulative and daily production (thin line) of filial chains during the life time (modified and redrawn from Falconi et al., 2015).

changes include the rearrangement of segmental gradients in the giant nerve fibers also and progresses antero-posteriorly. Eventually, the medial and lateral giant nerve fiber sensory fields and conduction properties are also re-established. Aeolosoma viride may serve as an example for the reproductive potential of asexual reproduction. The Products: 1. According to Falconi et al. (2015), the pygidial budding results in the formation of a chain composed of two– five zooids, designated as zA, zB, zC, zD and zE, based on their position in the antero-posterior axis of the body (Fig. 4.3B). 2. The worm produces 51 progenies during its Life Span (LS) of 64–66 days. The process: 3A. The proportion of a parental zooid is ll, 45, 40 and 5% for the 5th, 4th, 3rd and 2nd zooids, respectively. 3B. The number of chaetigers/zooid is decreased from 8 in zA to 5 in zD after deep reductions to 1 and 2 in zB and zC, respectively. 4. In this worm, a single budding area is located in the sub-terminal part of the posterior end of the parental zooid. In this area, a histochemically recognizable cell mass of the coelom produces the chaetiger. It is from this chaetiger, the secondary zooids arise at the rate of one every 24 hours. In the secondary zooids also, the same fragmentation process is repeated from the same chaetiger. 5. The maturation time, i.e. the interval between origin of a zooid and its separation from the chain is 80 hours. 6. The last zooid in a chain that has completed growth and cephalic differentiation separate itself from the chain at the rate of 1 at every 22 hours. The dynamics: 7. The production of cumulative number of zooids produced as a function of time

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shows a logistic trend with a raising slope lasting for 20 days or 30% of LS and a gradual asymptotic duration of 45 days or 70% of LS (Fig. 4.4). Senescence: 8. Correspondingly, the number of progenies produced is dropped to zero at the age of 60th day after an initial peak to 2.2 offspring/d. 9. From their observations on 1,559 fissions in the formation of G1 zooids, Falconi et al. (2006) have found that the worm attains senescence at ~ the age of the 60th day of its life. Notably, the negative effect is imprinted on the progenies produced during the senescent age. Not only age but also density and the consequent non-availability of food can induce irreversible effect on reproduction in sedimentivorous worms like Paranais leidyi. The worms that have been cultured at high densities (16–18 worms/10 cm2) without renewal of fresh sediments crashed its asexual reproductive potency to zero level. The F1 progenies produced during the crashing period, even when optimally fed, suffer 100% mortality and reduction in the proportion of clonal worms to 11%, in comparison to 25 and 58%, respectively in optimally fed worms. Hence, the effect of nonavailability of food in the parental worm is also imprinted in F1 generation of the worm.

4.7  Restoration of Sexual Reproduction Germ cells constitute a key cell type. Being the sources of gametes, they are required to manifest sexual reproduction. In some polychaetes and oligochaetes, clonal reproduction has evolved independently at multiple numbers of times. Typically, a few of these species undergo many rounds of clonal generations (e.g. Pristina leidyi, Ozpolat and Bely, 2015) but cloning is terminated in semelparous oligochaetes (e.g. Stylaria lacustris, Schierwater and Hauenschild, 1990) or interspersed by short bouts of sexual reproduction (e.g. Nais barbata). These clonal annelids re-establish by transmission of the germ line post-embryonically. Homologs of the germline (piwi, vasa and nanos) and other multi-potent somatic stem cell genes are reported to express in the gonads as well as in proliferative tissues like Segment Addition Zone (SAZ) (e.g. Platynereis dumerilii, Gazave et al., 2013), regenerative blastema (e.g. Nereis virens, Kozin and Kostyuchenko, 2015) and fission zone (e.g. Enchytraeus japonensis, Tadokoro et al., 2006). Hence, both Primordial Germ Cells (PGCs) and somatic stem cells are characterized by the expression of the similar set of genes. Besides, PGCs and Mesodermal Posterior Growth Zone (MPGZ) cells or somatic stem cells are indistinguishable in morphology and expressed the germline markers vaso, nanos and piwi. Recently, the PGCs of P. dumerilii have been found to incorporate the proliferation marker 5-ethyl2’deoxyuridine (EdU), a marker for proliferation shortly before gastrulation, which coincides with the emergence of four small blastomeres from the mid-

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Figure 4.5 Distribution of PRlle-piwi1 positive ventral cells (PPVCs) along the body of Pristina leidyi. Asterisk indicates the axial position of individuals PPVCs in the worm. Arabic numbers indicate the mean number of PPVCs per segment in the six body regions along the body (modified and redrawn from Ozpolat and Bely, 2015).

blast lineage. Hence, the so called ‘secondary mesoblast cells’ constitute the definitive PGCs in P. dumerilii. In contrast, the cells of the MPGZ incorporate EdU only from the pre-trochophore stage onwards (Rebscher et al., 2012). Using the EdU pulse labeling technique, it has been possible to trace the independent emergence of PGCs from the vasa, piwi and PL10 expressing MPGZ even earlier than that of somatic stem cells. Interestingly, the Oogonial Stem Cells (OSCs) only express piwi and vasa in embryos of the leech Helobdella robusta, while Spermatogonial Stem Cells (SSCs) only express nanos (Cho et al., 2014). Regarding the restoration of germline lineage, two publications are considered. In Enchytraeus japonensis, the gonad can regenerate from any body segment produced by fission during clonal reproduction. Using homolog piwi gene (Ej-piwi) as a marker, Tadokoro et al. (2006) have found that Ej-piwi are distributed widely in the body as single cells. These cells serve as a reservoir of germ cell precursors and migrate into the regenerating tissue, where they eventually form the gonadal primordium and give rise to germ cells upon sexualization. These germline stem cells are distinct and differ from the somatic lineage arising from the neoblasts. In P. leidyi, PRlle-piwi1 is expressed in isolated spindle-shaped cells on the dorsal surface of the ventral nerve cord (Fig. 4.5). Following fission, the number and configuration of Piwi-Positive Ventral Cells (PPVC) are transmitted across asexual generation through migration along the ventral nerve cord. As a result, asexually reproducing P. leidyi expresses PRlle-piwi1. Amazingly, the clonal strain of P. leidyi, in which Ozpolat and Bely (2015) have accomplished their findings, has been reproducing asexually alone for over 60 years and has undergone clonal reproductive cycle for 1,000–3,000 generations.

4.8  Clonal Stem Cells Using available information in the representative species belonging to 10 families of polychaetes and oligochaetes, an attempt has been made to trace

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the possible direction of clonal reproduction. The direction may provide a clue for the anlagen of the multipotent stem cells that manifest clonal reproduction (Table 4.9). In oligochaetes, it is the multipotent neoblasts that manifest clonal reproduction. At this juncture, two facts have to be noted. 1. Fragmentation can occur between any two post-cephalic segments in Enchytraeus japonensis (Myohara, 2012). However, the loss of neoblasts by some posterior segments by Lumbriculus variegatus limits the fragmentation from the trunk to the anterior abdominal segments (Morgulis, 1907). The extensive loss of neoblasts limits the fragmentation to the mid-body segments alone in Stylaria lacustris (Chu and Pai, 1944). 2. Paratomy may involve either fragmentation or budding (see Table 4.9). The number of the budding zone is limited to only one in many oligochaetes (e.g. Aeolosoma viride, Falconi et al., 2015) or rarely to two–four in a few naidids (Table 4.9). Consequent to these differences in the distribution pattern of neoblasts in segments, the direction of clonal reproduction is altered. With the budding zone limited to a single chaetiger at the posterior end of the parental zooid, the direction is from posterior to anterior in A. viride. Conversely, it is antero-posterior in S. lacustris, as the site of fragmentation is located in the stock. In polychaetes, the clonal direction varies and is also complicated. Firstly, there are no equivalents of neoblasts. Secondly, some polychaetes are sedentary and tubiculous, while others are errants. Thirdly, within tubiculous polychaetes, cloning may be by architomic fragmentation, as in sabellids or paratomic budding, as in serpulids. Surprisingly, the direction originates from the posterior end in all the thus far investigated sabellids (Table 4.10). In Sabella pavonina, though all the eight fragments commence clonal regeneration, their survival increases from the posterior end. The posterior origin of clonal direction is also true for the paratomic serpulids (Faulkner, 1930), as represented by Salmacina dysteri. Understandably, the tubiculous sabellids and serpulids hold the clonal stem cells at the posterior segments in the depth of the tube, as the crown and thorax are subjected to more intense predation. Incidentally, the neurosecretion required for regeneration also arises from the ventral ganglia of posterior segments in the sedentary polychaetes (see p 111). With fission limited to the stock in the paratomic tubiculous Polydorella dawydoffi, the clonal direction is unusually antero-posterior (Table 4.9). Among the errant polychaetes too, the direction varies. In the dorvilleid Parougia bermudensis, the direction originates also from posterior but more frequent from the mid-body segments (Table 4.9). Understandably, it is bidirectional in the cirratulid Dodecaceria concharum and the syllid Procerastea halleziana, as they possess one or two seminal segments in the mid-body, from which anterior and posterior segments are developed. In Pygospio elegans, the mid-body fragment develops both anterior and posterior; however, the anterior develops the posterior but the posterior may not. Neither the

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Table 4.9 Clonal direction in polychaetes and oligochaetes. Arrows indicate the clonal direction and Arabic numbers indicate the position of amputation Family

Habit/Type

Sabellidae Bispira brunnea

Tubiculous/architomy

Pseudobranchiomma schizogenica

Tubiculous/architomy

Pseudopotamilla reniformis

Tubiculous/architomy

Sabellastarte sp

Tubiculous/architomy

Sabella pavonina

Tubiculous/architomy

Potamilla torelli

Tubiculous/architomy

Serpulidae Salmacina dysteri

Tubiculous/paratomy

Aeolosomatidae Aeolosoma viride

Errant/paratomy

Dorvilleidae Parougia bermudensis

Errant/architomy

Spionidae Polydora colonia

Tubiculous/architomy

Cirratulidae Dodecaceria concharum

Errant/architomy

Syllidae Procerastea halleziana

Errant/architomy

Spionidae Pygospio elegans

Tubiculous/architomy

Enchytraeidae Enchytraeus japonensis

Errant/paratomy

Lumbriculidae Lumbriculus variegatus

Errant/paratomy

Naididae Stylaria lacustris

Errant/paratomy

Spionidae Polydorella dawydoffi

Tubiculous/paratomy

Direction

13th seg

12%

64%

24%

70 seg

120 seg

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Table 4.10 Clonal direction and potential anlage of clonal stem cells (vertical bars) in errant and sedentary polychaetes (for details see Table 4.9) Family

Clonal Type

Clonal Direction

Errant polychaetes Dorvilleidae

Architomy

Syllidae

Architomy

Cirratulidae

Architomy Sedentary/Tubiculous polychaetes

Sabellidae

Architomy

Serpulidae

Paratomy

Spionidae

Architomy



Architomy Paratomy

errant and sedentary habit nor architomy and paratomy in spionids seem to play a determining role in the clonal direction. In the absence of neoblasts, more than one type of clonal stem cells may be present at different anlage of polychaetes.

5 Epitoky

Introduction Epitoky is a spectacular phenomenon, unique to a few errant polychaete species and is not known to occur in any other aquatic invertebrate taxa. Since 1868, this phenomenon has attracted the attention of a large number of zoologists. The transformation (metamorphosis) from benthic atokous form to a brief epitokous pelagic existence devoted to mating involves many morphological, physiological and behavioral modifications. It involves the following structural modifications: A. Morphology (i) body length and segment number (e.g. Raricirrus variegatus, Dean, 1995), (ii) enlarged eyes, both in cell number and volume (e.g. Odontosyllis polycera, Daly, 1975), (iii) broadened vascularized biramous parapodia with formation of spatulate natatory chaetae (e.g. O. polycera, Daly, 1975) and B. Anatomy (iv) atrophy of the gut, (v) histolysis of body wall to provide resources for gametogenesis (e.g. O. polycera, Daly, 1975), structural changes in musculature involving reduction in longitudinal muscle (cf Samuel et al., 2012, p 108) and reorganization of peripheral muscle cells in each segment and elaboration of structures for the release of gametes (O. polycera, Daly, 1975); these changes are radical in Autolytus but moderate in Odontosyllis. C. The physiological changes include adjustments in metabolic pathways facilitating higher muscular activity to sustain vertical swimming capacity (e.g. Nereis virens, Chatelain et al., 2008). D. Behavioral changes comprise of response to luminescence (e.g. Odontosyllis enopla, Fischer and Fischer, 1995) and pheromones (e.g. Perinereis dumerilii, Hardege et al., 1998) to synchronize swarming and spawning with precise timing guided by the lunar cycle (e.g. Odontosyllis luminosa, Gaston and Hall, 2000). As epigamic epitokes die soon after spawning, they lose 75–79% of available energy allocated for gametogenesis (e.g. Nereis pelagica, Olive et al., 1984, Perinereis cultrifera, Cassai and Prevedelli, 1998a). Not surprisingly, these epitokes release a few million eggs (e.g. Glycera dibranchiata, Creaser, 1973).

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5.1  Types and Characteristics Essentially, there are two major types of epitoky: epigamy and schizogamy. In epigamy, the reckless mating act commences with a burst of sustained vertical swimming activity, lifting the worm to the pelagic zone, where at the climax, a huge cloud of gametes is discharged and is terminated by death due to total exhaustion. For instance, it takes an entire life span of 4 years in Glycera dibranchiata to prepare for the event (Fig. 5.1). In fact, irreversible transition from somatic growth to reproductive development occurs several months prior to breeding. Briefly, all the epigamic polychaetes are semelparous. In schizogamy, only a part (usually the stolon) of the body is transformed into an epitokous sexual stage (Fig. 5.1). Amassed with a load of gametes, equipped with a stolonial head and other sensory structures like the eyes, the stolon breaks off from the atokous benthic worm and migrates vertically for a brief existence in pelagic zone. An epitokous stolon lacks a mouth and a pharynx and its independent life exclusively is devoted to mating, followed by eventual death, i.e. the stolons are semelparous. The unchanged benthic parental stock, however, survives, continues to feed, regenerates the lost segments and then reproduces again (Franke, 1999). In these worms, schizogamy has restored iteroparity from the reckless semelparity.

Figure 5.1 Schematic representation of semelparous epigamic epitoky in non-syllid errant polychaetes and iteroparous schizogamous epitoky in errant syllid polychaetes. 1. Atokous form, 2. Sexual maturation, 3. Epitokous form, 4. Stolonic epitokous form, 5. Spawned eggs by swarmed epitokous females, 6. Trochophore larvae, 7. Regenerated parental atokous stock (modified from Franke, 1999).

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5.2 Epigamy With taxonomy being in a fluid status and description of life history limited to < 3% of polychaetes (Giangrande, 1997) and the number of description increasing at three species/y (see p 41), it is to be expected that reviewers have also limited the description of epigamy citing a few examples alone. Secondly, a few reports on the incidence of epitoky are not reliable. Despite their admirable effort to assemble information on dispersal of polychaetes, Carson and Hentschel (2006) have wrongly indicated the incidence of epitoky in seven glycerid species. However, when checked with Strathmann (1987) and Shanks (2001), the cited information has not been found. With these constraints, the first attempt has been made to list the incidence of epigamy and schizogamy in Table 5.1 and Table 5.3, respectively. Incidence of epigamy is limited to 61 + species in a dozen errant polychaete families. Nereididae includes the maximum incidence in 19 species. Rightly, the epitokous form is named as ‘heteronereis’. Epitoky occurs in these polychaetes from the Arctic to Antarctic (e.g. Capitella capitata, Table 5.1). Notably, half a dozen nereidid species are reported from estuaries (e.g. Hediste japonica) and rivers (e.g. Dendronereis aestuarina). In epigamics, especially in the nereidid, sex ratio is skewed in favor of males, except in Perinereis vancaurica tetradentata and Eunice siciliensis. Understandably, the former broods embryos and larvae in gelatinous mass (Arias et al., 2012). In nereidids, the ratio ranges from 1 ♀ : 1 ♂ in the pair-forming Nereis limbata to 1 ♀ : 3 ♂ in the riverine H. diadroma and D. aestuarina (Table 5.2). Hence, it is likely that the excess number of males ensures the highest Fertilization Success (FS). Regrettably, no information on FS is yet available for any epigamic epitokous species or for that matter, any broadcast spawning non-epitokous polychaetes. Simple plankton collection of eggs prior to the completion of swarming can readily provide the desired information (cf Pandian, 2010). Incidentally, 53% polychaetes are broadcast spawners (p 38) and, external fertilization occurs in 53% of polychaetes (p 44). Hence, it is difficult to justify whether epitoky has achieved a higher FS than non-epitokous broadcast spawning polychaetes or epitoky is simply a reckless mating act. Incidentally, some hesionids like Kefersteinia cirrata swarm and spawn for 2 days during June–July in the UK waters (Olive and Pillai, 1983). In fact, Fage and Legendre (1927) have listed K. cirrata, Oxydromus propinquius, Ophiodromus flexuosus and Podarke pallida as epitokes. The epitokes of a few nereidids like Hediste spp and Odontosyllis enopla display neither the striking structural epigamous modifications nor look like heteronereis. However, these hesionids are not listed in Table 5.1, as they do not display epigamous modifications and are not semelparous. Likewise, the nephtyid

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Table 5.1 Incidence of epigamic epitoky in errant polychaetes Aerocirridae Flabelligela macrochaeta (Mexican Pacific, Salazar-Vellajo and Londano-Mesa, 2004) Amphinomidae Eurythoe complanata (Bay of Bengal, Fauvell, 2010) E. parvecarunculata (Bay of Bengal, Fauvell, 2010) Aphroditidae Drieschia pelagica (Bay of Bengal, Fauvell, 2010) Capitellidae Capitella capitata (Arctic, Antarctic, Mediterranean, Lopez-Jamar et al., 1986) Cirratulidae Aphelochaeta glandularia (USA, see Petersen, 1999) A. monilaris (USA, see Petersen, 1999) Caulleriella viridis (French coast, see Petersen, 1999) Cirratulus cirratus (Gibson, 1981) C. incertus (UK, see Petersen, 1999) Dodecaceria caulleryi (Berrill, 1952) D. concharum (Danish waters, see Petersen, 1999) D. fimbriata (Northeast USA, Gibson, 1979) Ctenodrilidae Ctenodrilus serratus (Indo-Pacific, Atlantic, Harms, 1993) Monticellina heterochaeta (Mediterranean, Martin and Gil, 2010) Raricirrus variabilis (Virgin Islands, 17º N, Dean, 1995) Tharyx perbranchiata (Mexican, Pacific, Salazar-Vellajo and Londano-Mesa, 2004) Eunicidae Eunice afra (Tropical, Bisby et al., 2005) E. fucata (Atlantic, Mayer, 1902) E. schizobranchia (Subtropical, Martin and Gil, 2010) E. schemacephala (Red Sea, Mexico, Wehe and Fiege, 2002) E. siciliensis (Mediterranean, Hofmann, 1974) E. torquata (Tropical, Fishelson, 1971) E. viridis (Pacific, Hauenschild et al., 1968) Glyceridae Glycera alba (Northeast Atlantic, Mediterranean, Gusso et al., 2001) G. americana (Indo-Pacific, Western Atlantic, Fish Forum) G. capitata (Belgium coast, Degraer et al., 2006) G. dibranchiata (Maine, USA, Creaser, 1973) G. macrobranchia G. gigantea (Pacific, cosmopolitan, Gibbs, 1978) G. oxycephala (Tropical, Salazar-Vallejo, 1996) G. tenuis (Tropical, Pacific, Salazar-Vellajo and Londano-Mesa, 2004) Table 5.1 contd. ...

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...Table 5.1 contd. Nereididae Nereis succinea (Brazil, Aguiar and Santos, 2017) N. virens (Canada, Chatelain et al., 2008) (Andaman Island, Muruganatham et al., 2015) Dendronereis aestuarina (Freshwater in the southwest coast of India, Jayachandran et al., 2015) Hediste diadroma (Japan, Hanafiah et al., 2006) H. japonica (Japan, Sato, 1999) H. osawai (Japan, Hanafiah et al., 2006) H. oxypoda sensu (Japan, Hanafiah et al., 2006) Marphysa disjuncta (New Zealand, Read, 2004) N. falcaria (Australia, Read, 1974) N. fucata (Atlantic, Gilpin-Brown, 1959) N. grubei (Schroeder, 1967) N. japonica (Japan, see Fischer, 1999) N. limbata (Hauenschild and Hauenschild, 1951) N. pelagica (Tropical, Salazar-Vallejo, 1996) Perinereis cultrifera (Cassai and Prevedelli, 1998) P. nuntia brevicirrus (see Gilpin-Brown, 1959) P. vancaurica tetradentata (Mediterranean, Arias et al., 2012) Platynereis dumerilii (Garcia-Alonso et al., 2013) Tylorhynchus heterochaetus (Japan, Koya et al., 2003) Ophelidae Euzonus flabelligerus (Baltic, Rower, 2010) Phyllodocidae Mystides caeca (Mediterrean, Martin and Gil, 2010) Nereiphylla castanea (Indo-Pacific, Salazar-Vellajo and Londano-Mesa, 2004) Paranaitis polynoides (Western central Atlantic, Salazar-Vellajo and Londano-Mesa, 2004) Phyllodoce cuspidata (Northeast Pacific, Macdonald et al., 2010) P. groenlandica (East Pacific, Altantic, Salazar-Vellajo and Londano-Mesa, 2004) P. hartmanae (Pacific, Macdonald et al., 2010) P. longipes (Tropical, Vittor, 2002) Protomystides confusa (Tropical, Western Atlantic, Salazar-Vallajo, 1996) Pterocirrus foliosus (Central American Sea, Salazar-Vallajo, 1996) Syllidae Autolytus alexandri (Franke, 1999)

Nephtys caeca with a straightforward annual reproductive cycle is also not included, as it is polytelic (Olive, 1978). But both Hediste spp and O. enopla are listed, as the former is semelparous and the latter displays luminescence, a typical characteristic of epitokous genus Odontosyllis. Notably, many authors have reported either a fraction or an entire population undergoing reversible epigamy. For example, Creaser (1973) has observed a few partially milted males of Glycera dibranchiata returning to the bottom and burrows. In

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Table 5.2 Sex ratio of some epigamic epitokes Species

Sex Ratio ♀ : ♂

Reference

Nereidae Nereis limbata

1.0 : 1.0

Hauenschild and Hauenschild (1951)

N. fucata

1.0 : 1.0

Gilpin-Brown (1959)

N. brevicornis

1.0 : 1.25

See Gilpin-Brown (1959)

N. japonica

1.0 : 2.10

Hanafiah et al. (2006)

N. diadroma

1.0 : 3.0

Hanafiah et al. (2006)

Dendronereis aestuarina

1.0 : 3.0

Jayachandran et al. (2015)

Perinereis vancaurica tetradentata (brooder)

1.0 : 0.1

Arias et al. (2012)

Glyceridae Glycera dibranchiata

1.0 : 1.24

Creaser (1973)

Eunicidae Eunice siciliensis

1.8 : 1.0

Hofmann (1974)

O. enopla, a reversible epigamic epitoky is displayed. This Bermudan fireworm does not undergo structural modifications except for the large eyes and swimming chaetae; all the post-swarming worms return to a benthic life. In laboratory reared worms, the post-swarming females are filled with a new set of oocytes 4 months after the swarming (Fischer and Fischer, 1995). Rearing the anterior fragment of the epitokous Eunice siciliensis, Hofmann (1974) has observed the following sequence of development: (i) within 10–14 days, all the anterior fragments begin caudal regeneration. (ii) more than 50 segments with parapodia are developed by the 30th day. (iii) between 10th and 16th month, almost all of them have developed gonadal sacs and germ cells. Briefly, the anterior part of the epitoky—when reared under optimal conditions—can survive, completely regenerate genital segments and produce another epitoky. In the recipient mature (non-epitokous) N. diversicolor, implantation of cerebral ganglia from the immature donor is reported to induce repeated gametogenic cycles but not spawning. Still, the publication by Golding and Yuwono (1994) has indicated that the trait of iteroparous gametogenic cycling is retained in the epitokous nereidids. Hence, the regenerative potency of the anterior fragment is inherent but for its initiation, promotion of gametogenesis and epitokous development, a gonadotrophic neurosecretion originating from the prostomium is required. Apparently, the genes responsible for iteroparity in these epigamics are indeed retained but are not expressed and results in an expensive semelparity.

Epitoky  159

Table 5.3 Incidence of schizogamic epitoky in syllid polychaetes Autolytus brachycephalus (Schiedges, 1980) A. charcoti A. edwardsii (Okada, 1935) A. magnus A. prolifer (see Franke, 1999) A. purpureimaculata (Okada, 1933) Brania pusilla (Temperate, Mediterranean, Harms, 1993) Eusyllis blomstrandi (Indo-Pacific, Atlantic, Arctic, Harms, 1993) Exogone hebes (NewFoundland, Pocklington and Hutcheson, 1983) Exogone naidina (Indian ocean, Atlantic, Arctic, Harms, 1993) Haplosyllis spongicola (Indo-Pacific, Mediterranean, Bisby et al., 2000) Haplosyllides floridana (see Franke, 1999) Grubeosyllis clavata (Tropical, Circumglobal, MarineSpecies.org) Myrianida pinnigera (Subtropical, Mediterranean, Martin and Gil, 2010) Odontosyllis enopla (Bermuda, Fischer and Fischer, 1995) O. hyalina (Indonesia, Lummel, 1932) O. luminosa (Belize, 16˚.5’ N, Gaston and Hall, 2000) O. phosphorea (Bermuda, Tsuji and Hill, 1983) O. polycera (New Zealand, Daly, 1975) O. undecimdonta (Japan, Inoue et al., 1993) Pionosyllis lamelligera (see Franke, 1999) P. neapolitana (see Franke, 1999) P. pulligera (see Franke, 1999) P. procera (see Franke, 1999) Proceraea cornuta (North Atlantic, North Pacific, Nygren, 1999) P. okadai (North America, Calif Acad Sci, 2015) P. picta (Subtropical, Martin and Gil, 2010) Sphaerosyllis hystrix (Tropical, Mediterranean, Wehe and Fiege, 2002) Streptosyllis verrilli (see Franke, 1999) S. websteri (see Franke, 1999) Syllides japonica (see Franke, 1999) Syllis amica (Ruppert et al., 2004) S. gracilis (Indo-Pacific, Mediterranean, Harms, 1993) S. ramosa (Phillipines, Wikipedia) S. vittata (Central American Mediterranean, Salazar-Vellajo, 1996) Trypanosyllis asterobia (Okada, 1933, 1937) T. coeliaca (Tropical, Salazar-Vallajo, 1996) T. crosslandi (Bay of Bengal, Fauvel, 2010) T. gemmipara (Tropical, Salazar-Vellajo and Londano-Mesa, 2004) T. ingens (Northeast Pacific, Canada, Lamb et al., 2011) T. zebra (Mediterranean, Western Pacific, Salazar-Vellajo and Londano-Mesa, 2004) Typosyllis prolifera (Atlantic, Mediterranean, Wikipedia) T. hyalina (see Franke, 1999) T. pulchra (Tropical, Mexican Pacific, Salazar-Vellajo and Londano-Mesa, 2004) T. variegata (Tropical, Mexican Pacific, Salazar-Vellajo and Londano-Mesa, 2004)

160  Reproduction and Development in Annelida

5.3 Schizogamy The speciose (700 species) syllids display an incredible diversity in the reproductive phenomenon. Almost all the reported incidences of schizogamy occur in 45 + syllid species (Table 5.3); hence schizogamics make up only 6.4% of Syllidae. The schizogamic syllids reproduce by means of sexual stolons. The stock can generate one (e.g. Streptosyllis verrilli, iteroparous?) to 15 successive stolons (e.g. Typosyllis prolifera, see Franke, 1999). In Autolytus prolifer a single female can produce 1.4 (sacconereis) stolons/mo (Hauenschild, 1953). Indeed, the syllids have discovered stolonization as the most successful reproductive mode and have elaborated them into fascinating diverse types. Accordingly, schizogamy is divided into two subtypes: scissiparity, where the existing segments in the stock are transformed into a stolon and is followed by fragmentation. The fission plane is a fixed one, for instance, the 14th segment in Proceraea picta. It is further divided into paratomic stolonization, (e.g. T. prolifera) and architomic stolonization (e.g. T. hyalina and T. variegata, see Franke, 1999, Polycera cornuta [Fig. 5.2A], Myrianida pachycera [Fig. 5.2B]). The gemmiparous subtype is characterized by budding, in which the segments, destined from the very beginning to form a stolon, rapidly proliferate. Successive terminal buddings lead to the formation of stolon chains. For example, Trypanosyllis asterobia forms multiple collateral budding with a large number of successive posterior stolons (Fig. 5.2G). In T. gemmipara, a bundle of stolons are simultaneously formed within a limited proliferation area near the posterior end of the stock (Fig. 5.2H). A single stolon is formed/mo in T. prolifera and the number of regenerated stock segments is in a sort of temperature-dependent monthly rhythm (Franke, 1985). In Syllis ramosa, collateral budding results in a complex stock consisting of numerous branches extending through the canal system of the host sponge. Special branches bear a single stolon at their tip (Fig. 5.2E, F). Within the syllids, the subfamily Autolinae is the most complex. On the basis of sexual dimorphism, especially cephalic appendages, the female and male stolons are named as sacconereis and polybostrichos, respectively. Procerastea halleziana undergoes rapid multiplication by a process of repeated fragmentations followed by regeneration of anterior and posterior ends in each fragment. The first fragmentation occurs between the 7th and 8th chaetigerous segments, resulting in the head and seven segments forming a fragment. This is followed by 3 fragmentations, resulting in 3 ramets each of with 2 segments, and another 3 ramets each with 3 segments, behind which 4 or 5 fragmentations per se with 4 fragments each followed by divisions per se again 2 fragments each with 3 segments up to pygidium. The mode of fragmentation is expressed in the following formula in species belonging to five different genera.

Epitoky  161

Procerastea: Hd 7 + 2 + 2 + 2 + 3 + 3 + 3 + 4 + 4 + 4 + 4 (+ 4) + 3 + 3…..Py Autolytus: (Hd 7 + 2) + 2 + 2 + 3 + 3 + 3 + 4 + 4 + 4 + 4…………….Py Pionosyllis: (Hd 7 + 2) + 2 + 2 + 3 + 3 + 3 + 4 + 4 + 4 + 4………Py Myrianida: (Hd 7 + 2 + 2 + 2) + 3 + 3 + 3 + 4 + 4 + 4 + 4 + 3 + 3 + 4 + 4 + 4 + 4 + 4 + 4 + 3 + 3..Py Trypanosyllis: Hd (7 + 2 + 2 + 2 + 3 + 3 + 3 + 4) + 4 + 4 + 4 + 3 + 3 + 4 + 4 + 4 + 3 + 3 + 4 + 4 + 4 + 4 + 3 + 3 + 3….Py Hd = 7 = Head segments, Py = Pygidial segment, Head segments + Anterior region within brackets are usually not fragmented (Allen, 1923, Berrill, 1952).

Figure 5.2 Some patterns of schizogamic epitoky in syllids. A. Scissiparous vertical Polycera cornuta, B. Gemmiparously budding Myrianida pachycera, C. Autolytus prolifer, D. Scissiparous laterally budding Exogone rubescens, E and F. Show individual and highly branching Syllis ramosa, G. Multiple collateral budding Trypanosyllis asterobia and H. Simultaneously bundled stolons at the posterior end of T. gemmipara (all are freehand drawings from Franke, 1999 and others).

162  Reproduction and Development in Annelida

Many hermaphroditic schizogamic syllids reproduce through stolons. As sex change occurs in the stolons of the hermaphrodites like Syllis vittata (Durchon, 1975), it is possible that germ cells are also transmitted through successive stolons. However, it is not known whether this sort of transmission also occurs in gonochoric epitokous syllids. In Autolytus edwardsii, fertile eggs produced in the terminal stock segments are passed on to the stolons, where they undergo further development with a support of nurse eggs produced in abortive stolonial ovaries (see Franke, 1999). As egg development is restricted to vitellogenesis alone in the stolon, research inputs are required to know whether the stolons of gonochoric have also received germ cells from the stock. Schiedges (1979) has successfully hybridized Autolytus prolifer and A. brachycephalus by feeding them on laboratory-reared hydrozoid Eirene viridis. The life span of the hybrid is ~ 550 days. A detached sacconereis holds 2, 8–16 and 0–2 segments in its anterior, swimming mid-body and caudal zones, respectively, in comparison to 2, 4–20 and 0–1 segments in polybostrichos. A swarming polybostrichos commences the encircling dance around the sacconereis and begins to milt. It can inseminate eggs of several sacconereis prior to death. The stolonization potential increases from five numbers on the 8th day to 45 on the 180th day and is followed by gradual decrease to 5–10 stolons up to 540th day. Apparently, the stolonization process also undergoes age specific senescence (cf Martinez and Levinton, 1992, see also p 138), perhaps due to the reduction of juvenile hormone, the nereidine (see Golding, 1967). Besides questioning Hamond’s (1969) idea of monophagy, Schiedges (1979) has also found that the hybrid produces scissiparous stolons initially but switches to gemmiparous stolons subsequently.

5.4  Vertical Migration Understandably, the spectacular epitoky has attracted the attention of a large number of zoologists over long years. But only a few of them have cared to report the depth, from which vertical migration is undertaken by the epitokes to reach the surface waters, where swarming and mating occur. For the first time, Table 5.4 lists almost all available information on the depth, from which the migration is undertaken. Of 14 epigamic species, 11 of them undertake the migration over a distance of < 175 m. Similarly, of 13 schizogamics, the migration for 14 species is also limited to < 320 m. Only a capitellid (Capitella capitata), a glycerid (Glycera oxycephala) two ctenodrillid species and a syllid (Eusyllis blomstrandi) migrate over distances ranging from 1,310 to 4,000 m. In preparation of this migration, the epitokous form, for example, the epigamic Raricirrus variabilis grows to a larger size in length (14.6 mm), width (1.3 mm) and in chaetigerous segments (32), in comparison to the atokous form with

Epitoky  163

Table 5.4 Reported depths of collected epitokous species. For reference, see Tables 5.1 and 5.2 Species

Depth (m)

Species

Epigamy

Schizogamy

Nereididae

Syllidae

Depth (m)

Nereis falcaria

6

Grubeosyllis clavata

2

N. fucata

52

Odontosyllis enopla

2

O. phosphorea

3

O. luminosa

6

Typosyllis variegata

10

Capitellidae Capitella capitata

1500

Eunicidae Eunice schemacephala

150

Trypanosyllis coeliaca

43

E. siciliensis

33

Odontosyllis polycera

70

Glyceridae

Syllis gracilis

75

Glycera capitata

17

Haplosyllis spongicola

189

G. americana

37

Exogone naidina

210

G. macrobranchia

55

Syllis ramosa

250

G. oxycephala

2951

Phyllodocidae Phyllodes groenlandica

38

Nereiphylla castanea

45

Paranaitis polynoides

82

Phyllodoce longipes

175

Trypanosyllis zebra

320

Eusyllis blomstrandi

4000

Ctenodrillidae Monticellina heterochaeta

1310

Raricirrus variabilis

4000

5.8 and 0.5 mm with 27 chaetigers (Dean, 1995). In the schizogamics too, the number of swimming segments is increased to 50% of the total number of segments and their chaetiger length is also increased to 1.5 mm length in Odontosyllis polycera (Daly, 1975). Swarming males of O. enopla have less number of segments (~ 85) than females (~ 116) (Fischer and Fischer, 1995). Except for these bit and pieces of information, no other relevant information is yet available on morphological changes in the epitokes. The duration of swarming lasts for 25 minutes in O. luminosa, which undertakes a vertical migration from 6 m depth (Gaston and Hall, 2000). O. enopla climbs a distance of just 2 m to reach the surface waters. Incidentally, the worms seem to swim fast. High speed swimming has been documented in film shots of Platynereis dumerilii (Fischer, 1985) and epitokous Autolytus prolifer (Fischer et al., 1992). The climb involves a concave bent on the dorsal side of the trunk followed by lateral waves passing from posterior to anterior (Fischer and Fischer, 1995). It is difficult to imagine how the little (15 mm length) R. variabilis can sustain

164  Reproduction and Development in Annelida

its climb to cover a vertical distance of 4,000 m. Certainly, these small worms may adopt a different strategy like reduction in buoyancy to climb over 4,000 m distance. Till this day, this area remains a virgin field of research. An attempt has been made to correlate vertical distance climbed by the epitokous worms with their body length and/or number of segments and/ or body weight. Even with a computer search only limited information is available on body length and segment of epitokes (e.g. Glycera dibranchiata: Maryland 7–26 cm, Nova Scotia, 13–36 cm, Wiscasset 18–51 cm [Creaser, 1973], Eunice viridis, 20 cm body length [Caspers, 1984], 80 cm body length, 300–700 parapodial segments in Eunice siciliensis [Hofmann, 1974], 13 mm length, 60 segments in Monticellina heterochaeta [Martin and Gil, 2010], Syllis vittata, 25 mm length [Salazar-Vellajo, 1996]). Almost parallel trends are obtained for the relationship between body size and vertical distance traveled by epigamics and schizogamics (Fig. 5.3). Surprisingly but certainly, the ability

Figure 5.3 Effect of body size on vertical distance climbed by epigamic and schizogamic polychaetes. Data for vertical distribution are taken from Table 5.4. Due to non-availability of information on body length of many of these worms, only approximate trends are shown.

Epitoky  165

to travel vertical distance decreases with increasing body size in both epigamic and schizogamic epitokes. Of course, with more inputs of data, the level may change but not the trends. Among the epigamics, the ability for vertical climb decreases in the following order: Ctenodrillidae < Capitellidae < Phyllodocidae < Glyceridae < Nereididae < Eunicidae. Besides body length, departures from the stream-line body shape by curves, bends, projections and other structures may considerably decelerate the vertical migration, especially in the schizogamics (see Fig. 5.2).

5.5  Swarming Phenomenon In epitokes, the swarming phenomenon is timed by a complex hierarchy of endogenous and environmental factors (e.g. temperature, photoperiod, luminescence). Long day length and elevation in temperature have timed the warmer spring-summer (in south the austral spring-summer) as a favorable season for swarming in temperate and tropical epigamics (Table 5.5). In semelparous epigamics, swarming occurs in the terminal end of their life during the warmer season, during July in the Mediterranean Eunice siciliensis and August in the Indian riverine Dendronereis aestuarina (Fig. 5.4A, B). In the epigamics, seasonal rhythms seem to play a role in timing the swarming phenomenon. In schizogamics also, long daylength and elevated temperature stimulate stolonization and short daylength and low temperature inhibit it (Franke, 1985). Swarming rhythms: On the other hand, swarming in schizogamics is timed by a combination of annual, lunar and diel rhythms. It repeatedly occurs during a specific warmer season May–June in Odontosyllis luminosa. In Typosyllis prolifera, each cycle lasts for 31 days, which includes 17 days of regenerative phase and 14 days of stolonization phase (Franke, 1986a). Hence, the schizogamics can undertake repetitive swarmings. For example, O. phosphorea undertakes as many as nine swarming cycles within a season between June 30th and October 30th (Fig. 5.4C). In it, the timing of swarming is progressively postponed from ~ 30 minutes after sunset in July to ~ 60 minutes after sunset in October (Tsuji and Hill, 1983). Tuned to lunar rhythm, swarming occurs during a specific lunar phase (Table 5.5) and is reported to occur 3–5 days around that specific lunar phase. Amazingly, tuned to lunar cycle, swarming is precisely timed by the diel rhythm. Notably, it occurs during dusk in most species and dawn in T. prolifera. Dusk and dawn are selected to minimize their visibility and predation. Nocturnal high tides (e.g. Syllis amica) and neap tides (O. phosphorea) are also selected to ensure the maximum dispersal.

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Table 5.5 Swarming timed by annual, lunar and diel rhythms in some epitokes Epigamics Nereis fucata

April, Atlantic (Gilpin-Brown, 1959)

N. falcaria

January, austral summer, Australia (Read, 1974)

N. virens

July, Andaman Islands, India (Muruganantham et al., 2015)

Dendronereis aestuarina

August, Southwest Indian river (Jayachandran et al., 2015)

Glycera dibranchiata

July, USA (Creaser, 1973)

Eunice siciliensis

July, Mediterranean (Hofmann, 1974)

E. viridis

October–November, austral spring, Samoan Islands (Caspers, 1984)

Odontosyllis enopla

May, Bermuda (Markert et al., 1961, Fischer and Fischer, 1995) Schizogamics Annual rhythm

O. enopla

May, Bermuda (Markert et al., 1961, Fischer and Fischer, 1995)

O. luminosa

May–June, Caribbean Belize (Gaston and Hall, 2000)

O. phosphorea

June–October, South California (Tsuji and Hill, 1983)

O. polycera

October, austral apring, New Zealand (Daly, 1975)

Syllis amica

Mid-July–mid-August, France (Herpin, 1925)

Typosyllis prolifera

March–October, Adriatic Sea (Franke, 1985) Lunar rhythm

S. amica

A few days after the first quarter phase of the moon at nocturnal high tide

O. enopla

A few days immediately after the full moon

O. phosphorea

First and third quarter phases of the moon during neap tides

O. polycera

Last quarter phase of the moon

E. viridis

Third quarter phase of the moon

O. polycera

Precisely 30-minutes after sunset

O. enopla

Between 15 and 55-minutes after astronomical sunset

O. phosphorea

60-minutes after sunset

O. luminosa

Between 55 and 75-minutes after astronomical sunset

T. prolifera

Between 30-minutes before and 20-minutes after sunrise

Diel rhythm

Besides temperature and photoperiod, some epitokes also engage luminescence to attract the opposite mating partners. Incidence of luminescence has been described in the epigamic Odontosyllis enopla (Fischer and Fischer, 1995) and schizogamics O. hyalina (Lummel, 1932), O. undecimdonta (Inoue et al., 1990), Pionosyllis pulligera (see Franke, 1999) and O. luminosa (Gaston and Hall, 2000). It is a secretory process and

Epitoky  167

Figure 5.4 Swarming season and lunar periodicity in epitokes A. Eunice siciliensis, B. Dendronereis aestuarina and C. Odontosyllis phosphorea. Note a single event in the epigamics but repeated events in the schizogamics (Figure is compiled and redrawn from Hofmann, 1974, Jayachandran et al., 2015, Tsuji and Hill, 1983).

the luminescence slime is found even in one-month old O. phosphorea. Bright flashes of it are also observed in post-swarming O. enopla that has returned to benthic life. Hence, the role played by luminescence in swarming in O. enopla and O. phosphorea is not clear. However, swarming luminesced female O. enopla is reported to attract males, which also luminesce. P. pulligera displays vivid luminescence during swarming. The males are attracted by the luminescence; with their large eyes, they can precisely locate the luminescent female (see Franke, 1999). In O. luminosa, which is distributed throughout the tropical waters of the Western Hemisphere, the glow is bright and visible upto 30–50 m distance. Interestingly, it is the luminescent glow of O. luminosa in the Caribbean that is reported to have provided hope for the nearby land mass to the totally exhausted Christopher Columbus in 1492.

5.6  Pheromones and Spawning In the pelagic realm, swarming leads to high density (e.g. 14,800/m2 Dendronereis aestuarina, Jayachandran et al., 2015) of mating partners and it lasts for 30 minutes in Odontosyllis luminosa (Gaston and Hall, 2000). During this brief pelagic stay, the pairing of mating partners is followed by speciesspecific and the sex-specific nuptial dance and other behavioral signals that eventually lead to the shedding of gametes. A series of publications by Boilly-Marer (1974, 1984), Boilly-Marer and Lassalle (1980), Boilly-Marer and

168  Reproduction and Development in Annelida

Lhomme (1986), Zeeck et al. (1988, 1996, 1998) and Hardege (1999) on the non-epitokous Platynereis dumerilii and Arenicola marina (Hardege et al., 1996) have contributed to our knowledge on pheromones and their role to induce spawning in these polychaetes. Their findings, though not immediately relevant to epitokes, are briefly summarized. Essentially, the following is the sequence of mating and spawning: 1. Male discharges an egg-releasing pheromone, 2. The Cysteine-Glutathione-Disulfide (CGD) pheromone stimulates a female to swim at high velocity in narrow circles surrounded by swarming males (Ram et al., 1999), 3. After an induction period of 30–40 seconds, the female spawns, 4. Subsequently, the male emits a cloud of sperm. P. dumerilii detects a pheromone signals by their modified cirri. The bipolar neurons innervating the chemoreceptors transmit the electrophysiologically measureable excitation along the ventral nerve cord to the brain ganglia. From the coelomic fluid of males, a few low molecular and low polarity fractions have been extracted. These fractions induce the female to spawn. Of > 50 volatile substances, 5 methyl-3-heptanone, 3, 5-octadien2-one and possibly 3-methyl-uric acid and xanthane are important. The heptanone is a Mate Recognition Pheromone (MRP), 3, 5-octadien-2-one is an Egg Releasing Pheromone (ERP) and uric acid acts as Sperm Release Pheromone (SRP) (see Andries, 2001). The actual concentration of the heptanone produced by the worm is 2.5-fold higher the threshold required to induce spawning in female. However, the 5 methyl-3-heptanone successfully induces 20% of females only. To induce 96% females, coelomic fluid has to be added to the heptanone. In Nereis succinea, injection of L-glutamic acid, inosine and quanosine induces egg release alone. Hence, P. dumerilii is reported to use a complex pheromone bouquet to co-ordinate the nuptial dance and gamete release. Spawning act: With mating and spawning limited to a single event in semelparous epigamics, the mature worm begins to twists its body in a peculiar screw-like manner, breaks off from the infertile anterior body part and releases the gametes through ruptures of the body wall (e.g. Eunice siciliensis, Hofmann, 1974). In schizogamics, once a pair is formed, the female begins to quiver rapidly with periodical spinning circles and releases a cloud of gametes (e.g. O. luminosa, Gaston and Hall, 2000) and coelomic fluid (see Andries, 2001). These gametes stimulate the encircling males to milt. Sometimes, several males may encircle a single female (polyandry, cf polygyny in the hybrid male of Autolytus spp inseminating eggs of many females, Schiedges, 1979). The sequence of events from formation of heteronereis/stolon to spawning is briefly summarized in Fig. 5.5. In polychaetes, especially the epitokous syllids, extirpation of the proventriculus induces stolonization in non-reproductive worms (Franke,

Epitoky  169

Figure 5.5 Sequence inductions that lead to the formation of heteronereis/stolon until gamete shedding in epitokes. MRP = Mate-Recognition Pheromone, ERP = Egg-Release Pheromone, CGD = Cysteine-Glutathione Disulfide, SRP = Sperm-Release Pheromone.

1980). Amputation of the prostomium causes rapid disintegration of growing oocytes followed by spermiogenesis (Kahmann and Franke, 1984). These contributions are elaborated in Chapter 7.

6 Sex Determination

Introduction In annelids, sexuality includes gonochorism, parthenogenesis and hermaphroditism, which comprises of simultaneous, protandric, protogynic and serial hermaphrodites. More than 76% of annelids are gonochores. Hermaphroditic oligochaetes (18.7% of annelids), hirudineans (4.0%) and polychaetes (0.6%) make up the remaining 24% (see p 64). In gonochoric polychaetes, either male or female heterogametic chromosomal mechanisms of sex determination may be in operation. Sex ratio represents the cumulative end product of sex determination and sex differentiation processes. Table 2.2 shows that only in a few polychaetes, the expected ratio of 0.5 ♀ : 0.5 ♂ is reported. In many others, the ratio is, however, significantly deviated. These deviations indicate (i) amenability of sex chromosomes to overriding autosomal genes and alteration of the sex differentiation process exactly toward the opposite sex, and (ii) environment-dependent polygenic autosomal sex determination (Premoli et al., 1996). However, it must be indicated that our knowledge on sex determination in annelids is elementary, despite the presence of many freshwater oligochaetes with a short life span (e.g. naidids). With the presence of about 190 clonal species, the scope for their restoration to sexuality even after 1,000–3,000 rounds of clonal cycles may make the subject a fascinating one.

6.1  Karyotype and Heterogamety Spermatogonia (male) and regenerating tail (female) of polychaetes have been used as tissues for karyotyping. In Neanthes japonica, the chromosome size ranges from 2.3 to 5.8 µm and most of them are metacentric (Sato and Ikeda, 1992). The number of 2n chromosome ranges from 12 in Pristina aequiseta to 54 in Aporrectodea rosea (Table 6.1). In Serpula vermicularis, the reported

Sex Determination  171

Table 6.1 Chromosome number in annelids Species

Number

Ditrupa arietina Ficopomatus enigmaticus Filograna implexa Hydroides elegans H. norvegica Placostegus tridentatus Pomatoceros triqueter Serpula vermicularis Spirorbis spirorbis S. corallinae S. tridentatus Janua pagenstecheri Circeis spirillum Dinophilus gyrociliatus Capitella capitata Neanthes japonica Hediste atoka H. diadroma H. japonica Nereis diversicolor N. acuminata Polydora curiosa

20 26 20–44 26 22 20 24–26 14–28 20 20 20 20 20 XX-XO ZW-ZZ 28 XX-XY 28 XX-XY 28 XX-XY 28 XX-XY 28 18, 22, 24, 28 XX-XY

Reference

Polychaetes see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) see Kupriyanova et al. (2001) Prevedelli and Vandini (1999) Petraitis (1985a, b) Sato and Ikeda (1992) Tosuji et al. (2004) Tosuji et al. (2004) Tosuji et al. (2004) Christensen (1980) Pesch and Pesch (1980) Korablev et al. (1999)

Aquatic oligochaetes Aeolosoma hemprichi Branchiodrilus sowerbyi Dero indica Frederiova bubosa Henlea ventriclosus Limnodrilus claparedianus L. hoffmeisteri L. undekemianus Pristina aequiseta Tubifex blanchardi T. costatus T. tubifex Lumbricillus lineatus

56 24 36 32 34 25 24 25 12 50 25 50, 100, 150 26, 39, 52, 65

Naidu (2005) Naidu (2005) Naidu (2005) Naidu (2005) Naidu (2005) Christensen (1980) Naidu (2005) Christensen (1980) Naidu (2005) Marotta et al. (2014) Christensen (1980) Marotta et al. (2014) Coates (1995)

Earthworms Allolopophora caliginosa A. chlorotica A. iterica A. nocturna A. terrestris

36 32 32 36 36

Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Table 6.1 contd. ...

172  Reproduction and Development in Annelida

...Table 6.1 contd. Species

Number

Reference

Earthworms Aporrectodea trapezoides A. rosea Bimastos eiseni B. tenuis Eisenia foetida E. rosea f.typica E. rosea f.mut E. venata Eiseniella tetraedra Dendrobaena mammalis D. rubida D. subrucunda Dendrodrilus rubidus D. octaedra Octolasion tytaeum O. cyaneum O. lacteum Lumbricus castaneus L. festiwas L. friendi L. rubellus L. terrestris

36, 54 54, 90, 108 32 48 22 54 33 36 72 34 68 68 34, 68, 102 108, 124 38, 54, 72 190 38 36 36 36 36 36

Jaenike and Selander (1979) Jaenike and Selander (1979) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Jaenike and Selander (1979) Jaenike and Selander (1979) Jaenike and Selander (1979) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952) Muldal (1952)

Leeches Glossiphonia complanata Glossiphonia complanata concolor Glossiphonia heteroclita Glossiphonia heteroclita papillosa Theromyzon tessulatum Branchellion torpedinis Hemiclepsis marginata Pontobdella muricata Dina lineata Erpobdella octoculata Erpobdella testacea Nephelopsis obscura Trocheta subviridis Trocheta bykowskii Haemopis sanguisuga Hirudo nipponia Placobdella papillifera Theromyzon rude Hirudo medicinalis H. vertana H. orientalis

28 28 16 16 16 12 32 20 18 16 16, 22 22 22 22 26 16 24 14 28 26 24

Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) see Davies and Singhal (1987) Yang et al. (1997) Davies and Singhal (1987) Davies and Singhal (1987) Utevsky et al. (2009) Utevsky et al. (2009) Utevsky et al. (2009)

Sex Determination  173

numbers are 14 and 28 for specimens collected from the Mediterranean and Norwegian fjord, respectively. Similarly, they are 20 for Filograna implexa collected from the UK waters but 44 for that from Norway. It is not clear whether the Norwegians specimens are tetraploids. The 2n number of Nereis acuminata is 18, 22, 24 or 28 (Pesch and Pesch, 1980); it is also not clear whether the one with 28 chromosomes is a triploid. Karyotyping has revealed the presence of male (XX-XY) heterogametic chromosomes in Neanthes japonica (Sato and Ikeda, 1992), Hediste atoka, H. diadroma and H. japonica (Tosuji et al., 2004). All these nereidids have 14 pairs of chromosomes including an unusually large sized Y chromosome. The spionid Polydora curiosa is also male heterogametic (Korablev et al., 1999), in which the Y chromosome is also larger than the X chromosome (Jablonka and Lamb, 1990). Possibly, the Y chromosome of polychaetes accumulates multiple transposable elements. Experimental crosses have also revealed the presence of male heterogamety in Dinophilus gyrociliatus (XX-XO, Prevedelli and Vandini, 1999) and female heterogamety in Capitella capitata (ZW-ZZ, Petraitis, 1985a, b).

6.2  Gametic Compatibility In gonochores, reproduction is characterized by homospecific compatibility between egg and sperm and heterospecific incompatibility between them. Nevertheless, there are exceptions for each of heterospecific compatibility and homospecific incompatibility in annelids. Successful hybridization between the syllid Autolytus prolifer and A. brachycephalus as well as survival and reproduction of their hybrids have been described earlier (p 162). Another example for successful hybridization is between the nereidids Hediste diadroma and H. japonica. These two epitokes are sympatric only in the Omuta-gawa River, Japan. Their gametes are released without involving pairing between male and female partners and the nuptial dance. Their gametes are reciprocally compatible and hybrids between them are viable until 23rd day after fertilization. The differences in the developmental stages between these two nereids and their hybrids are listed in Table 6.2. The hybrid larvae are intermediate phenotypes but with a greater maternal influence in characteristics like the relative length of chaetae and lecithotrophic larval duration. Neither the pre-mating gametic incompatibility nor hybrid viability appears to ensure reproductive isolation between these nereids (Tosuji and Sato, 2006). However, two points may have to be noted: 1. The hybrids between Autolytus spp sexually mature and successfully generate stolons. But the hybrids between Hediste spp are not reported to sexually mature and

174  Reproduction and Development in Annelida

Table 6.2 Time course of important developmental stages in Hediste diadroma and H. japonica and their hybrid offspring (condensed and compiled from Tosuji and Sato, 2006) Development Stage

H. diadroma

H. japonica

H. diadroma egg × H. japonica sperm

H. japonica egg × H. diadroma sperm

Formation of I polar body II Polar body formation

100 minutes

90 minutes

120 minutes

120 minutes

130 minutes

115 minutes

180 minutes

180 minutes

Trochophore hatching

38 hours

44 hours

38 hours

44 hours

Metatrochophore

72 hours

68 hours

72 hours

72 hours

Notochaeta 4th chaetiger

16–30 days

7–8 days

12–23 days

9–10 days

5th chaetiger

> 30 days

10–20 days

~ 31 days

~ 30 days

reproduce, albeit both the species have the same number of 2n chromosomes (28), which morphologically look alike (Tosuji et al., 2004). 2. Fertilization success (FS) is ~ 95% in Hediste spp, indicating the near 100% compatibility between their respective eggs and sperm. Notably, the heterospecific compatibility between H. diadroma egg and H. japonica sperm, as indicated by FS, is reduced to ~ 80%, in comparison to that (~ 85%) between H. japonica egg and H. diadroma sperm. This minor level of heterospecific incompatibility can reach a very high level between two morphotypes within a species and surprisingly within populations of a single species. In Spirobranchus polycerus, two morphotypes are recognized. Of them, the seven-(opercular) horned morph is hermaphrodite and two-horned morph gonochore. Gametic compatibility and hence FS for the homomorphotypes is ~ 98%. Marsden (1992) has attempted to fertilize eggs of the gonochoric morph with sperms from hermaphroditic morph. None of the eggs have cleaved, a marker of egg-sperm compatibility. Hence, the gametic incompatibility between these two morphs within the species S. polycerus has been confirmed. Another serpulid Galeolaria caespitosa is widely distributed around Australia from the eastern coast at Sydney to the southern coast of Adelaide, covering a coastal distance of 2,200 km. In it, gametic compatibility and cross-fertilization occur among populations across temperate Australia. However, fertilization assays have revealed the pre-mating asymmetrical differences between very distantly located populations with near complete incompatibility between eggs from Sydney with sperm from Adelaide. However, the reverse cross is reasonably compatible (Styan et al., 2008). The low level of gametic incompatibility observed between Hediste spp and high level of incompatibility reported between the two morphotypes within S. polycerus and that reported for the distantly located populations within G. caespitosa can be explained from our knowledge on echinoderms.

Sex Determination  175

In them, the presence of bindin in the sperm acrosome serve as an adhesive responsible for sperm attachment to the vitelline layer of homospecific eggs. Hence, bindin plays an important role in determining whether the gametes of two species are compatible and fertilize each other. However, the role of bindin is not a simple lock- and key-mechanism, in which molecular change(s) in sperm need not necessarily result in analogous change(s) in the egg also and vice versa. As a consequence, heterospecific gametic incompatibility may be complete or incomplete. Gametic compatibility is defined as the ratio of mean percentage of eggs fertilized in homospecific crosses at a sperm concentration required to fertilize ~ 90% of eggs. In such species, analogous molecular change(s) have either occurred or not occurred in eggs and sperms, the ratio of reciprocal heterospecific compatibility is 1.00 (see Pandian, 2018). However, the ratio begins to decrease with slight changes in sperm of H. japonica but not in the egg of H. diadroma. The ratio is also significantly decreased to almost zero with major changes in eggs of G. caespitosa from Sydney but not in its sperm from Adelaide. Off from these, allopatric isolation may lead to post-mating isolation due to karyotypic changes. Low dispersal and sexual selection are characteristics of coastal polychaetes like the monogamous N. acuminata with male parental care. From a karyotypic study, Weinberg et al. (1990) have reported that the N. acuminata populations from the North Atlantic have karyotypes with 11 pairs of small acrocentric chromosomes (2n = 22), while their Pacific counterparts have nine pairs of large metacentric or submetacentric 2n = 18 chromosomes. The Atlantic and Pacific North American populations have remained allopatric for a long time. They are now almost different species and post-mating reproductive isolation has evolved as an incidental byproduct of allopatric divergence between populations.

6.3  Sex Ratio and Variations Male heterogamety has been recognized in the gonochoric Dinophilus gyrociliatus. This has been described earlier (p 90). However, a couple of sentences are required for the sake of continuity. In D. gyrociliatus, a single ovary simultaneously produces many small (XO) male eggs and one larger (XX) female egg. This unique mechanism allows the mother to overcome the chromosomal mechanism of sex determination by selective fertilization of larger eggs by X bearing sperms and small egg by sperm without sex chromosome. A number of environmental factors alter the ratio between small and larger eggs. Optimal temperature 28ºC and salinity of 30‰ maintain the ratio at 1 ♀ : 3 ♂ by producing 1 large and 3 small eggs (Akesson and Costlow, 1991). Similarly, the optimal diet tetramin increases fecundity

176  Reproduction and Development in Annelida

and thereby alters sex ratio (Prevedelli and Simonini, 2000). The presence of multiple females can also significantly skew sex ratio toward males (Minetti et al., 2013). Through experimental crosses, heterogamety is identified in Capitella capitata. The 27 crosses between females and males of C. capitata have produced 2,110 offspring with sex ratio of 0.47 ♀ : 0.53 ♂. But 31 crosses between hermaphrodites and males have yielded 3,264 offspring; of them 96.7% are males. Apparently, females, males and hermaphrodites are characterized by ZW, ZZ and ZZ, respectively (Petraitis, 1985a, b). However, two subsequent series of experiments made the picture a little more complicated. In the first series, the effects of density have been tested on the expression of gender and hermaphroditism. The second one has tested the maternal effect on the incidence of hermaphroditism. From these experiments, Petraitis (1991) has drawn the following inferences: 1. Families determine the age of female maturity. The overall age of maturity of females is 39 days. 2. The first transition from male to hermaphroditism occurs at the age of 76 days. 3. Males are more prone to become hermaphrodites, especially in isolation or in the presence of excess number of males. 4. ZW mothers produce ZW females but hermaphroditic mothers (ZZ) only ZZ offsprings (Fig. 6.1). 5. ZZ females are more likely to become hermaphrodites. 6. Higher density induces sex transition but isolated ones switches to hermaphroditism. Briefly, isolated males and females are prone to become hermaphrodites. Clearly, ZW and ZZ genes are harbored on respective sex chromosomes. The environmentdependent differential expression of ZZ/ZW gene(s) stimulate simultaneous function of male and female and thereby hermaphroditism. Or the expression of Z gene(s) is inhibited to produce females. Incidentally, Akesson (1982) has

Figure 6.1 Types of sex change in Capitella capitata (modified and redrawn from Petraitis, 1991).

Sex Determination  177

described the dominant yellow colored YY eggs and recessive (yy) eggs in Ophryotrocha diadema and effect of these genes on survivorship and spawning interval. This publication may not be of immediate relevance. Unlike the widely distributed G. caespitosa, populations of Ophryotrocha labronica are found around the Italian coast. With differences in the biokinetic range for salinity tolerance, the Naple (Na) and Venice (Ve) populations are further divided into Na II, III and IV as well as Ve II and Ve IV. With gametic compatibility and successful hybridization between the representatives of these populations, male ratio among F1 progenies ranges from 0.35 in a cross between ♂ Ve II and ♀ Ve II to 0.84 in another cross between ♂ Na III and ♀ Ve II (Table 6.3). When Na IV ♂ is crossed with Na IV, Na III and Ve II females, the male ratio is increased from 0.56 to 0.64 and 0.82. Hence, the female partner is responsible for the increased male ratio. It is also responsible for the increased ratio from 0.36 to 0.47 and 0.84 in combinations of ♂ Na III × ♀ Na III, ♂ Na III × ♀ Na IV and ♂ Na III × ♂ Ve II. Clearly, Ve II alters the ratio in favor of males. However, a cross between ♂ Ve II × ♂ Ve II has yielded the lowest male ratio, indicating the overriding impact of Ve II male on the increase of female ratio (Lanfranco and Rolando, 1981). From their studies on heritable variation in sex ratio of O. labronica, Premoli et al. (1996) have proposed an updated version of an old hypothesis by Bull et al. (1982). Accordingly, sex is determined by a multilocus genetic system, facilitating the combined effects of a female major sex gene (as it is proposed for Capitella capitata) and masculanizing modifiers. Polygenic sex determination is a quantitative trait. It is rare in gonochoric animal species and evolutionary unstable (see Pandian, 2011). From their experiments, Premoli et al. (1996) have shown that O. labronica has a major sex determining gene with two alleles. Meiotic segregation imposes a sex ratio of 0.5 to the Table 6.3 Effects of crossing Naple and Venice populations of Ophryotrocha labronica on male ratio (condensed and compiled from Lanfranco and Rolando, 1981) Crosses Between Populations

Male Ratio

♂ Na IV × ♀ Na IV

0.56

♂ Na IV × ♀ Na III

0.64

♂ Na IV × ♀ Ve II

0.82

♂ Na III × ♀ Na III

0.36

♂ Na III × ♀ Na IV

0.47

♂ Na III × ♀ Ve II

0.84

♂ Ve II × ♀ Ve II

0.35

♂ Ve II × ♀ Na IV

0.51

178  Reproduction and Development in Annelida

oocytes of female. Variations from 0.2 to 0.8 in sex ratio are explained by the presence of polygenic system in the male parent, which may override the sex imposed by progamic maternal mechanism. Some evidences are brought for the heritable variations in sex ratio of O. labronica. Though attractive, the hypotheses of Premoli et al. (1996) remains to be tested in other annelid species, in which the factors responsible for variation in sex ratio are also inheritable.

7 Sex Differentiation

Introduction In animals, sex determination and differentiation are successive but diverse processes that have evolved independently a number of times (Hodgkin, 1990). Hormones and neuroendocrines act as chemical messengers of genetic cascade that realizes sexualization and maintenance of sex as well as regulation of reproductive cycles in animals. In annelids, researches on these aspects have progressed from histological to surgical study to know the source and target of a specific endocrine as well as from injection (of a hormone) to in vitro and in vivo incubation studies to understand the action and biosynthesis of a hormone, respectively. Despite having a well-developed circulatory system, annelids do not possess a developed glandular endocrine system. No information is yet available on endocrines of hermaphroditic oligochaetes and hirudineans. Although most hirudineans are protandrics, they are not amenable to surgical study, as they do not have the ability to regenerate any tissue; most aquatic oligochaetes are too small (e.g. aeolosomatids measuring < 0.5 mm) for any surgical study. But the hermaphroditic earthworms are large in size (e.g. Australian giant earthworm Megascolides australis measuring 3 m) and have the ability to regenerate the ‘head’ including the prostomium harboring the ‘brain’ (see p 96). Our understanding of endocrine sexualization and regulation of reproductive cycles is based on findings from temperate polychaetes only; for their tropical counterparts, who have greater scope for aquaculture, no information is yet available. Using histology including immunohistochemistry (e.g. Weidhase et al., 2016), surgical (e.g. Pfannenstiel, 1978b) techniques, injection (e.g. Durchon, 1952) and incubation (e.g. Lawrence and Soame, 2009), subesophageal ganglia, prostomium and proventriculus have been identified as a source of endocrines in polychaetes. Expectedly, a fairly large number of publications is available; they have been reviewed from time to time; however, some of these reviews are restricted to nereidids alone (e.g. Andries, 2001). Unlike in crustaceans (Pandian, 2016) and echinoderms (Pandian, 2018), sex differentiation in polychaetes is a labile and protracted process.

180  Reproduction and Development in Annelida

7.1  Endocrine Regulation Seasonal changes in temperature and photoperiod influence biological cycles like phytoplankton production, which, in turn, regulate reproductive cycle of temperate polychaetes (e.g. Harmothoe imbricata, Garwood and Olive, 1982). Conversely, the breeding season in cosmopolitan species like Cirratulus cirratus is not restricted; C. cirratus females spawn mostly two–three times/y and sometimes four–five times/y (Olive, 1970). This may also hold true for tropical polychaetes. Within temperate polychaetes, the breeding cycle is characterized by (i) semelparity, as in some errant nereidids, (ii) iteroparity (a) but with extended period of vitellogenesis, as in errant nephtyids, (b) with a short period of vitellogenesis, as in errant phyllodocids and (iii) sedentary polychaetes like Arenicola marina. Notably, tubiculous polychaetes have not yet received attention by endocrinologists. Table 7.1 summarizes available information on known endocrines and their action in polychaetes (see also Porchet et al., 1989).

7.1.1 Nereidids Experimental grafting of the suboesophageal ganglion of immature Nereis diversicolor has led Golding (1967) to conclude that a single brain hormone is responsible for the promotion of somatic growth and inhibition of sexual maturation. Confirming his conclusion, Dhainaut (1970) has reported the accelerated growth of oocytes as small as 50 µm in Nereis pelagica. In Perinereis dumerilii, removal of the brain results in precocious sexual maturation but abnormality and degeneration of oocytes. However, these impairments are restored by replacing the brain (Hauenschild, 1966). Hence, the brain hormone also plays a gonadotrophic role by promoting progressive sexual development with gradual decrease in its titer. Briefly, the single brain hormone namely Juvenile Hormone (JS), the nereidin in nereidids performs a range of functions such as somatic growth and regeneration (see p 100), gonadotrophic support for oocyte growth and regulation of eleocyte activity as well as transition to sexual maturity and epitoky (see Andries, 2001, Lawrence and Soame, 2009). Incidentally, considerable asynchrony among smaller oocytes of neriedids is reported but the oocyte growth becomes synchronized with declining titer of JH and as the worm approaches epitoky (Fischer, 1974). Through a series of experiments, in which oocytes of different sizes (47 or 50 µm) have been incubated with no ganglia or ganglia from juvenile or mature P. dumerilii, the following have been inferred: (i) growth is significantly faster in oocytes of 47 and 50 µm with no ganglia than that with ganglia from juvenile, (ii) it is significantly faster on incubation with fresh or boiled ganglia from mature P. dumerilii, (iii) it is also faster in

Arenicola marina

Syllidae, Franke (1976, 1977)

Arenicolidae, Howie (1963)

Stolonization inhibition hormone

Ootrophic hormone

Inhibits stolonization

Oogonial sex determination

Vitellogenesis promotion

Spermatozoa are released from sperm cluster

Ripening and release of gametes

Proventriculus

Syllis prolifera

Dorvilleidae, Pfannenstiel (1978b)

Vitellogenesis promotion hormone

Promotion of oogenesis Promotion of spawning

Switches off action of JH

8, 11, 14-eicosatrienoic acid

Prostomium

Ophryotrocha spp

Phyllodocidae, Olive (1975)

Gonadotrophic hormone, Spawning hormone

+ ve effect on oocyte growth

Coelom

Prostomium

Eulalia viridis

Nephtyidae, Olive (1970)

Dopamine

+ ve effect on oocyte growth

Bentley et al. (1990)

Cerebral ganglia

Nephtys hombergii

Lawrence and Soame (2009)

Oxytocin

Vitellogenin synthesis

Oocyte development

–

N. succinea

Lawrence and Soame (2009)

Serotonin

17-β Estradiol

Coelomic maturation factor

–

P. dumerilii

Lawrence and Soame (2009)

Inhibition of JH

Gametogenesis inhibition

Prostomial maturation factor

–

N. diversicolor

Garcia-Alonso et al. (2006)

Feedback substance

Effect

Coelom

Eleocytes

N. virens

Durchon (1952)

Hormone Juvenile hormone (JH)

Prostomium

Coelomocytes, oocytes

Perinereis cultrifera

Nereidae, Golding (1967)

Watson and Bentley (1997)

Source Cerebral ganglia

Species Nereis, Platynereis

Family/Reference

Summary of known hormones, their source and action in some polychaetes

Table 7.1

Sex Differentiation  181

182  Reproduction and Development in Annelida

P. dumerilii oocytes, when incubated with fresh ganglia from mature N. virens as well as (iv) oocytes (70 µm) of N. succinea with the ganglia from mature P. dumerilii female (at low concentration) (Olive and Lawrence, 1990, Lawrence and Olive, 1995). Arguably, these results have brought evidences that the JH is not species-specific within neriedids. The thermostable structure and action of JH are highly conserved among the neriedids (Lawrence and Soame, 2009). Although the brain hormone from juvenile neriedids is reported to perform multiple functions, the incidence of a series of other hormones has also been reported. Estradiol-17β has been isolated from the coelomic fluid of N. virens, in which it promotes the secretion of vitellogenin by the eleocytes in mature females (Garcia-Alonso et al., 2006). The presence of an array of hormones has been reported such as serotonin (from immuno-positive staining) in N. diversicolor (Heuer and Loesel, 2007), oxytocin-like hormones in Perinereis vancaurica ganglia (Fewou and Dhainaut-Cortois, 1995, Matsushima et al., 2002) and vasopressin from P. dumerilii (expressed in developing forebrain, Tessmar-Raible et al., 2007). Melatonin is a key factor in the photoperiodic control of reproduction in vertebrates (e.g. fishes, see Pandian, 2013). As gonad maturation is controlled by photoperiod in many polychaetes, the possibility does exist for its presence in them also. Examining the effects of some of them, Lawrence and Soame (2009) have reported that dopamine and melatonin switch off the action of JH in P. dumerilii and N. succinea but serotonin and oxytocin have a positive effect on oocyte growth. Feedback substance: In Perinereis cultrifera, the pioneering research by Durchon (1952) has initiated an idea of a feedback factor. Injection of oocytes of 90–100 µm from a submature worm into the coelomic cavity of juvenile eliminates the inhibition of the brain hormone. Increasing size of the injected oocytes accelerates sexual maturity (see also Pfannenstiel, 1978a). Confirming Durchon’s findings, Porchet (1967) reported the down-regulation of juvenile hormone differs according to sex of the recipient. It results in precocious maturity and epitoky in males but abortive oocyte growth in female. Apparently, the feedback factor is sex specific. Hofmann (1974) suggested that the feedback substance emanates from coelomocytes and/or oocytes. However, its chemical identity is not yet known, though suspected to be a glycoprotein (Porchet and Cardon, 1976).

7.1.2 Nephtyids Reproduction in the nephtyids are characterized by iteroparity, intra-ovarian pattern and discrete spawning. Many temperate nephtyids live up to 6–9 years and display an annual reproductive cycle (Olive and Bentley, 1980). In them, reproduction often fails due to (i) inability to initiate gametogenesis (e.g. Nephtys caeca, N. cirrosa), (ii) premature gametogenic degeneration and oosorption and (iii) non-release of viable gametes by gravid worm (= spawning failure) (e.g.

Sex Differentiation  183

N. hombergii and N. cirrosa, Olive et al., 1985). The nephtyids sexually mature mostly at the age of 1+ year or latest by 2+ years. Unlike the nereidids, gametogenesis is well synchronized in nephtyids at all stages from meiotic prophase onward. Only fully grown oocytes are ovulated from the discrete ovary into the coelom. A long-term survey between 1978–1979 and 1983–1984 in the Tyne estuary, England has indicated successful spawning only during 1978–1979 and 1983–1984 (Olive et al., 1985). Our understanding of endocrine regulation of reproduction in nephtyids is based on the following findings: 1. Reproductive cycle is regulated by cyclic production of Gonadotrophic Hormone (GH) secreted and released from the subesophageal ganglia located in the prostomium and it supports the gametogenic development from October to March in N. hombergii. 2. Spawning Hormone (SH) is also released from the subesophageal ganglia during May. Injection of 10 µl of brain homogenate prepared from 40 ganglia/100 µl into gravid N. hombergii female induces spawning more successfully during May than April (Olive, 1970). Decerebration in the early gametogenic phase results in total failure of gametogenesis in N. hombergii. But the catastrophic effects of decerebration can be restored by implantation of whole alive subesophageal ganglia. This observation confirms that the cerebral subesophageal ganglia are the source of an Ootrophic Hormone (OH) (Olive and Bentley, 1980).

7.1.3 Phyllodocids The non-epitokous phyllodocids like Eulalia viridis are characterized by release of oocytes from the dispersed ovary into the coelom (extra-ovarian pattern), where they accumulate from the previous autumn. Unlike the nereidids and nephytids, which require a prolonged duration for oocyte development, vitellogenesis is rapidly completed during April–May in E. viridis (Olive, 1975), when Vitellogenesis-Promoting-Hormone (VPH) is released from the prostomium. The response of an oocyte to VPH is dependent on its size. Smaller oocytes (> 40 µm) are refractory and do not commence vitellogenesis in a decerebrated worm. However, vitellogenesis is restored with an implantation of the ganglia into the coelom. The medium sized oocytes (> 40 but < 100 µm) degenerate, when VPH is deprived by decerebration. But the large oocytes (> 100 µm) survive in the absence of VPH, suggesting that a normal feature of oocyte maturation is by the withdrawal of VPH during the final stages of oocyte maturation (Oliver, 1976). The role of VPH has been elucidated by Lawrence and Olive (1995) using bioassays and incorporation of 3H-leucine. Unlike in the nephytids, VPH promotes the uptake of external protein into the developing oocytes.

7.1.4 Arenicolids Oocyte development in Arenicola marina requires not only Prostominal Maturation Hormone (PMH) but also coelomic fluid (Watson and Bentley,

184  Reproduction and Development in Annelida

1997). Hence, the oocyte development is controlled by a two-steps process namely PMH and Coelomic Maturation Factor (CMF). The CMF is thermolabile, trypsin-sensitive, has a molecular mass of > 30 kDa and suggests a proteinaceous nature. Clusters of spermatogonia are released from the testis into the coelom. In the presence of Sperm Maturation Factor (SMF) 8, 11, 14-eicosatrienoic acid, the breakdown of sperm clusters occurs (Bentley et al., 1990). Ripening of gametes and their release are inhibited by decerebration and are restored by administration of prostominal homogenate, albeit only during the breeding season (Howie, 1963).

7.2  Differentiation and Lability Our understanding of endocrine sex differentiation is based on the findings from the dorvilleid protandric Ophryotrocha spp and gonochoric syllids.

7.2.1 Dorvilleids Pfannenstiel (1971, 1973, 1974, 1975, 1976, 1977a, b, c, 1978a, b) made a series of homospecific and heterospecific prostominal transplantations. In 1980s, when Pfannenstiel made excellent publications (mostly in German language), when our understanding of Primordial Germ Cells (PGCs) in invertebrates has been at its nascent stage. Hence, his findings are briefly summarized but with insertions (in italics) of observations in the present day context. 1. At 6–8 and 16-segment stages, O. puerilis puerilis matures as male and switches to female phase, respectively. 2. (a) Decerebrated juveniles attain male phase but not female phase, even after passing the critical 16-segment stage. (b) On transplantation of prostominal graft from a female donor, the decerebrated recipient differentiates into a female but that from a male donor induces no change. Clearly, a factor originating from the prostomium, i.e. Ootrophic Hormone (OH) (cf Olive and Bentley, 1980) is obligately required for differentiation into a female. In males, factors like dopamine and/or melatonin may switch off the action of juvenile hormone (JH/OH). (c) In females, a portion of the germ cells, i.e. PGCs retain their bisexual potency (it also occurs in fishes, see Pandian, 2013), while the other portion of PGCs is differentiated into oogonia under the influence of OH. 3. Due to Paarkultureffekt (pair rearing effect), one individual in a pair becomes a female within 3–5 days under the influence of OH but the other is pheromonally (?) induced to differentiate into a male around the 8th day. Clearly, the sex differentiation process in the protandric O. puerilis puerilis is labile with retention of a portion of PGCs with bisexual potency. 4. In O. labronica, the process of sex differentiation is more labile than in O. puerilis puerilis. In this protandric, juveniles differentiate into 73% of (phenotypic) primary males and 27% of (genetic)

Sex Differentiation  185

stable females (Fig. 7.1A). Following decerebration, 48% (i.e. 67% of the total) of the phenotypic males remain as (genetic) stable males but the remaining 25% phenotypic males differentiate into secondary (phenotypic) females (Fig. 7.1B). Clearly, a half (24%) of primary males (not amenable to OH) are genetic males and the other half (24%) are amenable to OH and differentiate into phenotypic females (Fig. 7.1C). 5. Even with amputation of prostomium, the stable genetic females and genetic males differentiate into females and males, respectively (Fig. 7.1D). Therefore, other factors like serotonin and dopamine

Figure 7.1 Decerebration and resulting sex ratio in A. juvenile, B. primary male and C. secondary female as well as D. stable female and male on sex ratio of Ophryotrocha labronica (modified freehand drawing from Pfannenstiel, 1978a).

186  Reproduction and Development in Annelida

may also be involved in sex differentiation. 6. All heterospecific transplantations involving the prostominal grafts from females or males of O. labronica into the decerebrated recipient O. puerilis puerilis females have induced differentiation females. Hence, the OH produced by the prostominal graft of O. labronica is compatible and completely compensates the OH required by the decerebrated females and males of O. puerilis puerilis. Like JH within nereidids, OH is also not a species specific hormone within these dorvilleids.

7.2.2 Syllids Stolonization and stolon morphology: In syllids, stolonization is the common mode of reproduction. The proventriculus (PV) produces a hormone that at high titer attenuates stolonization but promotes regeneration of the posterior end, from which a stolon is detached. Weidhase et al. (2016) have described the stolonization cycle and stolon morphology in a representative syllid Typosyllis antoni. Accordingly, female stolons are full of oocytes and males harbor two packages of sperms per segment. The syllid undergoes many successive stolon cycles. The stolons are not generated by segment addition but arise from a transformation of posterior segments. Following detachment of a stolon, the posterior end is regenerated prior to development of a new one. Hence, the syllids have retained the regenerative potency of the posterior even after the commencement of reproduction. Briefly, regeneration is alternated with stolonization. The number of transformed segments in stolons varies between individuals and may also vary between successive stolonization events in an individual from 5 to 18 segments in T. antoni. The anterior most stolon segment bearing eyes and antennae undergoes extensive morphological changes to become the stolon’s ‘head’. Notably, having originated from the posterior tip of the ventral nerve cord, the new structure represents the stolon’s ‘brain’ (Weidhase et al., 2016). Our understanding on sexualization and sex reversal in stolons is based on the researches of Durchon (1975), Hauenschild (1975) and Franke (1976, 1977). The PV is a muscular structure with radially arranged striated muscle cells surrounding the gut (see Weidhase et al., 2016). These cells consist of usually only one or two sacromeres of up to 100 µm length, being the longest known sacromeres within Metazoa (Smith et al., 1973). A histological study of the PV has revealed no sign of glandular or secretory structure. The ventrical and caeca are composed of glandular tissues but they are not involved in the reproductive and regenerative processes (Weidhase et al., 2016). The PV is shown to release Stolonization Inhibition Hormone (SIH), which inhibits stolonization during the breeding season. 1. Sensing the absence of incoming food during starvation, the PV inhibits stolonization in Syllis prolifera, which has just released a stolon. 2. Implantation of PV from a juvenile donor into a recipient without its own PV also inhibits stolonization. 3. However, that from a sexualized donor does not inhibit stolonization in the recipient.

Sex Differentiation  187

Apparently, SIH promotes somatic growth but with its declining titer, sexualization of the stolon is commenced. Hence, SIH of the syllids functions like JH of nereidids. Sex reversal: In syllids, male differentiation is a stable process. Natural sex change from male to female is not yet reported. In fact, spontaneous sex change is rare in syllids. However, extirpation of PV or removal of SIH not only induces precocious stolonization but also leads to sex reversal (Fig. 7.2A). To test sex differentiation in stolons, homosexual transplantation of PV from an undifferentiated male stock of S. prolifera donor was grafted into a recipient male stock. More than 70% of juvenile recipient stocks became sterile and the remaining 30% of them produced male stolons only. However, receiving PV graft from a sexualized donor, the recipient continued to release male stolons only. Heterosexual implantation of PV from juvenile female stock was grafted into male stock that has already released male stolon. Of 27 implantations, 52% of the recipient stocks became sterile. Obviously, heterosexual grafts are incompatible at least in half of the stocks. The remaining stocks produced both female and male stolons at the ratio of 0.37 ♀ : 0.11 ♂ stolons. Hence, sex ratio of the stock was noted as 0.52 sterile: 0.48 fertile (Fig. 7.2B). On transplantation from sexualized donors, the ratio is 0.21 sterile stocks: 0.79 ♂ stolons. It seems that the stocks of S. prolifera retain PGCs with bisexual potency and display a tendency toward the production of male stolon.

Figure 7.2 Stolonization and sexualization in Syllis prolifera. Effect of transplantations of proventriculus A. from juvenile and sexualized donor males on recipient’s sex ratio and B. from juvenile and sexualized female donors to male recipient’s sex ratio (modified freehand drawings from Franke, 1976, 1977).

188  Reproduction and Development in Annelida

To study the role played by cerebral and PV hormones in Typosyllis pulchra, Heacox and Schroeder (1982) removed either simultaneously prostomium and PV or one of them. In T. pulchra, the stolonization cycle lasts for 30 days and the stolon remains sexually undifferentiated until the 3rd day of the cycle. If prostomium and PV are removed on the 5th day, the stolon continues to undergo oogenesis. This observation is also confirmed by the occurrence of oogenesis in a recipient stock implanted with PV from an undifferentiated donor that has not yet stolonized. However, the removal of prostomium and PV prior to the 5th day induces sex reversal and the former female stock begins to produce stolon, in which spermiogenesis occurs. Further, the production of PV hormone is inhibited, when prostomium is removed during the initial days of stolon cycle. Hence, prostomium has an upper hand in regulation of the PV inhibitory hormone. Weidhase et al. (2016) have further elaborated to know the exact source in and around the PV by amputating at the (i) pharynx, (ii) pharyngeal tube and (iii) middle of the PV in T. antoni. Their findings are listed below: (i) Incomplete anterior regeneration on removal of anterior end of the PV, (ii) Deviation from the usual reproductive pattern, (iii) Accelerated and masculanized stolonization and (iv) Limitation of regeneration of posterior segment. Besides endocrines, density is an important factor in regulation of male ratio of stolons. Typosyllis prolifera flourishes on the thalli of Halopteris scoporia. Of 7,846 stolons surveyed by Franke (1986b), 25 and 27 stolons have been developing gametes of either sex in Porec and Pula populations of Yugoslovia, respectively. Irrespective of increasing density from 5 to 20 stolons/10 g thalli, male ratio of stolons remains around 0.5 in Rovinj population but decreases from 1.0 to 0.5 in Pula population. However, when stocks of these three populations are reared in singles, the ratio remains around 0.5. Hence, density plays an important role in sexually labile T. prolifera. But the level of lability is a population trait. When reared individually also, spontaneous sex change can occur in 2.5 and ~ 5.0% stolons in Rovinj and Pula stocks, respectively. At the 4th stolonization event, male ratio is 0.5 and 1.0 in Rovinj and Pula populations. Hence, Rovinj population is the least labile but that of Pula is the most labile. The spontaneous sex change begins to slowly but steadily increase in successive stolonization events. Consequently, the ratio increases from 20% in the 2nd stolonization event to 100% at the 5th event. Hence, masculanization is rapid and completes at the 6th event. In another two series of experiments, Franke (1986b) investigated the effects of density and combination of partners on the stolon’s male ratio of T. prolifera. The 1st series tested the effect of volume of water and presence of males, when female stolons were reared at 1 ♀/5 ml, 50 ♀♀/150 ml and 50 ♀♀ + 50 ♂♂/150 ml. The results indicated that (i) > 89% of female stolons reared in isolation in limited volume of water sex reversed into males; (ii) the sex reversal was reduced to 20%, when groups of female

Sex Differentiation  189

stolons were reared in larger volume of water; (iii) the presence of primary male stolons further reduced the reversal to 17%. Clearly, the co-presence of female and male stolons reduces spontaneous sex reversal. To study the effects of density and female-male combinations on male ratio during the 2nd, 3rd and 4th stolonization events in the Pula stock, Franke (1986b) has chosen the following three groups of combinations and three subgroups in each of them namely Group A: 2 ♀♀ : 0 ♂, 0 ♀ : 2 ♂♂, 1 ♀ : 1 ♂, Group B: 10 ♀♀ : 0 ♂, 0 ♀ : 10 ♂♂, 5 ♀♀ : 5 ♂♂ and Group C: 20 ♀♀ : 0 ♂, 0 ♀ : 20 ♂♂ and 10 ♀♀ : 10 ♂♂. The recalculated male ratios are plotted against the chosen female-male combinations at increasing density in Group A, B and C (Fig. 7.3A) and successive stolonization events (Fig. 7.3B). Following an initial increase of the ratio in the combinations of 0 ♀ : 2 ♂♂, the ratio progressively decreases, irrespective of the differences in combinations like 0 ♀ : 10 ♂♂ and 0 ♀ : 20 ♂♂. On the other hand, the ratio progressively increases with successive stolonization events but at different levels. The decrease in male ratio is in the following order for the chosen female : male combinations: 0 ♀ : 2 ♂♂ < 0 ♀ : 10 ♂♂ < 0 ♀ : 20 ♂♂. The trends for others in each of the groups fall below the respective ones. From these observations, the following may be inferred: 1. Male ratio increases with successive stolonization events 2. The ratio decreases with increasing density and 3. However, the presence of males alone at any chosen density increases male ratio to a higher level than that in which females alone is present. Hence, the presence of females reduces the male ratio and the reverse is true for the males.

Figure 7.3 Male ratio in stolons of Typosyllis prolifera. A. Effect of different female-male combinations on male ratio during the 2nd, 3rd and 4th stolonization events. B. Effect of successive stolonization events on male ratio in different female-male combinations (drawn using data reported by Franke [1986b] in his Table 5).

190  Reproduction and Development in Annelida

7.3  Pollutants and Reproduction Annelids play an important role in decomposition and bioturbation. In this process, they accumulate the pollutants; in its tissues, Tubifex tubifex may concentrate cadmium by ~ 35 times (Gillis et al., 2002). Not surprisingly, aquatic oligochaetes are reported to have been used in bioassays since the times of Aristotle (Lobo and Espindola, 2014). A fairly large volume of literature is available on this subject. However, this account elucidates selected mosaic representative examples on gametogenesis and reproductive output, regenerative potency and brooding in acidic water as well as inherent recovery ability from cadmium and pCO2 pollutants. For annelids, information related to the negative effects of pollutants is available on survival (e.g. Enchytraeus crypticus, Castro-Ferreira et al., 2012), body size (e.g. T. tubifex, Gillis et al., 2002) and reproductive output (e.g. Neanthes arenaceodentata, Oshida et al., 1981). However, information is not yet available on metabolic pathways through which endocrine disruption occurs. Many insecticides and metallic pollutants are reported to induce (i) spermatogonial damages in Eisenia foetida, (ii) incomplete sperm maturation in Branchiura sowerbyi and (iii) reduce sperm count in Pheretima guillelmii (Table 7.2). In a review, Yasmin and D’Souza (2010) listed the negative effects on growth and reproduction in Eisenia foetida exposed to copper oxychloride, malathion, achetochlor, chloropyrifos, cypernethrin or benomyl. For aquatic annelids, more information is available on cadmium (cd) pollution. Exposure to sedimental cd reduces body weight of T. tubifex from 2.7 mg to 0.9 mg and reproductive output from 15 offspring/adult to 5/adult (Table 7.2). Undertaking a long term (440 days) chronic exposure to trivalent cadmium covering three generations of Neanthes arenaceodentata (Table 7.3), Oshida et al. (1981) have reported the negative effects in the first generation and the possible recovery during the third generation. With increasing dose, the following negative effects are recorded in the parental generation: (i) decrease in the number of spawning pairs (6 to 5), (ii) extended interspawning interval (112 to 123 days), (iii) reduced brood size (255 to 78) and (iv) total offspring production (1,628 to 391). However, the worm recovers during the second regeneration by (i) an increase in spawning pairs (5 to 10), (ii) reduction in inter-spawning interval (123 to 118 days), (iii) increases in brood size (78 to 111) and (iv) total offspring production (391 to 1,112). Ocean acidification: Oceans cover 70% of the earth’s surface and hold 97% of its water and serve to buffer CO2. Thankfully, the daily uptake of atmospheric CO2 by the oceans is 22 million metric tons. Since the advent of the industrial era, oceans have absorbed 127 billion metric tons of carbon as CO2 from atmosphere. Carbon dioxide combines with water chemically. Hydrolysis of CO2 increases the hydrogen ion (H+) concentration with

Sex Differentiation  191

Table 7.2 Effect of pollutants on spermatogenesis and fecundity of annelids Species/References

Reported Observation

Branchiura sowerbyi Casellato et al. (2011)

Treated at (50–100 times above the normal) 0.1–0.3 mg fluoride/l for 5 months inhibits the completion of sperm maturation

Eudrilus eugeniae Yesudhason et al. (2012)

X-rays exposure fragments acrosome in the head and breaks the zig-zag tail

Lumbricus terrestris Cikutova et al. (1993)

Exposure to cadmium reduces sperm count

Eisenia foetida Corpas et al. (1995)

Exposure to lead damages spermatogonia and spermatocytes

Pheretima guillelmii Reinecke and Reinecke (1997)

Chronic exposure to heavy metals reduces sperm production

Hirudinaria manillensis Singhal and Davies (1996)

Exposure to organo-phosphorous insecticide Temphos at > 0.1 mg/l for 22 hours reduces fecundity

Tubifex tubifex Gillis et al. (2002)

Exposure to cadmium reduces body weight from 2.7 mg at sediment concentration of 3.8 µmol/g x 10–3 to 0.9 mg at 5 µmol/g x 10–3 and reproductive output from 15 offspring/ adult in control to 5/adult at 5 µmol/g sediment

concomitant reductions in pH and carbonate ion (CO32–) concentration. The progressive reduction in availability of carbonate (CO32–) renders the acquisition of biogenic calcium carbonate (CaCO3) by calcareous tubiculous worm more difficult and energetically costlier (cf Pandian, 2017). As a result, marine annelids may encounter a series of significant changes in oceanic pCO2 and pH levels in the future. Increases in oceanic pCO2 are predicted to occur at an unprecedented rate, leading to a significant decrease in pH, a phenomenon commonly known as ‘ocean acidification’. However, evolutionary adaptation occurs, when selection on existing genetic variation shifts the average phenotype of a population toward the fitness peak that matches with changing environmental conditions (Rodriquez-Romero et al., 2015) like elevation in pCO2 and declining pH. With a short life span, many polychaetes have served as an ideal model for multi-generational studies indicating initial negative effects and subsequent recovery. In a rare study, Pires et al. (2015) have investigated the effects of increase in temperature and decrease in pH on regenerative potency of Diopatra neapolitana (Table 7.4). Increase in temperature below 24ºC accelerates the regenerative process. But decreases in pH from alkaline to neutral level cause a drastic increase in regeneration duration from 52 to 63 days and decreases segment regenerative potency from 81 at pH 7.8 to 68 at pH 7.1. Rodriquez-Romero et al. (2015) have carried out two series of experiments in the errant Ophryotrocha labronica. In the first one, reproductive traits have

192  Reproduction and Development in Annelida

Table 7.3 Effect of different doses of trivalent cadmium chronic exposure on breeding characteristics in parental generation and recovery in second generation of Neanthes arenaceodentata (condensed and simplified from Oshida et al., 1981) Breeding Characteristics

Control

Cadmium Dose (mg/l) 0.0125

0.05

9 100 133 1199

5 123 78 391

10 130 258 2580

7 111 50 415

10 132 190 1395

10 118 111 1112

Parental generation Spawning pairs (no.) Inter-spawning interval (d) Brood size (no.) Total offspring (no.)

6 112 255 1628 First generation 9 153 292 2628

Spawning pairs (no.) Inter-spawning interval (days) Brood size (no.) Total offspring (no.)

Second generation Spawning pairs (no.) Inter-spawning interval (days) Brood size (no.) Total offspring (no.)

8 129 273 2186

Table 7.4 Effects of temperature on the duration and level of regeneration in Diopatra neapolitana (condensed and simplified from Pires et al., 2015) Temperature (ºC) 17 20 24

Regeneration Completed (d)

Chaetiger (no.)

61 48 43

75 82 89

pH 7.8 7.5 7.3 7.1

Regeneration Completed (d)

Chaetiger (no.)

52 54 57 63

81 77 72 68

been estimated in the worms exposed to elevated (pCO2 = 1,000 µatom) level for seven generations. The second one has involved the transplantation from elevated level to low (pCO2 = 400 µatom) level and vice versa. The biological assay has shown that all the tested traits are recovered and even improved, except the egg size, which has remained constant at 0.6 × 10–3 mm3. The increase for growth is from 0.9 to 1.4 chaetigers per day and for fecundity from 5.13 to 8.55 eggs per chaetiger in the first and seventh

Sex Differentiation  193

generation, respectively (Table 7.5), i.e. the increases represent improvement of 55% for growth and 67% for fecundity. With transplantations, growth rate (1.40 chaetigers per d), adult size (~ 15 chaetigers) and egg size (0.6 × 10–3 mm3) have remained constant. However, there are decreases in survival (from 82% at elevated-elevated transfer to 79% in low to elevated transfer) and fecundity (from 22.3 to 18.7 eggs per chaetiger) (Table 7.5). Studies on calcareous tube-building sedentary polychaetes are desirable, as they may find it difficult and costlier to acquire the required calcium carbonate with increasing pCO2 and decreasing pH. In the field of ocean acidification research, our knowledge is based on experimental studies involving a short or long term exposure to different levels of predicted future pH. In a model investigation, Lucey et al. (2015) have circumvented the limitations related to experimental studies by relating the dominance and distribution of the known polychaete species inhabiting a range of natural acidic waters from extremely low pH to low pH and ambient pH. They have found that on exposure to extreme low pH and low pH, the proportion of brooders within a species progressively increases, especially in interstitial species (Table 7.6). However, a small portion (6–17%) spawns in ‘chemical islands’ characterized by low pH. Broadcasters (e.g. Platynereis dumerilii) and others with short non-feeding pelagic phase (e.g. Pileolaris spp) choose to spawn in open waters with normal ambient pH. Table 7.5 Multi-generational responses of life history traits of Ophryotrocha labronica on exposure to pCO2. Also the effects of transplantation from elevated (pCO2 = 1,000 µatom) to low (pCO2 = 400 µatom) level and vice versa (compiled and simplified from Rodriquez-Romero et al., 2015) Multi-generational responses Trait

F1

F2

F3

F4

F5

F6

F7

Survival (%)

80

83

80

87

90

89

83

Growth rate (chaetigers/d)

0.9

1.3

1.3

1.4

1.5

1.3

1.4

Adult size (no. of chaetigers)

14

15

15

16

16

–

15

Fecundity (no./chaetiger)

5

7

10

10

9

–

9

Egg volume (× 10–3 mm3)

0.6

0.6

0.6

0.6

0.6

–

0.6

Transplant assay Trait

Elevated to Elevated

Elevated to Low

Low to Low

Low to Elevated

Survival (%)

82

78

85

79

Growth rate (chaetigers/d)

1.4

1.4

1.4

1.4

Adult size (no. of chaetigers)

15.4

15.1

15.4

15.0

Fecundity (no./chaetiger)

22.3

19.5

21.4

18.7

Egg volume (x 10–3 mm3)

0.6

0.6

0.6

0.6

194  Reproduction and Development in Annelida

Table 7.6 Adaptive strategies in response to ocean acidification and lowering pH in vent-inhabiting polychaetes (simplified from Lucey et al., 2015) Species/Family

Adaptive Strategy

Abundance (%) at Extreme Low pH

Low pH

Ambient pH

Positive increase in pelagic Platynereis dumerilii Nereididae

Broadcaster, External fertilization, PLK larvae

6

–

94

Pileolaris spp Serpulidae

Brooder, LEC non-feeding larva at pelagic

19

38

43

Spio decoratus Spionidae

Brooder, pelagic/benthic juvenile development

17

17

67

Positive increase in brooding Platynereis massiliensis Neredidae

Brooder, Mucus tube brooding Direct development

91

–

9

Exogone naidina Syllidae

Brooder, Direct development, Interstitial

27

35

38

E. meridionalis Syllidae

Brooder, Direct development, Interstitial

44

39

17

Syllis prolifera Syllidae

Stolonization, Swarming, Benthic fertilized eggs

48

21

21

Rubifabriciola tonerella Fabricidae

Intra-tubular brooding, Direct development

67

21

31

Parafabricia mazzellae Fabricidae

Intra-tubular brooding, Direct development

85

6

9

Brifacia araiargonensis Fabricidae

Intra-tubular brooding, Direct development

74

19

7

7.4  Vectors and Borers In crustaceans (see Pandian, 2016) and molluscs (see Pandian, 2017), parasites are known to disrupt the endocrine sex differentiation process. However, no parasite is reported to disrupt the endocrine sex differentiation in annelids. But many annelids, especially tubificids serve as vectors to transmit diseases. For example, Tubifex tubifex serve as an obligate vector of the sporidean Myxobolus cerebralis, the causative agent of salmonid whirling disease (Stevens et al., 2001).

Sex Differentiation  195

The cosmopolitan spionids Polydora and Boccardia compose a large number of burrowing species of ‘mudworms’. For example, P. websteri damages oysters by burrowing into the shell through boring aided by a viscous fluid, which dissolves the shell. Burrowing on the shell in and around the adductor muscle area by P. ciliata renders the shell to open readily and thereby increases predation of mussels like Mytilus edulis (see Pandian, 2017).

8 Vermiculture

Introduction One objective of this book is to elevate annelids from their academic interest to economic importance. For a long time, the usefulness of some annelids was known but is not adequately recognized. Due to page limitation, this chapter may not elaborate nascent technical details related to (i) rack culture system for Tubifex tubifex (Marian et al., 1989), (ii) culture of sedentary polychaetes (e.g. Sabella spallanzanii, Giangrande et al., 2014b), (iii) regeneration as a novel method for ornamental sabellids (Murray et al., 2013), (iv) compacted culture system (Garcia-Alonso et al., 2013) and (v) ideal method for changing of water from simulated beach set-up (Serebiah, 2015). Rather, it attempts to assemble widely scattered basic information to (i) identify the fast growing candidate species, (ii) assess biomass production in the contexts of (a) nutrients; (b) temperature and (c) rearing density; (iii) distinguish ‘layers’ from ‘brooders’; (iv) estimate offspring production in the candidate species characterized by (a) sexual, (b) parthenogenic and (c) clonal reproduction; and (v) explore the scope for increased biomass production in elevated ploids. Information on this fundamental biological inputs may direct future research on more profitable vermiculture utilizing waste as a resource, and discourage odd researches, in which costlier shrimp meat (e.g. Nielsen et al., 1995) and trout pellet (e.g. Memis et al., 2004) are used as feed in vermiculture. Briefly, ‘wealth from waste’ must be the theme of vermiculture.

8.1  Characteristics and Features Earthworms are known to (i) improve aeration, (ii) drainage, (iii) availability of nutrients to plants and (iv) integrate soil organic and mineral elements (Butt, 1993). For example, more than 80% of litter-fall is decomposed and its nutrients are made available to Salix gigantea plantation (Curry and Bolger,

Vermiculture  197

1984). The vermicomposts increase the yield of commercial crops (e.g. Lablab purpureus, Karmegam and Daniel, 2009, Brassica compensis, Guerrero III, 2009). In recent years, the dramatic increase in demand for polychaetes as baits by anglers is so high that even developed countries like UK and South Korea have begun to aquaculture them in a big way (Olive, 1999, E Costa et al., 2006). The role played by the enchytraeids in composting municipal waste and production of live feed for salmon and sturgeon is known, especially from subantarctic countries like Russia (see Fairchild et al., 2017). In waste water treatment plants, polychaetes (e.g. Marphysa sanguinea, Parandavar et al., 2015) and aquatic oligochaetes (see Ratsak and Verkuijlen, 2006) serve as biological agents to minimize activated sludge. In percolating filter sewage system of UK, Lumbricillus rivalis (incidence frequency, 91%), Enchytraeus buchholzi (57%), E. coronatus (22%), E. albidus (9%) and Fridericia spp (4%) are present. Their density fluctuates but is mostly in the range of 200/l for L. rivalis and > 300/l for E. coronatus (Learner, 1972). Paddy fields: In a very useful investigation, the International Institute of Rice Research has made a survey of the presence and dominance of aquatic oligochaetes in the Philippine ricefields (Simpson et al., 1993). The survey has recorded the following: (i) Nine species of aquatic oligochaetes are present: (a) tubificids: Limnodrilus hoffmeisteri, Branchiura sowerbyi, Aulodrilus limnobius, A. sp, (b) naidids: Dero digitata, Aulophorus hymanae, Pristina sp, (c) enchytraeids: Mesenchytraeus sp and (d) lumbriculid: Lumbriculidae spp, (ii) of them, L. hoffmeisteri (81%) and B. sowerbyi (13%) dominate, (iii) population increases with increasing soil carbon content up to 3.5% and application of nitrogen fertilizer up to 140 kg/ha, (iv) density frequency decreases from 45% at 5,000/m2 to 1% at 35,000/m2 and (v) release of soil organic substances possibly dependent on bacterial decomposition. The occurrence of Aulophorus vagus, A. furcatus and Dero digitata in Indian ricefields has been reported (Hegde and Sreepada, 2014). In sediments, where L. hoffmeisteri flourishes, the processes of bacterial de-nitrification and nitrification occur simultaneously. Aquatic oligochaetes accelerate the release of nitrate (NH4+N) and phosphate (PO43–) from overlying water and soil. However, population density is increased to a greater extend by organic fertilizer than inorganic ones (see Yokota and Kaneko, 2002). The de-nitrification rates are 90 and 50 mg N/m2/d in the presence and absence of L. hoffmeisteri, respectively; the corresponding values for nitrification are 69 and 29 mg N/m2/d (Chatarpaul et al., 1980). Hence, the loss due to de-nitrification is almost completely compensated by nitrification. However, there are other reports indicating the stimulation of sediment nitrification at low density of Tubifex tubifex but inhibition of it at high density (Pelegri and Blackburn, 1995). Research inputs in this field are urgently required to know whether the harvest of L. hoffmeisteri and T. tubifex at appropriate intervals shall reduce the application of nitrogen fertilizer and provide live feed as a byproduct.

198  Reproduction and Development in Annelida

Size and Density: A notable feature is that with decreasing body size of annelids, cultivable density can be increased (Fig. 8.1). Individual size decreases from 7 g in Marphysa sanguinea (Parandavar et al., 2015) and Aporrectodea longa (Butt, 1993) to a few milligrams in the tubificids Branchiura sowerbyi (100–150 mg, Aston, 1968), Tubifex tubifex (7–17 mg, Marian and Pandian, 1984, Finogenova and Lobasheva, 1987) and L. hoffmeisteri (9.4 mg, Nasciomento and Alves, 2009) and to a few micrograms (recalculated highest live weight of aquatic enchytraeids: 900, 300, 240 and 180 µg for Lumbricillus rivalis, Cognettia cognettia, Marionina southerni and Achaeta eiseni, respectively, see Lindegaard et al., 1994). As aeolosomatids weigh < 100 µg, their growth is measured in number alone (e.g. Aeolosoma viride, Falconi et al., 2015). However, cultivable density increases in the following order: 50/m2 in Perionyx ceylanensis (Karmegam and Daniel, 2009) > 2,000/m2 in M. sanguinea > 20,000/m2 in T. tubifex > 35,000/m2 in smaller aquatic oligochaetes. Most remarkably, cultivable density of earthworms is far less than that of most aquatic worms. Differences in food availability and motility to acquire food from the substratum/culture medium may be responsible for the observed low density requirement of earthworms. The highest organic substance, in

Figure 8.1 Effect of body weight on density of some oligochaetes (data assembled from different sources, see text).

Vermiculture  199

which Perinereis spp (Palmer, 2010) and Lumbricus terrestris (Butt et al., 1995) have been reared, is in the range of 4.5% for the former and 3.9% for the latter. Hence, food availability may not be the reason for the low density requirement of earthworms. The reason seems to the denser substratum, through which earthworms have to move to acquire food. Both earthworms and nereidids move by alternate muscular contraction and relaxation. However, the resistance encountered by earthworms against movement in the denser soil substratum is enormous, in comparison to the ‘lose’ sediment submerged in aquatic medium. Consequently, the reported values for motility is 6.7 cm/d for earthworm (e.g. estimated distance of 200 cm/10 months in Aporrectodea longa, Butt et al., 1995) and 7,344 m/d for nereidid (e.g. estimated potential distance of 85 mm/s, see Pandian, 2016). Of course, the nereidid cannot sustain motility for all the 24 hours in a day and the earthworm motility may be altered by organic content and soil texture. Despite reduced motility, earthworms seem to require a larger volume of substratum to acquire food. In low nutrient sediments, polychaetes may switch to acquire dissolved organic substances from the surrounding medium (p 21–22) but earthworms can only increase the gut loading frequency up to 6 times (p 23). As a result, they require a larger volume of substratum. The calculated highest cultivable density values for earthworm range between 50/m2 for P. ceylanensis weighing 0.8 g and 159/m2 for Aporrectodea caliginosa weighing 0.8 g. It is possible to increase density with worms smaller than 0.2 g (Johnston et al., 2014). Taxa and Reproduction: Reproductive modes impose a profound impact on individual growth and biomass production and are important in vermiculture. Of 13,000 and odd polychaete species, only 207 are hermaphrodites and three are parthenogens (Table 8.1). Among 79 clonal species, the majority of them are capable of bidirectional cloning, i.e. a single parent (genet) produces two offsprings (ramets). In the errant stolonizing syllids, a parent is capable of producing up to 18 stolons (ramets). In some tubiculous sabellids, a single genet is shown to produce three to eight ramets (e.g. Potamilla torelli, Sabella pavonina, Sabellastarte spp, see Fig. 3.7, Table 4.9). However, the syllids and Table 8.1 Taxa (no.) and reproductive modes of annelids (compiled from Tables 1.2, 2.4, 2.5, 4.3, 6.1) Taxa

Species

Parthenogens

Ploids

Parthenogens + Ploids

Cloners

Polychaetes

13,002

3

3

–

79

Oligochaetes

3,175 432

5

5

6

0 9

Earthworms

670

6 + 3?

?

?

1,113

?

4

?

0

Naidids

175

?

?

?

88

Aeolosomatids

27

?

?

?

12

Enchytraeids Tubificids

200  Reproduction and Development in Annelida

sabellids have not yet received much attention by aquaculturists. Notably, much efforts are being made to cultivate nereidids, which may or may not be cloners. Among oligochaetes, earthworms are unidirectional cloners, i.e. on fission, only a single ramet is developed and the other dies. However, of 432 earthworm species, 56 of them are parthenogens and five of these parthenogens are polyploids (Table 2.7). These earthworms do not grow as fast as gonochorics. Of 70 enchytraeid species, 6 + 3(?) are parthenogens and 9 cloners. They are multidirectional cloners, i.e. each genet produces as many as eight ramets (e.g. Enchytraeus japonensis) and is smaller in size and weight, in comparison to sexually reproducing enchytraeids. In fact, the fast growing cloning naidids and aeolosomatids may prove to be great biomass producers only from wastewaters. Collection and Harvest: Not many authors have reported methods for collection and harvest of annelids. With decreasing size, they become more and more important. Earthworms are collected by digging and hand picking (e.g. Aporrectodea trapezoides, Fernandez et al., 2010). Terrestrial enchytraeids can be extracted from soil cores up to 10 cm2 area and 6 cm depth by wet funnel method (for details see O’Connor, 1957). Aquatic oligochaetes are collected by sieving through a hand net at depth of 0.5 cm (Ratsak and Verkuijlen, 2006). To collect Dero digitata, a net with 2000 µm copper mesh screen is first sieved to remove large debris and subsequently a net with 300 µm mesh screen is used (Mischke and Griffin, 2011). More worms of Uncinais uncinata are collected, when 100 µm mesh screen is used instead of a 300 µm screen (Lohlein, 1999). Briefly, the mesh size is progressively reduced with decreasing size of aquatic worms. The need for a simple but effective method of harvesting is obvious. In this regard, Marian and Pandian (1984) have made a key finding. T. tubifex is too sensitive to day light. More than 90% of worms remain at the surface between 20 and 24 hours midnight, when they can be sieved and collected. However, an easier and more effective method is to employ the self-assemblage behavior of annelids. In many oligochaetes, self-assemblage is a behavioral response to stress. An exposure to hypoxic water containing 1.5 mg O2/l, the self-assembled T. tubifex forms a ball, enabling the easiest method of harvest. Stressed by immersion into water, Eudrilus eugeniae self-assemble into a ball (Daisy et al., 2016, Fig. 3.6). Research is required to know whether the other aquatic annelids also self-assemble, when stressed by hypoxia. Feeds and ingredients: In aquaculture, feed is a single most important item, as nearly 60% cost is associated with the feed. Many worms show great promise as live feed for fragile early stages and brooders, and ingredient in processed feeds, as well. As feeds, they are more advantageous than the pelleted and pelagic feeds. Their color (white worm, red worm, the tubifex) and wriggle attract predators. They do not impair water quality, when added

Vermiculture  201

to aquaculture system. Being benthic, they remain alive at the bottom and are not easily flushed out from the rearing tanks, as the traditional feeds like artemia nauplii. Some of them are highly productive. During the 1940s, Russian biologists have made an admirable contribution to mass culture of white and red worms. Stalked culture boxes can readily produce 30 kg worm/d (see Fairchild et al., 2017). However, their size, even at hatching, is two-times larger than the first instar artemia larva (Table 8.2). Hence, they may serve as feed more for the larvae of fishes than shrimp. Interestingly, night-crawling earthworm E. eugeniae has been used as bait since the 1940s. Dietary fatty acids and some vitamins are obligately required to ensure optimal growth of finfish and shellfish in aquaculture farms. Of 14% fat present in E. eugeniae, nearly half of it is constituted by saturated and unsaturated fatty acids (Guerrero III, 2009). Available information on levels of fatty acids in Enchytraeus albidus and Nereis virens is listed in Table 8.2. The presence of 14% fat in E. eugeniae and 10–25% lipids in E. albidus are 2 to > 5 times more than that present in commercial pellet feeds (cf Leelatanawit et al., 2014). A comparative study on sperm production in the broodstock of tiger shrimp Penaeus monodon fed on commercial feed and polychaetes has indicated that the sperm cell production is 20 × 103/ml and 37 × 103/ml in the former and latter, respectively (Leelatanawit et al., 2014). At School of Biological Sciences, Madurai Kamaraj University, India, red tilapia, guppy, molly, fighter fish, barbs and tetras have been reared for nearly 15 years. To ensure quality eggs on expected dates, their brooder females obligately require to be fed at least for 7–10 days with tubifex (e.g. Kavumpuruth, Table 8.2 Fatty acid contents (mg/g dry substance) of Enchytraeus albidus and Nereis virens (condensed and compiled from Evjemo and Olsen, 1997, Brown et al., 2011, Fairchild et al., 2017). * coffee ground waste, † stale bread waste 20 : 5n–3

22 : 6n–3

n–3

n–6

SFA

MUFA

PUFA

34 42

154* 62†

36

246

22

66

27

119

Enchytraeus albidus (1.0–1.5 mm at hatch) 5 6

1 0

11 11

126 39

52 33

Artemia franciscana (0.42–0.52 mm, I instar) 36

22

80

9

22

Calanus finmarchicus (0.22–0.79 mm) 9

18

31

4

9

Brachionus plicatilis (0.13–0.34 mm) 13

18

62

4

9

Nereis virens 16.0

16.1

18 : 2ω6

20 : 1ω6

20 : 4ω6

20 : 5ω3

22 : 1ω11

22 : 1ω3

22

8

12

10

2

12

12

6

202  Reproduction and Development in Annelida

1992). Incidentally, many annelids are capable of synthesizing essential fatty acids and vitamins or they engage symbiotic microbes to synthesize them. The presence of many microbes in the gut of many annelids is reported (e.g. Sruthy et al., 2013). Subramanian et al. (2017) have demonstrated that E. eugeniae engage Bacillus endophyticus to synthesize riboflavin to facilitate regeneration (see p 108, 110). Protein content of annelids ranges between ~ 59% in E. albidus (Fairchild et al., 2017) and 65% in earthworms (Guerrero III, 2009). Vermimeal can replace fishmeal as an ingredient and as protein source for Oreochromis niloticus and Macrobrachium idella (Guerrero III, 2009). Studies on the vermicost of E. eugeniae, using biodegradable waste for culture, have indicated that the efficiency and cost-effectiveness in reducing chemical fertilizer for crop cultivation up to 100% (Guerrero III, 2009). Rearing Atlantic halibut in a recirculating culture system with N. virens in raceway water containing pellet waste, fecal waste or both, Brown et al. (2011) have estimated the cost-profit of the polyculture system. For every unit of feed input at US$1.04, the halibut fetches US$9.5 plus 0.3 kg sludge, which produces 0.11 kg N. virens valued at US$3.3.

8.2  Candidate Species To identify the fast growing candidate species for vermiculture, information on growth-age relationship is required. Unfortunately, description of life history characteristics is limited to 3% of polychaetes (p 41). As a result, information available for polychaete is limited, while a reasonable amount of information is available for earthworms, enchytraeids and tubificids with relatively longer history of their cultivation. Growth and reproduction are important factors in biomass production and are considered separately.

8.2.1 Growth Earthworms: In earthworms, there is a lime-lag between sexual maturity and cocoon laying (see also Fig. 8.9A). Their growth trends are more or less sigmoidal. At the end of log-phase of growth, cocoon production commences. Subsequently, their body weight may be retained, as in Aporrectodea trapezoides (Fig. 8.2A) or decreased, as in Eisenia foetida (Fig. 8.2C). Considering 150 days rearing period, growth achieved by the investigated earthworm decreases in the following order: A. longa > Eudrilus eugeniae > Lumbricus terrestris > E. foetida > A. trapezoides and Octolasion cyaneum > Perionyx ceylanensis > Hyperiodrilus africanus. Fed on paper pulp and reared singly at 20ºC, A. longa grows to a size of 6.5 g within 150 days. Reared in group of two to four and fed on waste dung manure at 25ºC, E. eugeniae grows to 4.4 g within the same period of

Vermiculture  203

150 days. Both of them are ‘layers’, as they are sexually mature but have not yet commenced laying cocoons. At 30ºC, E. eugeniae grows to 2.3 g on the 60th day (Dominguez et al., 2001) and is likely to reach ~ 4.2 g on the 150th day (Fig. 8.2B). Hence, A. longa may be more suited to temperate zone and E. eugeniae to tropics. Notably, the Spanish E. eugeniae grows significantly faster, even when reared in a group, but the South African grown individually achieves 3.5 g body weight at 25ºC (Viljoen and Reincke, 1989). A search for an E. eugeniae that achieves super-fast growth is required for tropical vermiculture. It is known that temperate animals grow larger than their tropical counterparts. Not surprisingly, the tropical earthworms like E. foetida, P. ceylanensis and H. africanus do not attain a large size, as temperate earthworms. However, it is not clear why the growth rate of tropical earthworms is at abysmal slow rate. As moisture plays an important role in sustaining growth (Fig. 8.2E), it is not clear whether the relatively lower moisture content prevailing in humid tropical soil (e.g. 10–40% in most soils of Tripura, India, Bhattacharjee and Chaudhuri, 2002) reduces growth rate. Expectedly, growth is accelerated up to 20ºC in L. terrestris and 25ºC in E. eugeniae (Fig. 8.2B). Earthworms survive and grow on litter and/or waste manure. E. foetida grows nearly two-times faster, when fed on cow dung manure than on dry leaves (Fig. 8.2C). Hitherto, semi-moist, half-digested dung of cow, horse or pig have been found to sustain growth of earthworms. However, fecal pellets of goats and sheep do not support cocoon laying in them (Siddique et al., 2005). Raw soil, which supports poor growth, can be enriched with saw dust and coconut husk. The ‘skin’ of coconut pod contains fiber (used for rope manufacture) and husk. The husk is not burnable but retains moisture for a long duration. In coconut-cultivating Asian countries, the coconut husk is a waste and remains as environmental hazard. Hence, research is required to estimate the moisture retention capacity and enriching ability of raw soil by coconut husk. Apart from these environmental factors, growth of earthworms can considerably be altered by internal factors like parthenogenesis and polyploidy. O. cyaneum is a parthenogen (see Table 2.5) and a pentaploid (see Table 6.1). Likewise, A. trapezoides is also a parthenogen and occurs as diploid or triploid (Table 6.1). Unfortunately, Fernandez et al. (2010) have not indicated the ploidy status of A. trapezoides. Metaphire houlleti is suspected as parthenogen (Kaushal et al., 1999, not shown in Fig. 8.2A). The growth trends of sexually reproducing diploid A. longa and E. eugeniae are considerably at higher levels than those of parthenogens. Hence, parthenogenic earthworms are not good candidate species for vermiculture. Polychaetes: Within polychaetes, researchers have concentrated mostly on commercially valuable (as bait) nereidids. A few nereidids are semelparous, while others are iteroparous. The former allocate much of available resource

204  Reproduction and Development in Annelida

Figure 8.2 Growth of selected earthworms A. Hyperiodrilus africanus (Tondoh and Lavelle, 1997), Perionyx ceylanensis (Karmegam and Daniel, 2009), Octolasion cyaneum (Lowe and Butt, 2008), Eisenia foetida (Siddique et al., 2005), Lumbricus terrestris and Aporrectodea longa (Butt, 1993), A. trapezoides (Fernandez et al., 2010) and Eudrilus eugeniae (Dominguez et al., 2001) B. Effect of temperature on L. terrestris (Lowe and Butt, 2005) and E. eugeniae (Dominguez et al., 2001) C. soil and manure on E. foetida (Siddique et al., 2005), D. density on P. ceylanensis (Karmegam and Daniel, 2009), O. cyaneum and L. terrestris + Allolobophora caliginosa (Curry and Bolger, 1984) and E. soil moisture on E. foetida (Reinecke and Venter, 1987) (figures are all redrawn).

Vermiculture  205

on gamete production, whereas the latter allocate bulk of resource on somatic growth (Cassai and Prevedelli, 1998b). Irrespective of any of them, harvest of ‘layers’ is to be made prior to sexual maturity, i.e. prior to the commencement of allocation to gametic production. Growth of polychaetes has been measured as body length in units of cm (e.g. Glycera dibranchiata, Fig. 8.3A) or number of segments (e.g. Perinereis rullieri, Fig. 8.3D) or weight (e.g. Marphysa sanguinea, Fig. 8.3B), making comparison difficult. In natural habitats, Nereis virens requires 2 years to attain sexual maturity at the size of 35 cm body length (Creaser and Clifford, 1982). Olive (1999) has suggested that optimal rearing conditions and feeding can reduce this period to 1 year. Hence, many attempts have been made to use costly enriched feeds. For example, the feeding regime of Neanthes arenaceodentata consists of initial inoculation of 15 mg dry powder of green alga Enteromorpha sp, subsequent addition of 15 mg powder on the 44th, 51st and 70th days and costly prawn flakes thrice weekly (Pesch et al., 1987). The others have gone for high protein fish feed pellets (hpffp) for M. sanguinea (Parandavar et al., 2015) and shrimp meat for N. virens (Nielsen et al., 1995). In fact, sedimented pellets and feces due to excess feeding are reported to reduce feeding in Pseudopolydora kempi japonica (Miller and Jumars, 1986). Figure 8.3B shows that the fastest growing M. sanguinea attains a body weight of 7 g fed on hpffp at the density of 500/m2. Considering 150 days period in the investigated polychaetes, growth decreases in the following order: N. virens > M. sanguinea > Perinereis nuntia (Fig. 8.3A–C). These worms have been fed on uneaten pellet + halibut fecal waste, hpffp or waste water containing ~ 4% organic matter, respectively. As in earthworms, increase in temperature accelerates growth of polychaetes (Fig. 8.3D); in P. rullieri, a reduction in ration decreases growth (Fig. 8.3E) and animal (nauplii) or protein enriched diet (e.g. N. aranaceodentata) enhances growth better than algal tetramin (Fig. 8.3D). Clearly, nereidids require relatively protein-rich feed than earthworms. With substratum containing 3–4% organic substance, earthworms achieve 4–6 g body weight but polychaetes > 1 g (e.g. Perinereis spp, Fig. 8.3C). With increasing protein enrichment in diet, the production cost of nereidids shall be higher than that of earthworms. Briefly, the waste water recirculating system of Brown et al. (2011) with an area of 12.6 m2 containing 1,110 worms (i.e. 407 g/0.37 g initial size), i.e. 87 no./m2 N. virens reared in waste water containing uneaten pellet and halibut feces seem to be the optimal system and best candidate species to produce 2 g sized worm within 80 days. Tubificids: The live feed tubificids are potentially important in commercial vermiculture. Branchiura sowerbyi, Tubifex tubifex, Limnodrilus sp and Lumbriculus sp can produce ~ 15 kg/m2 (Lietz, 1987). B. sowerbyi: The relatively large sized B. sowerbyi is a deserving cultivable candidate. It grows to 150 mg in 180 days (Fig. 8.4A). However, it attains

206  Reproduction and Development in Annelida

Figure 8.3 Growth in selected polychaetes. A. Glycera dibranchiata (drawn from data reported by Creaser, 1973) and Nereis virens (drawn considering about 35 cm growth in 2 y, Creaser and Clifford, 1982). B. Marphysa sanguinea (growth data pooled from 0.5, 1.0 and 2.0 g size reared at 500 or 2000 worms/m2, compiled and redrawn from Parandavar et al., 2015) and Nereis virens fed on waste in recirculating culture system (Brown et al., 2011). C. Perinereis nuntia and P. helleri in a sand filter system (redrawn from Palmer, 2010). D. Effect of temperature and food quality on growth of Perinereis rullieri (compiled and redrawn from Prevedelli, 1992) and Neanthes arenaceodentata (in dotted line, redrawn from Pesch et al., 1987). E. Effect of ration on growth of 400 mg weighing Nereis virens, redrawn from Nielsen et al., 1995).

Vermiculture  207

Figure 8.4 Growth and reproductive output in Branchiura sowerbyi. A. Growth as function of age (drawn from information reported by Aston, 1968). B. Growth and cocoon production as function of age (compiled and redrawn from Lobo and Alves, 2011). C. Effect of temperature on growth (redrawn from Aston et al., 1982). D. Effect of nutrients (cellulose content) on growth and egg production (compiled and redrawn from Aston, 1984).

maturity at the age of 30 days and subsequently lays cocoons almost once a week in two temporally separated batches (Fig. 8.4B); the first batch lasts from the 35th to the 190th day and the second from the 217th to the 360th day. More cocoons and eggs (see Fig. 2.8B) are laid in the first batch. Hence, harvest of ‘layers’ can be made on the 28th day at ~ 30 mg size and ‘brooders’ on the 190th day. As in other worms, elevation in temperature accelerates growth up to 25ºC (Fig. 8.4C). Notable is that it grows to 30 mg size within 30 days, in comparison to 7–17 mg size of T. tubifex (Fig. 8.5A). Further, B. sowerbyi is more herbivorous than T. tubifex, as it can grow to 40 mg size within 21 days, when fed on filter paper containing 6–9% cellulose (Fig. 8.4C). T. tubifex grows to a size of 7 and 17 mg, when reared on cow dung manure and activated sludge, respectively (Fig. 4.5A). Marian and Pandian (1985) have noted the entry of chironomus into the open continuous flow system. With increasing chironomus density from 0 to 7/cm2, the number and production of tubifex are reduced from 146 to 10 no./cm2 and 181 to 13 mg/cm2. Improving the culture system with recirculating water system and rack culture tanks covered to prevent the entry of chironomus and mosquitoes, Marian et al. (1989) have reduced water consumption from 38,000 l to 193 l with a production of 5.6 kg tubifex/mo. In Bangladesh, Jewel et al. (2016) have also reared tubifex with cow dung supplemented with yeast and the like. Their production system has yielded less tubifex than that of Marian and Pandian (1984). Hossain et al. (2011) have also reared tubifex feeding on costlier combination of 30% soybean, 20% mustard oil, 20% wheat bran, 20% cow dung and 10% sand and have achieved tubifex production of 659 mg/cm2. Clearly, protein requirement of tubifex is higher than that of B. sowerbyi (see Fig. 8.4D). Oplinger et al. (2011) have made a series of experiments to estimate the effects of feed type, ration, temperature and density on juvenile and biomass production. However, initial stocking level, which is known to have a profound effect on production, was not kept uniform; for example, it was 31 in cow dung manure, instead of 50 in feed type experiment. It also ranged

208  Reproduction and Development in Annelida

Figure 8.5 A. Tubifex tubifex growth as function of age in worms reared in activated sludge in Leningrad, Russia and natural silt (from Finogenova and Lobasheva, 1987), cow dung manure in Madurai, India (from Marian and Pandian, 1984) and lettuce waste (from Kosiorek, 1974). Dark shade indicates the cocoon biomass. B. Effects of (B1) food quality, (B2) food quantity, (B3) temperature and (B4) density on juvenile and biomass production (drawn from recalculated data reported by Oplinger et al., 2011).

from 166 to 397 worms, instead of 50 in temperature series experiment. Secondly, tubifex was reared in circular beakers and biomass production was reported in mg/mg stock, instead of reporting in mg/cm2. Hence, the results are not comparable with those of others. Thirdly, with elevation of temperature from 18ºC to 27ºC, both juvenile and biomass production was reported to decrease (Fig. 8.5B3). T. tubifex is cosmopolitan and almost all the worms belonging to different annelid taxa are reported to accelerate growth with elevation of temperature. As far as possible, their data were recalculated and summarized in Fig. 8.4B. With increasing protein enriched feed type and ration, the number of juvenile production is increased but only up to 0.4–0.5/ adult/d; however, the increase is upto 0.7/adult/d with increasing density and the decrease from 2.7/adult/d with increasing temperature. Biomass production is also increased in the following order: increasing density < increasing ration < decreasing temperature < feed type. Briefly, stocking density has an overriding role in biomass production of tubifex. Enchytraeids: Of 670 species, nine are clonal and another nine may be parthenogens (Table 8.1). A few like Enchytraeus bigeminus are parthenogens and polyploids (Christensen, 1973). Most enchytraeids are terrestrial and more abundant in arctic and temperate zones (density: > 100,000/m2) than in humid tropics (< 10,000/m2) (Schmelz et al., 2013). Their density can also reach 21,500/m2 for Marionina southerni in Lake Esrow, Denmark (Lindegaard et al., 1994) and as high as 300,000/m2 in moorland (see Gonclaves et al., 2017). Their body weight ranges from 15–240 µg in M. southerni (Lindegaard et al., 1994) to 770 µg in E. bigeminus (Christensen, 1973). As they are too

Vermiculture  209

small, their growth is measured in body length or number of segments or mostly in numbers. As mentioned earlier, clonals are more of academic interest and gonochorics are of economic importance. Life span of cloners is half of that of gonochores. For example, it is 15 days in E. japonensis in cloners but 29 days in gonochores. When the worm grows to 60–80 segments size, it spontaneously autotomizes into 5–10 fragments and each one of them is regenerated into a complete worm in 4–5 days but the ramet’s size is reduced to < 50% (Yoshida-Noro and Tochinai, 2010). Reared in 200 ml jar containing sand and abundant porridge, E. bigeminus grows to a maximum size of 770 µg within 2–3 weeks. Initially, it clonally reproduces until the density of 300–350 is reached, when it switches to sexual reproduction. In natural habitats, clonal reproduction occurs during favorable conditions but with a commencement of winter, it switches to sexual reproduction. In a 200 ml jar, the maximum biomass production during 8 weeks experiments can be 2.2 kg (Christensen, 1973). Incidentally, density also imposes profound effect on body size and juvenile production in F0 and F1 generations of E. crypticus. With increasing density, reductions are 50% in body size and 75% in juvenile production (Gonclaves et al., 2017). Information on growth of tropical aquatic enchytraeids is not available. Available information pertains to temperate enchytraeids thriving in activated sludge arising from sewage waters. Considering a 50-day period, growth of the investigated worms decreases in the following order: Lumbricillus rivalis < L. lineatus < E. albidus < E. crypticus < E. coronatus (Fig. 8.6A). In almost all these worms, temperature accelerates growth up to 20ºC (Fig. 8.6B). As no decline in the acceleration is observed in Lumbricillus lineatus even at the 20ºC, it may grow even faster up to 30ºC. Biomass production of Cognettia sphagnetorum is ~ 200 worms per day (Springett, 1970). In Lake Esrow, annual net production of M. southerni is in the range of 5.1 Kcal/m2/y (Lindegaard et al., 1994). Doubling time (Dt): Gravimetric or linear measurements to estimate growth in smaller worms become increasingly difficult, time-consuming and requires sophisticated balance. Growth can also be measured in numbers or specific growth/intrinsic growth rate. But the latter suffers from the following: (i) it decreases with increasing body size (e.g. Dero dorsalis, Ratsak and Verkuijlen, 2006). It may not be possible to fix a rate for a species and compare it with another species, as their body lengths may differ considerably. (ii) it can not be used in clonal species, as they only regenerate the missing body parts. A better parameter is the doubling time. Dt represents the period from the day, when sexual or clonal reproduction is commenced to that day, when F1 offspring begins to reproduce. An analysis of available data suggests that body size and temperature impose profound effects on Dt. Expectedly, Dt increases with increasing size in enchytraeids and tubificids. For example, Dt is < 3.61 days in relatively larger (7–17 mg) tubifex than in the smallest enchytraeids

210  Reproduction and Development in Annelida

Figure 8.6 Growth of selected gonochoric enchytraeids. A. Growth as function of age at 20ºC in gonochoric Lumbricillus rivalis, Enchytraes albidus (o represents data reported by Reynoldson, 1943), E. coronatus and E. buchholzi (in dotted line) in sewage percolating filters (redrawn from Learner, 1972), L. lineatus (redrawn from Reynoldson, 1943) and E. crypticus (redrawn from Bicho, 2015). Cross bars indicate the size and age at sexual maturity. B. Effect of temperature on growth of body length L. lineatus, L. rivalis, E. coronatus (redrawn from Learner, 1972).

(< 2 days). It requires the longest duration of 21 days in the largest (100–150 mg) Branchiura sowerbyi (Fig. 8.7A). It is known that 54% of naidids are cloners (p 127); hence, they are considered separately. In the naidids too, a linear direct relationship between body size and Dt becomes apparent (Fig. 8.7B); irrespective of sexually or clonally reproducing species within relatively shorter temperature ranges from 20ºC to 30ºC. Also, irrespective of clonal or sexual reproduction, naidids are capable of biomass production. In them, Dt increases from 2 days in the smallest gonochoric Chaetogaster langi at 20ºC to 3.3 days in Dero dorsalis at 30ºC and > 6 days in the largest cloner Nais elinguis. However, relatively a wider range of temperature does influence Dt

Figure 8.7 A. Doubling time in some aquatic oligochaetes. B. Doubling time in naidids, G = gonochore, C = cloner, ● = 20ºC, ■ = 25ºC and ▲ = 30ºC (drawn from data reported by Ratsak and Verkuijlen, 2006). C. Doubling time as function of temperature in enchytraeids and B. sowerbyi (drawn from data reported by Aston et al., 1982).

Vermiculture  211

Table 8.3 Effect of density on successful pairing, reproduction and fecundity in Neanthes arenaceodentata (condensed from Pesch et al., 1987) Trait

Density (no./840 cm2) Control

Survival (%)

40

80

160

97

90

81

74

0.54

0.55

0.56

0.53

Age at spawning (d)

85

101

102

109

Paired reproducing female (%)

54

54

49

23

Fecundity (egg no./brood)

695

881

622

598

Sex ratio

Paired but (a) not successful females (%)

5.3

0

0.8

0.5

(b) laid eggs (%)

5.1

5.5

13.0

11.0

(c) no egg laying (%)

35

41

37

66

in enchytraeids and tubificids. With increasing temperature, Dt decreases but at different levels. In Lumbricillus rivalis, it decreases from 45 days at 8ºC to 11 days at 20ºC; the corresponding values for Enchytraeus coronatus are 24 days at 8ºC and 11 days at 20ºC. For B. sowerbyi, it increases from 11 days at 25ºC to 20 days at 33ºC but decreases from 21 days at 21ºC. Apparently, the optimum for Dt is 25ºC (Fig. 8.7C). From this analysis too, B. sowerbyi and T. tubifex are suggested as more suitable candidate species.

8.2.2 Reproduction In polychaetes, nereidids are cultivated. They are all gonochores and are not cloners. Some of them like Nereis virens are epitokous and semelparous (Table 5.1), while others like Marphysa sanguinea are atokous and iteroparous. Information on their reproduction, especially fecundity has been elaborated in Chapter 2. From the point of maintaining brooders, only one publication is available. A vast majority of oligochaetes are hermaphrodites. In them, a few are parthenogens (e.g. Lumbricus terrestris, Table 2.5) and a very few are parthenogens cum polyploids (e.g. Dendrodrilus rubidus, Lumbricillus lineatus, Tables 2.7, 6.1). For maintenance of brooders in oligochaetes, relatively more information is available. Polychaetes: Pesch et al. (1987) have investigated the effect of density on pair formation and reproduction in Neanthes arenaceodentata. With increasing density from 40, 80, and 160/840 cm2, i.e. 48, 95 and 190 no./m2, survival, male ratio and age at spawning are marginally affected (Table 8.3). However, the number of successfully paired females is decreased from 54% to 23% and fecundity from 881 eggs/brood to 598 eggs/brood at the highest density of 160 no./840 cm2. Further, the number of females, which are

212  Reproduction and Development in Annelida

unable to pair (0 to 0.8%) and which are unable to lay eggs (41 to 66%) is significantly increased. Considering the control levels, maintaining brooders of N. arenaceodentata at ~ 50 no./m2 is the recommended density. It must be noted that N. arenaceodentata is a fairly small nereidid. Nereis virens, the recommended candidate nereidids, grows to > 40 cm (Fig. 8.3A). Hence, the density at which larger brooding nereidids can be maintained may be in the range of only a few. However, research inputs are required. Oligochaetes: During their life time, oligochaetes lay cocoons in two temporally separated batches. The second batch contains less number of eggs/cocoon than the first one (Fig. 8.8A). Further, fertility of eggs decreases at elevated

Figure 8.8 Reproduction in earthworms. A. Monthly and cumulative cocoon production in Lumbricus terrestris (simplified and redrawn from Butt et al., 1994). B. Effect of feeds on cocoon production in Eisenia foetida (simplified and redrawn from Siddique et al., 2005). Window shows the effect of moisture on cocoon production in E. foetida (redrawn from Reinecke and Venter, 1987). C. Effect of culture density on number and biomass of cocoon production in Metaphire houlleti (compiled and redrawn from Kaushal et al., 1999). D. Effect of temperature on cocoon production and hatching success in Eudrilus eugeniae (drawn from data reported by Dominguez et al., 2001).

Vermiculture  213

temperature (Fig. 8.9D). Hence, the maintenance of brooders beyond 2 years in Lumbricus terrestris (Fig. 8.8A), 190 days in Branchiura sowerbyi (Fig. 8.4B) and 60 days in Eisenia foetida (Fig. 8.8B) is not recommended. With advancing age, fecundity is considerably decreased and fertility of egg is also decreased. Considering the respective initial values and 50% decrease in fecundity with advancing age, it is suggested that brooders older than 1 year in L. terrestris (Fig. 8.8A) and 50 days in Lumbricillus rivalis (Fig. 8.9B) are not maintained. Food quality and moisture play an important role in cocoon production. Cow dung manure and maintenance of 75% moisture facilitate production of more number of cocoons, especially in tropical earthworms, as in Eisenia

Figure 8.9 Reproduction in aquatic oligochaetes. A. Sexual maturity and generation time (discontinuous line) as a function of age at different temperatures in Lumbricillus rivalis (thick lines) and Enchytraeus albidus (drawn from data reported by Learner, 1972). B. Fecundity as function of age in L. rivalis (drawn from data reported by Kirk, 1971). C. Effect of temperature on fecundity of Branchiura sowerbyi (redrawn from Aston et al., 1982). D. Effect of temperature on cocoon production and egg fertility in L. lineatus and E. albidus (drawn from data reported by Reynoldson, 1943).

214  Reproduction and Development in Annelida

foetida (Fig. 8.8B). Incidentally, E. foetida brooders older than 80 days may not be as productive as the young ones. Irrespective of rearing in single or batch, the suspected parthenogen Metaphire houlleti invests 8 mg on cocoon biomass (Fig. 8.8C). As age advances from 75 days to 350 days in Perionyx ceylanensis too, the cocoon production rate (no./worm/d) also decreases from 1.15 to 1.00 in singles and 1.25 to 1.15 in batches of eight worms (Karmegam and Daniel, 2009). Hence, density seems not to affect cocoon production. However, more research inputs are required. In general, oligochaetes are sensitive to temperature. The period required for sexual maturity decreases with increasing temperature in Lumbricillus rivalis and E. albidus. However, there is a time-lag between sexual maturity and the day, on which the first cocoon is laid, i.e. generation time. This timelag between maturity and generation time is maintained with increasing temperature, though the trend is linear in L. rivalis but L-shaped in E. albidus (Fig. 8.9A). The thermal optima for brooders is 25ºC for Eudrilus eugeniae (Fig. 8.8D), 29ºC for B. sowerbyi (Fig. 8.9C) and 15ºC for L. lineatus and E. albidus (Fig. 8.9D). Of course, these two enchytraeids are temperate and subarctic species. A feature that deserves to be noted is that the optimum temperature for somatic growth differs from that for cocoon production. For example, it is 25ºC for growth in B. sowerbyi (Fig. 8.4C) but 29ºC for cocoon production (Fig. 8.9C). However, it is 28ºC for growth in E. albidus and L. lineatus (Fig. 8.6B) but 15ºC for cocoon production (Fig. 8.9D). For E. eugeniae, 25ºC is the optimum for both growth and reproduction (Fig. 8.2B, 8.8D). Oligochaete embryos are enclosed in eggs, which, in turn, are enclosed in a cocoon. No information is available on the mechanism of hatching, and the time course and sequence, through which hatching occurs. Firstly, fertility of eggs is an important factor in determining hatching success. It decreases on either side of the optimum temperature in L. lineatus and E. albidus (Fig. 8.9D). Clearly, on either side of 15ºC, fertilizability of eggs and sperm is reduced. The presence of infertile eggs within cocoons may be responsible for the reported decrease in hatching success of Eudrilus eugeniae on either side of 25ºC, the optimum temperature (Fig. 8.8D). In view of wide differences in hatching success reported for oligochaetes, information on cocoon structure and hatching mechanism is required. Considering earthworms, for which relatively more information is available, hatching success ranges for 20% in Eutyphoeus gammiei to 100% in Metaphire houlleti fed on moist filter paper. In E. foetida too, it is 54 and 63% in worms fed on cow manure and dry leaves. It is not clear whether the water content of feed and/or desiccation plays a role in determining hatching success. For cocoons of M. houlleti, which have been fed on moist filter paper or incubated in distilled water, the success is 100%. It is then difficult to comprehend 70–79% hatching success reported for the cocoons of Aporrectodea longa, L. terrestris and Octolasion cyaneum incubated in water at 15ºC, especially 47% success reported for A. longa cocoons incubated in water medium at 20ºC. Hence, attempts have been made to relate cocoon

Vermiculture  215

Figure 8.10 Effect of cocoon size (in numbers) and incubation period (in alphabets) on hatching success of earthworms. 1/A Perionyx excavatus, 2/B Pentoscolex corethrurus, 3/C Lampito mauritii, 4/D Polypheretima elongata, 5/E Dichogaster modigilianii, 6/F Drawida nepalensis (P), 7/G Eutyphoeus gammiei (data from Bhattacharjee and Chaudhuri, 2002), 8/H Metaphire houlleti (P?) (data from Kaushal et al., 1999), 9/I Aporrectodea longa, 10/J Lumbricus terrestris (P) and 11/K Octolasion cyaneum (data from Butt, 1993).

size and hatching success as well as incubation period and hatching success (Fig. 8.10). On plotting, all the available values are found scattered widely. Still, cocoon size and incubation period seem to determine the level of hatching success. The longer the incubation, the possibilities for desiccation and/or predation are greater. Hence, incubation of cocoon in safe aquatic medium is recommended.

8.2.3  Polyploids and Parthenogens In animals, cell volume of triploids and polyploids triply and multiply increases; but, the maximum attainable body size by polyploids is regulated by reducing the number of cells (e.g. fishes, Pandian, 2011). However, both cell volume and cell number are increased in polyploid bivalves (Pandian, 2017). As eggs of oligochaetes are enclosed in a cocoon, ploidy induction is difficult to achieve. However, studies on cell number and cell volume in natural polyploid annelids are required to know whether or not the cell volume and number are increased. The ensuing information seems to demand research input in this area. Bimastos eiseni and B. tenuis exist only as diploid and triploid, respectively. So are Eisenia venata and E. rosea. Dendrobaena rubida exist only as diploid but its sister species D. subrucunda

216  Reproduction and Development in Annelida

exists only as tetraploid. In Octolasion, O. lacteum exists only as diploid, while its sister species O. cyaneum as polyploid. Interesting information on their growth has been assembled (Table 8.4). Triploid B. tenuis grows to a larger length (52 mm), width (3 mm) and with more number of segments (105), in comparison to diploid B. eiseni. Body lengths of triploid B. tenuis (10%), E. rosea (56%), tetraploid D. subrucunda (40%) and polyploid O. cyaneum (26%) grow to larger sizes than their respective diploid sister species. It is not clear whether triploid Bimastos, Eisenia, tetraploid Dendrobaena and Octolasion grow both in cell volume and number. Notably, E. rosea triploid grows 1.6 times larger than its diploid counterpart E. venata. Hence, these two earthworms may be subjected to studies on cell volume and number Table 8.4 Body size in some diploid and polyploidy earthworms (modified from Muldal, 1952) Species

Ploidy

Length (mm)

Diameter (mm)

Segment (no.)

Bimastos eiseni

2n

47

2–5

93

B. tenuis

3n

52

3

105

Eisenia venata

2n

34

3–4

102

E. rosea

3n

55

3–4

135

Dendrobaena rubida

2n

43

3.5

75

D. subrucunda

4n

60

4

85

Octolasion lacteum

2n

97

4

126

O. cyaneum

9n

122

7.5

127

Table 8.5 Polyploidy, parthenogenesis and reproduction in Tubifex tubifex and Limnodrilus hoffmeisteri (inferences from Poddubnaya, 1984) Traits

Semelparous

Iteroparous

Tubifex tubifex Life span (days)

100

245

Age of maturity (days)

55

60

380 72

Reproductive cycle (no.)

1

2–3

2–6

Fecundity (egg no./cycle)

36

40

23

% in population

60

40

60 349

Limnodrilus hoffmeisteri Life span (days)

135

181

Age of maturity (days)

78

90

66

Reproductive cycle (no.)

1

2–5

2–4

Fecundity (egg no./cycle)

65

35

64

% in population

27

69

78

Vermiculture  217

on a priority. If found that the triploid E. rosea grows by increasing both cell number and cell volume, as in bivalves, then it is a matter of great academic and economic importance. Rearing parthenogenic Tubifex tubifex and Limnodrilus hoffmeisteri for two years, the former Soviet scientist Poddubnaya (1984) has briefly summarized the results in Table 1 of the publication. The results reported therein are confusing and difficult to understand. However, two groups in each species are recognizable with a short and long life span. Describing fecundity of the Italian T. tubifex, Pasteris et al. (1996) has recognized two cohorts with production of less and more number of eggs (see Fig. 2.8D, E). Investigating the ploidy status of the Italian T. tubifex, Marotta et al. (2014) have reported that the chromosome number of T. tubifex can be 2n = 50, 4n = 100 and 6n = 150. It is in this context, that an attempt has been made to understand and infer some information from Table 1 of Poddubnaya. The life span is 100, 245 and 380 days for T. tubifex and 135, 181 and 349 days for L. hoffmeisteri (Table 8.5). Those with the shortest life span have a single reproductive cycle. Hence, these diploids may be semelparous. The other two groups have two to five reproductive cycles and are iteroparous. Fecundity per reproductive cycle is reduced to ~ 50% in the long living T. tubifex and in L. hoffmeisteri. It is likely that they are tetraploids. However, more research on the effect of ploidy on growth and reproduction is urgently required, especially in the commercially valuable T. tubifex.

9 Summary and New Findings

As in other invertebrates, taxonomy of annelids is in a fluid but dynamic state. The number of described species, which has remained ~ 2,000–3,000 until the 19th century, has now grown to 17,000. Correspondingly, publications on polychaetes alone have also increased from a dozen/decade until 1960 to ~ 300/decade during 2,000–2,010 (Fig. 1.2). Yet, description of life history characteristics is limited to 3% of polychaetes and is increasing at the rate of three species/y. Some 81, 12 and 7% annelids are marine, freshwater and terrestrial inhabitants. Vertical distribution of the soft-bodied annelids ranges from 4,900 m depth (tubificid, Abyssidrilus stilus) to 2,000 m altitude (enchytraeid, Buchholzia appendicularia). They thrive in unusual habitats like hydrothermal vents (siboglinids) and sub-terranean aquatic system (stigobiont oligochaetes belonging to 42 genera in 27 families). For the first time, relevant information has been highlighted on gutless oligochaetes and polychaetes, osmotrophism and anaerobiosis in some annelids. For example, constantly migrating between the nutrient-rich anoxic and oxic zones in the sediments, the gutless tubificids have successfully colonized an ecological niche, so far unoccupied by any other interstitial fauna (Giere et al., 1984). A vast majority of polychaetes are gonochores, whereas almost all clitellates are hermaphrodites characterized by internal fertilization and laying cocoon containing a few eggs. Polychaetes display fascinating and incredibly diverse reproductive modes. From his extensive survey, Wilson (1991) has estimated that 47% of polychaetes brood their eggs and remaining 53% are broadcast spawners. Another estimate has indicated 42% are brooders and 58% are broadcasters. In 45% of polychaetes, fertilization is external; it is also external in another 9% of brooders. With hermaphroditism in clitellates, gonochorism is limited to 76% of annelids. Hermaphroditism occurs also in 23 families of polychaetes. From their survey, Schroeder and Hermans (1975) have reported the incidence of hermaphroditism in 67 polychaete species. Without validating and/or updating the report of Schroeder and Hermans, many reviewers have virtually repeated the same number of 67 hermaphroditic species. From an intensive survey of literature up to 2017, the number is updated to 207 species (Table 2.4). For the first

Summary and New Findings  219

time, a directory has been documented for the incidence of parthenogenesis in 75 annelid species, of which 56 are earthworms (Table 2.5). In annelids, fecundity decreases with advancing age in Lumbricus terrestris (Fig. 2.8A), and in tube-dwelling Streblospio benedicti (Fig. 2.8H) or increases with advancing age and body weight but beyond a particular age/body weight, it decreases in Tubifex tubifex (Fig. 2.8D, E). It is not clear whether with increasing body length, the girth grows differently in these worms or they undergo reproductive senescence at different ages and sizes. The flattened body in Ophryotrocha puerilis puerilis and dorso-ventrally compressed body in hirudineans provide a relatively larger surface area than that in cylindrical annelids; in the hirudineans and O. puerilis puerilis, fecundity continues to increase with increasing body size (Fig. 2.7). Besides body size and egg size, sexuality and oogenic anlage are shown to affect fecundity. Poecilogony is a rare reproductive mode reported only from polychaetes and opisthobranch molluscs. Whereas it is limited to two alternative morphs in a half a dozen opisthobranchs, the boccardian polychaetes exhibit three different morphs. In a reciprocal mating system, each partner fertilizes the eggs of the other. Amazingly, the partners simultaneously change sex several times in their life time in O. puerilis puerilis to ensure that no partner cheats the other. The account on regeneration has brought to light a whole range of new findings. Among annelids, hirudineans are not capable of either regeneration of missing body parts/organs or clonal reproduction; a few annelids are capable of regeneration of the entire anterior or posterior or anterior cum posterior fragment of the body. For example, the head of Lumbriculus variegatus can be regenerated 21 times and the tail 42 times and both together 20 times. Posterior regeneration is induced by neurosecretion from the brain in errant polychaetes. However, the ventral ganglia are capable of inducing posterior regeneration in sedentary polychaetes, whose projected ‘heads’ are more often subjected to sub-lethal predation. Notably, regeneration in annelids is a fairly complex but orderly process and proceeds from woundhealing to blastema formation and differentiation, segmental reorganization, and growth and elongation. It ranges from species that has no ability to regenerate even a single segment (e.g. Streblospio benedicti) to species that are capable of regenerating an entire worm from a single seminal (newly coined term) segment (e.g. Chaetopterus variopedatus). In polychaetes and oligochaetes, ectodermal, mesodermal and endodermal regeneration involves the injury-activated, dedifferentiated multipotent stem cells arising from the respective germ layers. Oligochaetes with chloragogue and polychaetes without it exhibit contrasting features. In chloragogue, reserves are inadequate to simultaneously meet the costs of both regeneration and reproduction and are temporally separated. But the sedentary polychaetes undertake them together at the cost of reduced reproduction. In annelids, the origin and loss of regenerative potency have occurred independently at multiple numbers of times (Bely, 2010). Hence, the seminal segments are

220  Reproduction and Development in Annelida

positioned at different locations in different taxonomic groups albeit mostly between the cephalic region and anterior trunk. The assemblage of incidences of regeneration in annelids by Zattara and Bely (2016) has paved the way for further analysis. The number of species capable of anterior, posterior and anterior cum posterior regeneration is 149, 206 and 144, respectively, i.e. only 0.88, 1.22 and 0.85% of annelids are capable of anterior, posterior and anterior cum posterior regeneration. The reasons for the less number of incidences of anterior regeneration are: (i) the anterior contains vital organs like the brain, heart and metanephridia and (ii) as new segments are regenerated from the pygidial zone, the regenerative potency diminishes faster in the ‘older’ anterior segments than in the ‘young’ posterior segments. When the respective number of these regeneratives is considered as fractions of 13,012 polychaete and 3,175 oligochaete species, the percentage values indicate that the potency is 1.5–2.0 times more prevalent in oligochaetes (1.57, 1.80, 1.42) than the respective ones (0.88, 1.22, 0.85) of polychaetes. In annelids, clonal reproduction is characterized by the following general features: (i) Reared under optimal conditions, a dozen annelid species are sustained by clonal reproduction alone up to 30–60 years. However, the Primordial Germ Cells (PGCs) can be transmitted upto 1,000–3,000 generations in the clones (e.g. Pristina leidyi). When stressed or induced, the cloners switch to sexual reproduction. (ii) Abundant food supply, low density and favorable temperature may either singly or in combination trigger clonal reproduction. (iii) Except for a couple of species, the duration required for completion of the sensitive cloning ranges between 3 and 8 days (Table 4.5). (iv) Intense predation alone may not manifest clonal reproduction; for example, of 290 tubiculous sabellids species, only 17 are cloners. (v) Unlike in echinoderms (Pandian, 2018), hermaphroditism and brooding in annelids do not prevent clonal reproduction. However, larval cloning is not yet reported in annelids. (vi) In clonal species, sex ratio is altered and gamete production is reduced. (vii) Clonal reproduction is usually succeeded by sexual reproduction but both can occur simultaneously in a very few naidids. (viii) Clonal reproduction can be grouped into architomy and paratomy. In architomy, fission is followed by the completion of regeneration and formation of new progeny (ramet). But it occurs even before the ramets are fully formed in paratomy. Assigning clonal species to their respective families in relation to architomy and paratomy has revealed the following new findings: (i) Of 100 and odd annelid families, clonal reproduction occurs only in 12 polychaete and five oligochaete families (Table 4.3). Architomy occurs exclusively in eight polychaete families but paratomy in aeolosomatids alone. (ii) In clonal reproduction, the incidence ranges from 2% of spionids to 54% of naidids. (iii) Further, it is limited to 79 polychaete species but to as many as 111 oligochaete species (Table 4.3), i.e. cloning occurs only in 0.61% of

Summary and New Findings  221

polychaetes but 2.14% of oligochaetes. Hence, clonal potency is ~ 4.6 times more prevalent in oligochaetes than that of polychaetes. Analyzing the data of clonal species, Zattara and Bely (2016) have viewed that cloning may have been derived from regeneration. This view may not be correct for following reasons: (a) Cloning in 111 oligochaete species do obligately requires the presence of neoblasts (Myohara, 2012). In the absence of neoblasts, almost all earthworms, many enchytraeids (e.g. Enchytraeus buchholzi, Myohara, 2012), naidids and tubificids are unable to clonally reproduce. (b) Without having the ability to regenerate anterior cum posterior regeneration, Chaetogaster diaphanus and C. diastrophus reproduce clonally. Clearly, the clonal potency of oligochaetes is vested with neoblasts and not from anterior cum posterior regenerative potency. (c) Even with anterior cum posterior regenerative potency, 31 out of 63 polychaete species do not reproduce clonally (Table 3.8). Hence, a search for the equivalent of neoblasts-like multipotent stem cells in polychaetes is required. Available information on the possible anlage for the clonal stem cells is assembled in Table 4.9 and 4.10. In majority of sedentary/tubiculous sabellids and serpulids, these cells are located deep at the posterior end, irrespective of architomic or paratomic fission. This is also true of the errant dorvilleid Parougia bermudensis and aeolosomatid Aeolosoma viride, and possibly the spionid Pygospio elegans. With the presence of neoblasts in all the non-cephalic segments, the segments are capable of clonal reproduction in E. japonensis and L. variegatus. Due to progressive loss of neoblasts in Stylaria lacustris and their equivalents in polychaetes, the stem cells are restricted to the seminal segments or to the mid-body. However, investigations are required to discover the equivalent of neoblasts in polychaetes. Epitoky is unique to errant polychaetes. The epitokes are divided into epigamics and schizogamics. In epigamy, the entire body of the worm undergoes epitokous transformation but only a fraction of it in schizogamics. As a result, the former is semelparous but the latter iteroparous. For the first time, a directory is documented listing the epigamic incidence in 62 species from 12 families (Table 5.1) and schizogamic incidence in 45 syllid species (Table 5.3). Hitherto, not a single author or reviewer has ever considered the energy demanding vertical migration over distances. Again for the first time, widely scattered information on vertical distance travelled by 15 epigamics and 13 schizogamics has been assembled (Table 5.4). Surprisingly, the vertical distance climbed by the epitokes decreases with increasing body size (Fig. 5.3). Obviously, the larger epitokes glycerids, nereidids and eunicids utilize muscular energy to climb up < 50 m distance. The smaller phyllodocids and ctenodrilids may engage reduction in buoyancy to migrate the vertical distance of > 100–4,000 m. This also holds true for schizogamics. In the epitokes swarming is timed by a combination of annual, lunar and diel rhythms. Temperature and photoperiod induce the formation of heteronereis in epigamics and stolon in schizogamics. The subsequent events are all induced by specific pheromones (Fig. 5.5).

222  Reproduction and Development in Annelida

In annelids, sex is determined by genes harbored on one or more chromosomes. Karyotypic heterogametism is described in five polychaete species. Selective breeding has led to the discovery of heterogametism in Dinophilus gyrociliatus (XX-XO) and Capitella capitata (ZW-ZZ) (Table 6.1). For the first time, a directory is assembled for chromosome numbers in annelids. By selective fertilization of large eggs by X-carrying sperm, D. gyrociliatus females have nullified the chromosomal mechanism of sex determination. In C. capitata, the expression of W gene(s) is relatively more stable and not amenable to environmental factors like density. But the gene(s) in Z chromosome are amenable. Consequently, phenotypic ZZ females and ZZ hermaphrodites are generated. The low level of gametic compatibility between Hediste spp and high level of incompatibility between two morphs of Spirobranchus polycerus and that for the distantly located populations in Galeolaria caespitosa can be explained by the role played by bindin present in the acrosome as an adhesive to attach the sperm to the vitelline layers of eggs. Despite having a well-developed circulatory system, annelids do not possess a glandular hormonal system. Our understanding of endocrine sexualization and regulation of reproduction cycle is based on temperate polychaetes alone. About a dozen neuroendrocrines/hormones secreted mostly by the brain are responsible for regulation of the reproductive cycle. The role played by stolonization inhibiting hormone arising from the proventriculus is responsible for sexualization in syllids. But its action can be reversed by prostominal hormone. The negative effects of cadmium and elevated pCO2 are neutralized in the third generation of polychaetes. Elevation of annelids from an academic interest to economic importance and ‘wealth from waste’ in vermiculture has been one of the objectives of this book. Possibly engaging symbiotic microbes, the annelids produce valuable fatty acids and vitamins, which cannot be synthesized by finfishes and shellfish. Hence, the worms can serve as valuable live feed in aquaculture. In the ricefields, tubificids and naidids play a complex role in nitrification and denitrification processes. Harvesting them at appropriate intervals may reduce the application of nitrogen fertilizer. Research inputs are urgently required in this vitally important field of rice cultivation. The need for research inputs on basic information on growth and reproduction in cultivable worms is emphasized. An early harvest of ‘layers’ is recommended. With minimal genetic diversity, parthenogens and cloners are not good candidates for vermiculture. Secondly, cloners rapidly increase in numbers but may not in biomass production. The fast growing Aporrectodea longa and Eudrilus eugeniae are recommended earthworm species for vermiculture in temperate and tropical zones, respectively. They can be harvested as baits at 4.2 and 2.3 g size on the 60th day (Fig. 8.2A). Cultivation of Nereis virens in wastewater containing waste feed and fecal waste in aquaculture system produces 2 g sized worms within 80 days. In rack culture system with water containing

Summary and New Findings  223

activated sludge or aerated sewage or cow dung manure, cultivation of Tubifex tubifex is profitable. For the first time Branchiura sowerbyi is recognized as a candidate species for vermiculture, as it can be reared in water containing waste paper and harvested at 30 mg size on the 30th day at 25ºC (Fig. 8.4D). Among enchytraeids, Lumbriculus spp and Enchytraeus albidus grow fastest at 20ºC. A search for suitable tropical enchytraeids is urgently needed. Oligochaetes lay cocoons in two temporally separated batches (Fig. 8.8A). The second batch contains less number of eggs than the first one. Hence, maintenance of oligochaete brooders till the day, when laying of the first batch of cocoons, is more profitable.

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Author Index

A Adiyodi, K.G., 35 Aguado, M.T., 2, 3, 5, 9, 121 Aguiar, T.M., 157 Ahearn, G.A., 22, Aiyer, K.S.P., 129, 139 Akesson, B., 29, 70, 88, 91, 126, 128, 130–131, 136, 138, 175, 177 Akesson, B., see Ocklemann, K.W. Alikunhi, K.H., 35 Allen, E.J., 128, 161 Alves, R.G., see Lobo, H. Alves, R.G., see Nasciomento, H.L.S. Andries, J.C., 168, 179–180 Anger, V., 126 Appleby, A.G., 23 Arias, A., 54, 56, 62, 128, 155, 157–158 Arnofsky, P.L., see Blake, J.A. Arp, A.J., 18 Aston, R.J., 198, 207, 211, 213

B Bahl, K.N., 25 Bain, B.A., see Govedich, F.R. Balavione, G., 111 Baldo, L., 67 Banse, K., 20 Barnes, R., 2, 5–6, 54, 56–57, 102 Bastida, R., see Martin, J.P. Bateman, A.J., 87 Beatty, R.A., 65 Beckmann, M., 52 Belov, V.V., see Britayev, T.A. Bely, A.E., 93–94, 98, 105–106, 116–117, 119–120, 123, 129, 219 Bely, A.E., see Ozpolat, D. Bely, A.E., see Zattara, E.E. Bentley, M.G., 181, 184 Bentley, M.G., see Olive, P.J.W., Williams, M.E., Watson, G.J.

Berglund, A., 49, 80, 87, 89 Berkeley, E., 128 Berkeley, C., see Berkeley, E. Berrill, N.J., 93, 95, 106–107, 113–114, 146, 156, 161 Bhattacharjee, G., 72, 203, 215 Bhaud, M.R., 51 Bicho, R.C., 209 Biggers, W.J., 52 Bilello, A.A., 107, 110 Bisby, F.A., 156, 159 Blackburn, T.H., see Pelegri, S.P. Blake, J.A., 9, 11–13, 32, 34, 36, 38–39, 46, 128, 131 Blakemore, J., 66 Bleidorn, C., 5, 6, 9 Boi, S., 33 Boilly, B., 94, 111 Boilly-Marer, Y, 167 Bolger, T., see Curry, J.P. Bomfleur, B., 50 Bondarenko, N.A., 10 Bonifazi, A., 3 Bonney, D.G., see Clark, R.B. Bouguenec, V., 29–30, 81, 104, 129, 133, 136, 138 Bowden, N., see Knight-Jones, P. Bridges, T.S., 78 Bridges, T.S., see Levin, L.A. Brinkhurst, R.O., 10, 30, 66, 141 Brinkhurst, R.O., see Appleby, A.G. Britayev, T.A., 81 Britayev, T.A., see Tzetlin, A.B. Brown, N., 202, 206–207 Bryan, J.P., 52 Bull, J.J., 177 Bullock, T.H., 111 Bunke, D., 56 Butt, K.R., 196, 198–199, 204, 212, 215 Butt, K.R., see Lowe, C.N. Buzhinskaja, G.N., 128 Bybee, D.R., 64

258  Reproduction and Development in Annelida

C Cammen, L.M., 21, 23, 25–26 Campbell, M., 145 Cannarsa, E., 45, 87, 89 Cardon, C., see Porchet, M. Carson, H.S., 37, 41–42, 155 Casellato, S., 191 Caspers, H., 3, 42, 164, 166 Cassai, C., 74–75, 78, 80, 82, 153, 157, 205 Castro-Ferreira, M.P., 190 Cavanaugh, C.M., 16, 18 Cazaux, C., see Bhaud, M.R. Chapman, A.D., 2, 5 Charnov, E.L., 87 Chatarpaul, L., 197 Chatelain, E.H., 153, 157 Chatelliers, M.C., 6, 12 Chaudhuri, P.S., see Bhattacharjee, G. Chevaldonne, P., 16, 19 Chia, F.S., see Qian, P.Y. Childress, J.J., see Arp, A.J. Cho, S.J., 149 Christensen, B., 66–67, 117, 126, 129, 132–133, 136, 139, 171, 208–209 Chu, J.W., 70 Chu, J., 115, 150 Cikutova, 191 Cinar, M.E., see Sahin, G.K. Clark, R.B., 111 Clavier, J., 98, 103, 119, 134 Clifford, D.A., see Creaser, E.P. Coates, K.A., 69, 171 Collado, R., 117, 129 Collado, R., see Schmelz, R.M. Collins, H.L., see Holmstrand, L.L. Cornec, J.P., 106–107 Corpas, I., 191 Costlow, J.D., see Akesson, B. Costopulos, J.J., 22 Cowart, J.D., 48 Creaser, E.P., 4, 58, 75, 78, 80, 82, 153, 156–158, 164, 166, 205–206 Cuomo, M.C., 52 Currie, D.R., 58 Curry, J.P., 196, 204

D D’Souza, D., see Yasmin, S. Dabral, M., see Joshi, N. Daisy, N.P., 112, 200 Daly, J.M., 53–54, 70, 73–74, 76–77, 82–83, 153, 159, 163, 166 Daniel, T., see Karmegam, N.

Dash, H.K., 23 Dash, M.C., 20–21, 23, 25–26 Dash, M.C., see Senapati, B.K. David, A.A., 92, 134–136 Davies, R.W., 20, 29, 77, 79–80, 172 Davies, R.W., see Singhal, R.N. Davila-Jimenez, Y., 58, 119, 121, 128, 131, 133–134, 136 Davison, A., 63 De Vlas, J., 100, 103 Dean, H.K., 11, 153, 156, 163 Dean, H., see Tovar-Hernandez, M.A. Decaens, T., see Jimenez, J.J. Degraer, S., 156 Dhainaut, A., 180 Dhainaut-Cortois, N., see Fewou, J. Diaz-Cosin, D.J., 56, 67–68, 71, 87 Dixon, D.R., 20, 26–27, 59–61 Dominguez, J., 87, 203–204, 212 Drewes, C.D., 145–146 Drewes, C.D., see Lesuik, N.M. Dualan, I.V., 92, 99 Dubilier, N., 14, 52 Durchon, M., 162, 179, 181–182, 186

E E Costa, P.F., 197 Eckelbarger, K.J., 31–32, 44, 54, 58, 75, 78 Engelstadtler, J., 67, 125 Ernst, W., 22 Erseus, C., 6, 11, 14, 32 Erseus, C., see Giere, O. Espindola, E.L.G., see Lobo, H. Evans, J.P., see Marshall, D.J. Evjemo, J.O., 201

F Fage, L., 155 Fairchild, E.A., 197, 201–202 Falconi, R., 117, 126, 129, 145–148, 150, 198 Faulkner, G.H., 140, 150 Faulwetter, S., 5 Fauvel, P., 156, 159 Felbeck, H., 18 Ferguson, J.C., 21–22 Fernandez, R., 29, 72, 200, 203–204 Ferraguti, M., 2, 6, 32–33, 127 Ferraguti, M., see Baldo, L., Boi, S., Jamieson, B.G.M. Fewou, J., 182 Fiege, D., see Wehe, T. Fielde, A.M., 97 Finogenova, N.P., 198, 208

Author Index  259 

Fischer, A., 35, 84, 120, 153, 157–159, 163, 166, 180 Fischer, U., see Fischer, A. Fischer, A., see Hauenschild, C., Fischer, E.A.,  87 Fishelson, L., 156 Fitzhugh, K., see Rouse, G. Fong, P.P., 29, 54, 62, 78, 81, 170 Forbes, V.E., see Ramskov, T. Fourtner, C.R., see Drewes, C.D. Fox, D.L., see Ruby, E.G. Franke, H.D., 90, 120, 127–128, 140, 144, 154, 157, 159–162, 165–168, 181, 186–189 Franke, H.D., see Kahmann, D.

G Gambi, M.C., 71, 74, 82, 126, 128 Garcia-Alonso, J., 157, 181–182, 196 Garraffoni, A.R.S., 40, 58, 60–61, 73 Garwood, P.R., 180 Gaston, G.R., 153, 159, 163, 166–168 Gates, G.E., 67, 97, 103, 118 Gazave, E., 148 Gelder, S.R., see Brinkhurst, R.O. Ghiselin, M.T., 89 Giangrande, A., 41, 49, 63, 74, 113, 155, 196 Giangrande, A., see Knight-Jones, P. Giani, N., see Bouguenec, V. Gibbs, P.E., 156 Gibson, G.D., 46, 84, 86, 93, 103, 116, 121, 128, 131, 133, 135–137 Gibson, G.D., see Smith, H.L., Mackay, J. Gibson, P.H., 66, 121, 128, 131, 133–134, 136, 156 Giere, O., 4, 6, 14–15, 218 Gil, J., see Martin, D. Gillis, P.L., 190–191 Gilpin-Brown, J.B., 157–158, 166 Goerke, H., 20 Goerke, H., see Ernst, W. Golding, D.W., 101–102, 158, 162, 180–181 Gomme, J., see Ahearn, G.A. Gonclaves, M.F.M., 208–209 Gordon, D.C., 24 Goss, R.J., 107 Govedich, F.R., 2, 4, 6 Gremare, A., 10, 26, 59–61 Greve, W., 33 Griffin, M.J., see Mischke, C.C. Grimm, R., 9 Grunig, C., see Pfannenstiel, H.D. Guerin, J.P., 84 Guerrero III, R.D., 197, 201–202 Gusso, C.C., 156

H Hadfield, M.G., 52 Hafer, J., 32 Hall, J., see Gaston, G.R. Halt, M.N., 128 Hamond, R., 162 Hanafiah, Z., 157–158 Hardege, J.D., 44, 153, 168 Harder, T., 52 Harley, M.B., 24 Harms, J., 156, 159 Harper, E.H., 102, 104 Hartman, O., 10 Hauenschild, C., 66, 90, 111, 156–158, 160, 180, 186 Hauenschild, A., see Hauenschild, C., Schierwater, B. Havey, J.M.L., see Gibson, G.D. Heacox, A.E., 101–102, 188 Hegde, P.R., 197 Hempelmann, F., 146 Henshaw, J.M., 43 Hentschel, B., see Carson, H.S. Heppell, S., see Bridges, T.S. Herlant-Meewis, H., 107, 145 Hermans, C.O., see Schroeder, P.C. Herpin, R., 166 Heuer, C.M., 182 Hilario, A., 19, 43 Hill, E., see Tsuji, F.I. Hill, S.D., 105–107, 110–111 Hirabayshi, K., 132 Hochachka, P.W., 25 Hodgkin, J., 179 Hofmann, D.K., 112, 120, 156, 158, 164, 166–168, 182 Holmstrand, L.L., 79, 82 Horridge, G.A., see Bullock, T.H. Hossain, A., 207 Howie, D.I.D., 181, 184 Hsieh, H.L., 43, 45, 54, 64 Huggett, D.V., see Levin, L.A. Hunt, H.R., 114 Hurtado, L.A., 16, 19 Hutcheson, M.S., see Pockilngton, P. Hutchings, P., 4, 5, 6 Hyman, L.H., 105, 106, 118, 136, 139

I Ikeda, M., see Sato, M. Inoue, S., 159, 166 Iori, D., 49 Irvine, S.Q., 46 Ivleva, I.V., 20

260  Reproduction and Development in Annelida

J Jablonka, E., 173 Jaenike, R.J., 66–67, 69, 172 James, R.J., 52 Jamieson, B.G.M., 34, 107 Jayachandran, P.R., 157–158, 166–167 Jenni, K., 84 Jewel, A.S., 207 Jimenez, J.J., 12 Johnston, A.S.A., 12, 199 Jollivet, D., 58, 78 Jones, M.L., 18 Jones, P.C.T., 21 Jorgensen, C.B., 22 Jorgensen, N.O.G., 22 Joshi, N., 27 Jumars, P.A., see Miller, D.C.

K Kahmann, D., 169 Kalidas, R.M., 108 Kaneko, N., see Yokota, H. Karaseva, N.P., 17 Karmegam, N., 197–198, 204, 214 Kaster, J.L., 24 Kato, K., 107 Kaushal, B.R., 203, 212, 215 Kavumpuruth, S., 201 Kawamoto, S., 106 Khan, R.A., 29 Kharin, A.V., 113, 118, 121, 132, 145 King, R.S., 106 Kirchman, D., 52 Kirk, R.G, 213 Kluge, B., 48 Knight-Jones, P., 83, 128 Kinght-Jones, P., see Nogueira, J.M.M., Tovar-Hernandez, M.A. Kobayashi, G., 17 Koene, J.M., 35, 45 Kolbasova, G.D., 127–128, 130, 133–137 Korablev, V.P., 171, 173 Kosiorek, D., 208 Kostyuchenko, R.P., 94 Koya, Y., 157 Kozin, V.V., 148 Kristensen, E., see Jorgensen, N.O.G. Kudenov, J.D., 100, 103, 121, 128, 134, 136 Kupriyanova, E.K., 40, 42–43, 64, 131, 171 Kutschera, U., 51, 63, 82

L Lamb, A., 159 Lamb, M.J., see Jablonka, E.

Lamont, P., see O’Connor, R.J. Lanfranco, M., 177 Langhammer, H., 114 Lardicci, C., 78 Lassalle, B., see Boilly-Marer, Y. Lassarre, P., 55 Lau, K.K., 52 Laufer, H., see Biggers, W.J. Lavelle, P., see Tondoh, E.J. Lawrence, A.J., 179–183 Lawrence, A.J., see Olive, P.J.W. Learner, M.A., 197, 209, 213 Leelatanawit, R., 201 Legendre, R., see Fage, L. Lesuik, N.M., 106 Levin, L.A., 29, 41, 78, 84–85 Levin, L.A., see Chu, J.W. Levinton, J.S., see Martinez, D.E. Lhomme, M.F., see Boilly-Marer, Y. Licciano, M., 114 Liebmann, E., 96, 102, 104, 113, 118 Lietz, D.M., 205 Lindegaard, C., 30, 198, 209–210 Lindsay, S.M., 13, 20, 92, 98, 100–101, 103, 106 Lobasheva, T.M., see Finogenova, N.P. Lobo, H., 30, 79, 81, 190, 207 Loesel, R., see Heuer, C.M. Lohlein, B., 28, 127, 129, 142, 200 Londano-Mesa, M.H., see Salazar-Vellajo, S.I. Lopez, E., 14 Lopez-Jamar, E., 156 Lowe, C.N., 68, 72, 79, 81, 204 Lucey, N.M., 193–194 Lummel, L.A.E., 159, 166 Lutz, R.A., 16

M Macdonald, T.A., 157 Mackay, J., 46 Mangum, C.P., 26 Marian, M.P., 4, 112, 196, 198, 200, 207–208 Marian, M.P., see Pandian, T.J. Markert, R.E., 166 Marotta, R., 19, 171, 217 Marsden, J.R., 52, 64, 174 Marsh, A.G., 17–18 Marshall, D.J., 45 Martin, A.M.S., 13 Martin, D., 156–157, 159, 164 Martin, E.A., 114, 121, 135–136 Martin, J.P., 29 Martin, P., 2, 6, 9, 10, 11, 12, 132 Martinez-Acosta, V.G., 92–93, 98, 121, 135 Martinez, D.E., 83, 126, 138, 162 Martinez, V.G., 93, 118, 129 Matsushima, O., 182

Author Index  261 

Mayer, A.G., 156 Mba, C.C., 72 McEuen, F.S., 44, 131 McHugh, D., 17, 39, 58, 60–61, 75, 78, 82 McLoughlin, N.J., see Davies, R.W. Memis, D., 196 Mercier, A., 87 Micaletto, G., 13, 73 Miller, D.C., 103, 205 Minelli, A., 2 Minetti, C., 90, 176 Mischke, C.C., 200 Miyamoto, N., 95 Mollison, J.E., see Jones, P.C.T. Moment, G.B., 100–101, 113–114, 118 Monahan, R.K., 89 Monroy, F., 34, 44 Moreno, R.A., 13 Morgan, T.H., 92 Morgulis, S., 93, 103, 115, 117–118, 132, 134–136, 150 Muldal, S., 171–172, 216 Muller, C., 92 Muller, M., 94, 111 Muller, M.C.M., see Paulus, T. Muller, M.C.M., see Westheide, W. Mullineaux, L.S., 19 Murray, J.M., 114, 116, 121, 128, 135–136, 196 Muruganatham, M., 157, 166 Mutayoba, S., see Newrkla, P. Myohara, M., 93–94, 110, 115–117, 119, 121, 123, 132–133, 136, 145, 150, 221

N Naidu, K.V., 127, 129, 141–142, 144, 171 Narita, T., 12, 20, 96 Nasciomento, H.L.S., 198 Nayar, K.K., 111 Needham, A.E., 111 Negm-Eldin, M.M., 21 Neuhoff, H.G., 24 Newmark, P.A., see King, R.S. Newrkla, P., 12 Nielsen, A.M., 196, 205–206 Nilsson, P., 129, 138 Nishi, E., 58, 128, 140 Nishihira, M., see Nishi, E. Niwa, N., 113 Nogueira, J.M.M., 128 Nogueria, J.M., see Radashevsky, V.I. Novikova, E.L., 113 Novo, M., 87 Nygren, A., 159 Nyholm, S.V., 16, 18

O O’Brien, J.P., 118 Obenat, S.M., 58 Ocklemann, K.W., 29 O’Connor, F.B., 200 O’Connor, R.J., 52 Okada, Y.K., 100, 108, 114, 159 Okrzesik, J., 113 Olive, P.J.W., 4, 31, 60, 155, 157, 180–184, 197, 205 Olive, P.J.W., see Garwood, P.R., Gremare, A., Lawrence, A.J. Oliver, J.S., 127–128, 131, 133–134, 136 Olsen, Y., see Evjemo, J.O. Oplinger, R.W., 208 Osani, K., see Sato, M. Oshida, P.S., 190, 192 Osman, I.H., 58 Ozpolat, D., 103, 126–127, 138, 148–149

P Paavo, B., 29, 79, 81 Pai, S., see Chu, J. Palmer, P.J., 199, 206 Pandian, T.J., 1, 4, 8, 10, 14, 16, 20–21, 24–26, 31, 47–50, 59, 61, 63–64, 70–71, 83–84, 92, 94, 122, 126–127, 131, 144, 155, 175, 177, 179, 182, 184, 191, 194–195, 199, 215, 220 Pandian, T.J., see Marian, M.P. Parandavar, H., 197–198, 205–206 Pasteris, A., 30, 79, 81, 217 Paterson, I.G., see Gibson, G.D. Patra, U.C., see Dash, M.C. Paulus, T., 106–108, 110-–111, 122 Pawlik, J.R., 52 Pearse, J.S., see Fong, P.P. Pechenik, J.A., see Qian, P.Y. Pelegri, S.P., 197 Penners, A., 111 Pesch, G.G., 171, 173, 205–206, 211 Pesch, C.E., see Pesch, G.G. Petersen, C.W., see Fischer, E.A. Petersen, M.E., 135, 156 Petersen, S., 22 Peterson, D.L., 71, 82 Petraitis, P.S., 58, 90, 171, 173, 176 Pezzani, S.E., see Obenat, S.M. Pfannenstiel, L.D., 33, 100, 129, 179, 181–182, 184–185 Pillai, G., see Olive, P.J.W. Pires, A., 98–99, 101, 191–192 Pleijel, F., see Rouse, G.W. Plyuscheva, M., 81 Pockilngton, P., 159

262  Reproduction and Development in Annelida

Poddubnaya, T.L., 28, 30, 66–67, 216–217 Policansky, D., 63, 89 Pollack, H., 24 Porchet, M., 180, 182 Porto, P.G., 46 Postwald, H.E., see Bilello, A.A. Pradillon, F., 18 Premoli, M.C., 27, 49, 57, 63, 72, 87–89, 170, 177–178 Preston, R.L., 22 Preston, R.L., see Stevens, B.R. Prevedelli, D., 38–40, 58, 70, 79, 81, 91, 171, 173, 176, 206 Prevedelli, D., see Cassai, C., Simonini, R. Probst, G., 108 Putter, A., 21 Purscke, G., 57, 128, 136 Purschke, G., see Westheide, W.

Q Qian, P.Y., 51–52 Qian, P.Y., see Harder, T., Lau, K.K., Qiu, J.W. Qiu, J.W., 89 Quiroz-Martinez, B., 10

R Radashevsky, V.I., 103, 128, 136 Ram, J.L., 168 Ramella, L., see Sella, G. Ramskov, T., 73 Randolph, H., 105–107, 111, 117, 119 Rasmussen, E., 131 Ratsak, C.H., 197, 200, 209–211 Read, G.B., 48, 84, 157, 166 Rebscher, N., 149 Reinecke, S.A., 191 Reinecke, A.J., 204, 212 Reinecke, A.J., see Reincke, S.A., Viljoen, S.A. Reish, D.J., 22 Rettob, M., 82 Reynolds, J.W., 66–67 Reynoldson, J.B., 209, 213 Rhoden, C., 6 Rice, S.A., 35 Rice, S.A., see Akesson, B. Rioja, E., 128 Rodriquez-Romero, A., 191, 193 Rolando, A., see Lanfranco, M. Rossi, A.M., 50 Rota, E., 9 Rouse, G.W., 5, 17, 32–33, 38–40, 46–47, 60–61 Rower, G., 157 Ruby, E.G., 26 Ruppert, E.E., 159 Rychel, A.L., 94

S Safarik, M., 92 Sahin, G.K., 7, 9, 10, 12 Salazar-Vellajo, S.I., 156–157, 159, 164 Samuel, S.C.J.R., 14, 106, 108–110 Santos, C.S.G., see Aguado, M.T. Sato, M., 48, 58, 75, 80, 84, 157, 170–171, 173 Sato, M., see Tosuji, H. Sawyer, R.T., 25, 80 Sayers, C.W., 50 Sayles, L.P., 103, 113–114 Schiedges, K.L., 159, 162, 168 Schierwater, B., 27, 126, 131–133, 144–145, 148 Schleicherova, D., 45, 54, 88 Schmelz, R.M., 2, 6, 9, 12, 117, 209 Schneider, S., 47 Schottler, U., 25–26 Schroeder, P.C., 33–34, 62–64, 89, 128, 157, 218 Schroeder, P.C., see Heacox, A.E. Seaver, E.C., see Irvine, S.Q. Selander, R.K., see Jaenike, R.J. Sella, G., 44, 54, 61, 64, 72, 75, 88–89 Sella, G., see Premoli, M.C. Senapati, B.K., 20 Serebiah, J.S., 196 Shanks, A.L., 155 Siddique, J., 72, 79, 203–204, 212 Siebers, D., 22 Sikes, J.M., see Bely, A.E. Simonini, R., 91 Simonini, R., see Prevedelli, D. Simpson, I.C., 197 Singhal, R.N., 191 Singhal, R.N., see Davies, R.W. Sket, B., 5, 6 Smirov, R.V., see Buzhinskaja, G.N. Smith, H.L., 46 Smith, C.R., 10 Smith, D.S., 186 Soame, J.M., see Lawrence, A.J. Southward, A.J., 15, 22 Southward, E.C., see Southward, A.J. Springett, J.A., 28, 209, 210 Sreepada, K.S., see Hegde, P.R. Sruthy, P.B., 202 Stephens, G.C., 22 Stephens, G.C., see Reish, D.J. Steven, V.R., see Preston, R.L. Stevens, B.R.., 22 Stevens, R.B., 194 Stolc, A., 126 Stone, R.G., 106–107, 115 Strathmann, M.F., 155 Strecher, H.J., 35 Sturzenbaum, S.R.., 5 Styan, C.A., 174 Subramanian, E.R., 14, 20, 107–108, 110, 202

Author Index  263 

Sudhakar, S., 96–97, 108, 129 Sugio, M., 118 Suomalainen, E., 65–66 Surholt, B., 26 Swalla, B., see Rychel, A.L.

T Tadokoro, R., 107, 148–149 Takeo, M., 110–111, 117 Tenore, K.R., 24 Tessmar-Raible, K., 182 Testerman, J.K., 22 Tettamanti, G., 106 Theede, H., 25 Timm, T., 27–28, 126 Tochinai, S., see Yoshida-Noro, C. Tondoh, E.J., 29, 79 Tosuji, H., 171, 173–174 Tovar-Hernandez, M.A., 54, 128, 130–131, 134 Traut, W., 91 Trontelj, P., see Sket, B. Tsai , C.F., 66, 68 Tsuji, F.I., 120, 159, 165–167 Tunicliffe, V., see McHugh, D. Tzetlin, A.B., 128

U Underwood, A.J., see James, R.J. Urcuyo, I.A., 18 Utevsky, S., 172 Uwe-Dahms, H., see Qian, P.Y.

V Van Cleave, C.D., 113, 118 van der Have, T.M., 4 Van Dover, C.L., 16, 19 Vandini, R.Z., see Prevedelli, D. Velando, A., 87–88 Venter, J.M., see Reinecke, A.J. Verkuijlen, J., see Ratsak, C.H. Viljoen, S.A., 203 Vittor, B.A., 157 Von Haffner, K., 100

W Watson, A.T., 116, 136 Watson, G.J., 44, 181, 183 Wehe, T., 156, 159 Weidhase, M., 98, 105, 112, 179, 186, 188 Weinberg, J.R., 87, 175 Westheide, W., 2, 5, 6, 33–35, 50, 73, 75, 112 Whitford, T.A., 100–101, 103 Williams, G.B., 52 Williams, J.D., 126, 131, 134, 144, 146 Williams, J.D., see David, A.A., Dualan, I.V., Whitford, T.A. Williams, M.E., 43–44 Wilson, W.H., 20, 35–36, 41–42, 131, 133, 134, 218 Wirtz, P. see Kutschera, U. Won, S., 2 Woodin, S.A., 103 Woodin, S.A., see Lindsay, S.M.

X Xiao, N., 65, 92, 96–97, 115, 119

Y Yang, T., 172 Yasmin, S., 190 Yesudhason, B.V., 191 Yokota, H., 197 Yoshida-Noro, C., 110, 135, 209 Young, J.O., 50 Yuan, S.L., 128, 134 Yuwono, E., see Golding, D.W.

Z Zajac, R.N., 58, 92, 104–105 Zattara, E.E., 93, 95, 98, 113, 121–124, 127–129, 141–142, 220–221 Zeeck, E., 168 Zoran, M.J., see Martinez-Acosta, V.G.

Species Index

A Abarenicola, 99 Abramis brama, 12 Abyssidrilus stilus, 10–12, 218 Acanthobdella, 8 Achaeta bilobisa, 66 A. eiseni, 198 Aeolosoma, 3 A. spp, 129 A. beddardi, 129, 142 A. furcatus, 129, 145 A. headleyi, 129, 142 A. hemprichii, 126, 129, 142, 171 A. hyalinum, 143–144 A. kashyapi, 129 A. niveum, 129, 142 A. quaternarum, 56, 129 A. singutare, 129 A. ternarium, 129, 142 A. titorale, 129 A. travancorense, 129, 142 A. viride, 126, 129, 142, 146–147, 150, 151, 145, 198, 221 Allolobophora caliginosa, 21, 115, 119, 171, 204 A. chlorotica, 72, 171 A. iterica, 171 A. molleri, 99 A. muldali, 66–67 A. nocturna, 171 A. terrestris, 171 A. chlorotica, 72, 171 Alkmaria romijni, 62 Alma emini, 26 Allonais inaequalis, 129, 142 A. lairdi, 129 A. paraguayensis, 120, 129, 136, 142 A. pectinata, 129, 143 A. rayalaseemensis, 129, 142 Alvinella ivanovi, 17 A. prompejana, 18–19 A. spiralis, 17 Alvinocaris lusca, 16 Amaena occidentalis, 61 Ampharete, 3

Amphichaeta, 3 A. raptisae, 106, 123, 129 A. sannio, 4, 129 Amphicorina brevicollaris, 60 Amphiglena marita, 60, 62 A. mediterranea, 60, 62 A. nathae, 60, 62 A. terebro, 60, 62 Amphinome, 2 Amphipholis squamata, 49 Amphipolydora abranchiata, 128 A. vestalis, 93, 103, 116, 121, 128, 131–133, 135–137 Amphisamytha galapagensis, 58 Amphitrite, 3, 8, 53 Amynthas bileatus, 66 A. catenus, 66 A. chilanensis, 66, 68 A. corticus, 66 A. cruxus, 66 A. diffringens, 66 A. gracilis, 66 A. hohuanmontis, 66 A. hupiensis, 66 A. shinanmontis, 66, 68 A. sheni, 68 A. tokioensis, 66 Aphelochaeta, 49 Aphelochaeta sp, 62 A. glandularia, 156 A. monilaris, 9, 156 Aphrodita, 1 Apisthobranchus, 11 A. glacierae, 62 Apodotrocha progenerans, 112 Aporrectdodea caliginosa, 66, 72, 199 A. longa, 72, 198–199, 202–204, 214–215, 222 A. rosea, 66, 69, 170, 172 A. trapezoides, 29, 66, 68–69, 72, 172, 200, 202–204 Aracia sinaloae, 54, 56, 62 Arcidea monicae, 11 A. ramosa, 11 A. simplex, 11 A. wassi, 11

Species Index  265 

Arcteonais lomondi, 129 Arenicola, 2, 3, 53–54 A. marina, 24, 26, 43–45, 98–100, 103, 111, 120, 123, 168, 180–181, 183 Aricia, 107 Artemia franciscana, 201 Asteromyzostomum grygieri, 63 Augeneriella alata, 60 Aulodrilus limnobius, 197 Aulophorus carteri, 129, 143, 210 A. flabelliger, 142 A. flagellum, 129 A. furcatus, 129, 143, 197 A. gravelyi, 129, 142 A. gwaliorensis, 142 A. hymanae, 129, 142, 197 A. indicus, 129, 142 A. michaelseni, 129, 142 A. moghei, 129, 142 A. tonkinensis, 29, 142 A. vagus, 197 Autodrilus sp, 129 A. japonicus, 129 A. pluriseta, 129 Autolytus, 144, 153, 161 Autolytus spp, 168, 173 A. alexandri, 157 A. brachycephalus, 159, 162, 173 A. charcoti, 159 A. cornuta, 120 A. edwardsii, 115, 159, 162 A. magnus, 159 A. pictus, 100, 108, 120 A. prolifer, 44, 159–163, 173 A. purpureimaculata, 159

B Bacillus endophyticus, 14, 20, 108, 110, 202 Balkaiodrilus maievici, 11 Bimastos, 35, 216 B. beddardi, 66 B. eiseni, 172, 215–216 B. gieseleri, 66 B. heimburgeri, 66 B. longicinctus, 66 B. palustris, 66 B. parvus, 66 B. tenuis, 172, 215–216 B. tumidus, 66 B. welchi, 66 B. zeteki, 66 Bispira brunnea, 58, 119, 121, 128, 131–134, 136, 151 Boccardia, 195 B. acus, 84 B. andrologyna, 84 B. chilensis, 84

B. proboscidae, 46, 84, 86–87 B. semibranchiata, 84 B. syria, 99 Bollandia anthipathicola, 62 Bothrioneurum righii, 129 B. vedjioskyanum, 129 Branchellion torpedinis, 172 Branchiodrilus hortensis, 129, 142 B. menoni, 129 B. semperi, 129, 144 Branchiomma bairdi, 56, 62, 128 B. cingulata, 62 B. curtum, 128 B. luctuosum, 62, 99 B. nigromaculata, 99, 111, 120 Branchiomaldane vincenti, 62 Brachionus plicatilis, 201 Branchipolynoe seepensis, 58, 73, 78 B. symmytilida, 19 Branchiura sowerbyi, 30, 77, 79, 81, 83, 171, 190–191, 197–198, 205–208, 210–211, 213–214, 223 Brania clavata, 63, 90 B. protandrica, 62 B. pusilla, 66, 120, 124, 159 B. pusilloides, 62 Brassica compensis, 197 Bratislavia unidentata, 129 Brifacia araiargonensis, 194 Buchholzia appendiculata, 12, 129, 218 Bugula neritina, 52 Bushiella sp, 62 B. abnormis, 62 B. atlantica, 62 B. granulata, 62 B. kofiadii, 62 B. quadrangularis, 62 B. similis, 62 Bythograea thermydron, 16

C Califa calida, 11 Calanus finmarchicus, 201 Caobangia abbotti, 60 C. billeti, 62 Capitella, 41 Capitella sp, 52 Capitella sp I, 48, 58, 73, 90, 99, 120 Capitella sp II, 120 C. capitata, 22, 25, 29, 58, 155–156, 162–163, 171, 173, 176–177, 222 C. hermaphrodita, 62 Capitomastus minimus, 62 Caulleriella lafolla, 7 C. parva, 62 C. viridis, 128, 156 Chaetogaster, 117

266  Reproduction and Development in Annelida

C. cristallinus, 129, 143 C. diaphanus, 106, 123, 129, 143–144, 221 C. diastrophus, 28, 106, 123, 129, 143, 221 C. langi, 129, 143, 210–211 C. limnaei, 129 C. limnae bengalensis, 129, 142 C. limnae limnae, 142 Chaetopterus, 3, 46, 107 C. variopedatus, 34, 93, 99, 106, 111, 113–114, 116, 121, 219 Chaetozone acuta, 9 C. corona, 9 C. gracilis, 10–11 C. setosa, 9 C. spinosa, 11 C. vivipara, 62 Ciona sp, 14 Circeis sp, 62 C. armoricana, 62 C. oshurkovi, 62 C. paguri, 62 C. spirillum, 171 Cirratulus cf cirrus, 112 C. cirratus, 99, 128, 156, 180 C. hedgepethi, 22 C. incertus, 156 Cirriformia spirobranchia, 22 Clymenella torquata, 22, 99, 103, 113–114, 120–121 Cognettia cognettia, 198 C. glandulosa, 129 C. sphagnetorum, 126, 129, 209 Contramyzostoma, 63 Cossura brunnea, 11 C. candida, 11, 13 C. modica, 11 C. pygodactylata, 11 C. rostrata, 11 Criodrilus lacuum, 27–28, 120 Cruistipellis tribranchiata, 129 Ctenodrilus serratus, 62, 128, 156 Cyclocirra, 63 Cystimyzostomum, 63

D Dasychone infrata, 111 Demonax medius, 60 D. pallidus, 62 Dendrobaena, 216 D. mammalis, 172 D. octaedra, 66, 172 D. rubidus, 66–67, 172, 215–216 D. subrucunda, 172, 215–216 Dendrodrilus rubidus, 66, 69, 172, 211 Dendronereis aestuarina, 155, 157–158, 165–167 Dero, 99 Dero sp I, 129

D. bauchiensis, 129 D. borelii, 129 D. carteri, 129 D. cooperi, 142 D. digitata, 120, 129, 145–146, 197, 200 D. dorsalis, 210–211 D. evelinae, 210 D. flabelliger, 129 D. furcata, 129 D. gravelyi, 129 D. huaronensis, 129 D. indica, 142, 171 D. limosa, 118 D. lutzi, 129 D. malayana, 129 D. multibranchiata, 210 D. nivea, 142 D. palmata, 142 D. pectinata, 142 D. plumosa, 142 D. ravinensis, 142 D. sawayai, 142 D. superterrenus, 129 D. tonkinensis, 129 D. vaga, 129 D. zeylandica, 142 Dichogaster bolaui, 66 Dina lineata, 172 D. modigilianii, 215 Dinophilus gardnieri, 120, 123 D. gyrociliatus, 28–29, 39, 58, 70, 77, 79, 81–82, 90, 171, 173, 175, 222 D. rostratus, 128 Diopatra, 107 Diopatra spp, 120 D. aciculata, 92 D. amboinensis, 99, 113 D. bilobata, 4 D. cuprea, 22 D. dexiognatha, 99 D. marocensis, 54, 56, 62 D. neapolitana, 99, 101, 191–192 Diplocardia singularis, 67 Diploria strigosa, 52 Dipolydora armata, 128, 135–136 D. caulleryi, 128, 131 D. commensalis, 13, 63, 92, 99 D. socialis, 128, 131 D. quadrilobata, 92, 99, 120 Ditrupa arietina, 171 Diurodrilus sp, 76 Dodecaceria berkeleyi, 128 D. caulleryi, 156 D. concharum, 66, 114, 115, 121, 128, 135–136, 150, 151, 156 D. coralii, 128

Species Index  267 

D. fewkesi, 128 D. fimbriata, 128, 156 D. fistulicola, 128 D. pulchra, 66, 121, 128, 131–136 Dorvillea bermudensis, 99, 120 Drawida hattamimiju, 66 D. nepalensis, 66, 72, 215 Drieschia pelagica, 156

E Eirene viridis, 162 Eisenia, 216 E. andrei, 45, 87–88 E. foetida, 34, 43–45, 57, 67, 72, 79, 87, 92, 96– 97, 99–102, 104, 106, 113, 115, 118–120, 172, 190–191, 202–204, 212–214 E. rosea, 66, 215–216 E. rosea f.mut, 172 E. rosea f.typica, 172 E. venata, 172, 215–216 Eiseniella tetraedra, 66, 172 Enapleris euchaeta, 62 Enchytraeus, 138 E. albidus, 22, 139, 197, 201–202, 209–214, 223 E. bigeminus, 129, 132–133, 136, 138–139, 208–209 E. buchholzi, 117, 12, 197, 210, 221 E. coronatus, 197, 209–211 E. crypticus, 190, 209–210 E. dudichi, 120, 129 E. fragmentosus, 120, 129 E. higentius, 126 E. irregularis, 139 E. japonensis, 106–107, 110, 115–118, 120–121, 129, 132–133, 135–136, 145, 148–151, 200, 209, 221 E. variatus, 29–30, 80–81, 83, 104, 129, 133, 136, 138–139, 144 Endomyzostoma scotia, 63 E. neridae, 63 Enteromorpha sp, 205 Eophila, 99 Ephesiella mizla, 62 Erpobdella octoculata, 51, 172 E. testacea, 172 Escarpia laminata, 17 E. southwarde, 17 E. spicata, 17 Euclymene oerstedi, 98–99, 103, 119–120, 134 Eudrilus eugeniae, 14, 20, 72, 97, 108–109, 112–113, 134, 191, 200–204, 212, 214, 222 Eulaeospira convexis, 62 Eulalia spp, 99 E. viridis, 120, 181, 183 Eukerria saltensis, 66 Eunereis longissima, 22 Eunice sp, 7

E. afra, 120, 156 E. fucata, 120, 156 E. schemacephala, 156, 163 E. schizobranchia, 120, 156 E. siciliensis, 112, 120, 155–156, 158, 163–164, 166–168 E. torquata, 120, 156 E. viridis, 3, 42, 120, 156, 164, 166 Eupolymnia nebulosa, 59–60 E. crescentis, 60, 75–76, 78 Euratella salmacidis, 62 Eurythoe complanata, 34, 99–100, 103, 111, 120–121, 128, 134, 136, 156 E. oerstedi, 134 E. parvecarunculata, 156 Eusyllis blomstrandi, 120, 159, 162–163 Eutyphoeus gammiei, 214–215 Euzonus flabelligerus, 157 E. mucronata, 4, 26 Exogone gemmifera, 63, 65 E. hebes, 159 E. meridionalis, 194 E. naidina, 62, 120, 159, 163, 194 E. rubescens, 161 E. verugera, 63

F Fabricia, 34 F. stellaris, 60 Fabricinuda trilobata, 60 Ficopomatus enigmaticus, 58–59, 63, 171 Filograna/Salamacina complex, 62 Filogranella elatensis, 128 F. gracilis, 128 F. implexa, 128, 171, 173 Flabelligela macrochaeta, 156 Frederiova bubosa, 171 Fridericia spp, 197 F. bisetosa, 66 F. connata, 66 F. galba, 67 F. ratzeli, 66 F. striata, 66 Fucus serratus, 52

G Galeolaria caespitosa, 34, 42–43, 45, 63, 74, 174–175, 177, 222 G. hystrix, 63 Glossiphonia complanata, 21, 51, 55–56, 172 G. complanata concolor, 172 G. heteroclita papillosa, 172 G. heteroclita, 172 Glossodrilus, 12 Glycera, 2 G. alba, 156

268  Reproduction and Development in Annelida

G. americana, 22, 156, 163 G. capitata, 156, 163 G. dibranchiata, 22, 58, 75–76, 78, 82, 153–154, 156–158, 164, 166, 205–206 G. gigantea, 156 G. macrobranchia, 156, 163 G. oxycephala, 156, 162–163 G. tenuis, 156 Goniada sp, 22 Grubeosyllis clavata, 120, 159, 163 G.neapolitana, 62

H Haementeria ghilianii, 25, 80, 82 H. parasitica, 44 Haemodipsa, 21 Haemonais waldvozeli, 143 Haemopis, 2 H. sanguisuga, 172 Halla parthenopeia, 49, 58 Halopteris scoporia, 59, 188 Haplosyllides floridana, 120, 159 Haplosyllis spongicola, 13–14, 62, 114–115, 120, 159, 163 Haplotaxis sp, 11 Harmothoe imbricata, 26, 74, 82, 120, 180 Hediste spp, 157, 173–174, 222 H. atoka, 171, 173 H. diadroma, 155, 157, 171, 173–175 H. japonica, 58, 70, 80, 82, 84, 155, 157, 171, 173–175 H. osawai, 157 H. oxypoda sensu, 157 Helicosiphon platyspira, 62 H. biscoensis, 62 Heliodrilus hachiojii, 66 Helobdella californica, 82 H. robusta, 149 H. stagnalis, 50–51 H. striata, 63 Hemiclepsis marginata, 172 Henlea ventriclosus, 171 Hesione sicula, 62 H. mazima (?), 62 H. pantherina, 62 Hesionides, 7 H. arenaria, 76 Hirudo, 2, 21 H. medicinalis, 4, 20, 29, 55–56, 70, 77, 79, 80, 82, 99, 106, 172 Hirudinaria manillensis, 191 H. nipponia, 172 H. orientalis, 172 H. vertana, 172 Homogaster elisae, 87 Hrabeiella periglandulata, 9 Hydroides, 2, 3

H. dianthus, 51, 99, 120 H. elegans, 52, 63, 65, 89, 171 H. ezoensis, 51 H. norvegica, 63, 171 Hyperiodrilus africanus, 29, 79, 202–204 Hypomyzostoma jasoni, 63 H. jonathoni, 63

I Idanthyrsuspennatus, 34 Ikosipodus carolensis, 76 Ilyodrilus templetoni, 28 Inanidrilus, 14, 16 I. leukodermatus, 14–16

J Janua brasiliensis, 52 J. pagenstecheri, 62, 171 Janus knightjohnsi, 63 Johanssonia, 21 J. arctica, 29 Josephella sp, 128 J. marenzelleri, 128

K Kirkegaardia, 99, 128 Kefersteinia cirrata, 31, 155

L Lablab purpureus, 197 Laetmonica product, 13 Lagis koreni, 62 Lamellibrachia spp, 18 L. anaximandri, 17 L. barhami, 17–19 L. columna, 17 L. juni, 17 L. luymesi, 17–19, 43 L. sagami, 17 L. satsuma, 17, 95–96, 114, 121 Lampito mauritii, 26, 215 Lanice conchilega, 22, 47, 75 Laonome albicingillum, 43, 45, 54, 56, 62, 64–65 Limnatus, 21 L. nilotica, 21 Limnodrilus, 66 Limnodrilus sp, 205 L. claparedianus, 120, 123, 171 L. hoffmeisteri, 24, 28, 30, 66, 99, 106–107, 171, 197–198, 216–217 L. mastix, 66 L. undekemianus, 171 Linopherus canariensis, 128 Lithothammoni, 52

Species Index  269 

Lumbricillus lineatus, 66, 69, 120, 123, 171, 209–210, 211–214 L. rivalis, 30, 197–198, 209–211, 213–214 Lumbriculus, 2 Lumbriculus sp, 205, 223 L. lineatus, 210 L. variegatus, 66, 92–93, 98–100, 103, 106–107, 115–118, 121–122, 129, 132, 134–136, 146, 150–151, 219, 221 Lumbriculidae spp, 197 Lumbricus, 8, 32 L. castaneus, 172 L. eiseni, 66 L. festiwas, 172 L. friendi, 172 L. rubellus, 172 L. terrestris, 35, 45, 66, 72, 79, 81, 83, 96–97, 102, 104, 113, 120, 172, 191, 199, 202, 203, 204, 211–215, 219 Lumbrinereis spp, 22 L. funchalensis, 58–59 L. latreilli, 48 Lysidice sp, 120 L. collaris, 124 L. ninetta, 124

M Macellicephala violacea, 62 Macrobrachium idella, 202 Maldane, 2 Manayunkia aestuarina, 4, 60, 62 M. caspica, 60 M. speciosa, 60 Marionina southerni, 198, 208–209 Marenzellaria viridis, 92, 99–101, 103 Marionina sp, 129 M. clevata, 28 Marphysa disjuncta, 157 M. sanguinea, 22, 39, 58, 71, 76, 78, 80, 82, 197–198, 205–206, 211 Mediomastus sp, 120 Megalomma cinctum, 128, 134 Megascolides australis, 179 Mercierella enigmatica, 26, 59–60, 63 Mesenchytraeus sp, 197 M. glandulosus, 66 Mesomyzostoma botulus, 63 M. katoi, 63 M. lanterbecqae, 63 M. leukos, 63 M. lobus, 63 M. okadai, 63 M. reichenspergeri, 63 Mesonerilla armoricana, 62 M. fagei, 62 M. roscovita, 62

Metalaeospira clasmani, 62 M. pixelli, 62 M. tenuis, 62 Metaphire hilgendorfi, 66 M. houlleti, 27, 203, 212, 214–215 M. peguana, 120 Micromaldane, 34 Microphthalmus, 34, 76 M. aberrans, 35 M. fragilis, 62 M. listensis, 62 M. oberrans, 62 M. sczelkowii, 62, 76 M. similis, 62 M. tyrrhenicus, 62 M. urofimbriatus, 62 Microscolex dubius, 66 M. phosphoreus, 66 Milliporea complanata, 21 Millsonia anomala, 21 Monticellina heterochaeta, 156, 163–164 M. serratiseta, 7 Munidopsis lentigo, 16 M. subsquamosa, 16 Myrianida, 145, 161 M. pachycera, 120, 160–161 M. pinnigera, 62, 159 Myriochele heeri, 127–128, 131–134, 136 Mystides caeca, 157 Mytilus edulis, 24, 195 Myxicola aesthetica, 99, 120, 124, 128 M. cf sulcata, 74 Myxobolus cerebralis, 194 Myzostoma cirriferum, 63 M. debiae, 63 M. deformator, 63 M. eeckhauti, 63 M. fuscomaculatum, 63 M. glabrum, 63 M. hollandi, 63 M. indocuniculus, 63 M. josefinae, 63 M. kymae, 63 M. laurenae, 63 M. miki, 63 M. pipkini, 63 M. pulvinar, 63 M. susanae, 63 M. tertiusi, 63

N Nainereis dendritica, 7, 22 N. laevigata, 62 Nais sp, 129 N. abissalis, 10–11 N. andhrensis, 129, 142 N. andina, 129, 143

270  Reproduction and Development in Annelida

N. barbata, 129, 143–144, 148 N. bretscheri, 129, 143 N. commensalis, 118 N. communis, 121, 126, 129, 132, 143–145 N. elinguis, 22, 120, 129, 143, 210–211 N. paraguayensis, 118, 139 N. pardalis, 129, 142 N. pseudobtusa, 129, 143, 210 N. simplex, 129, 143 N. stolci, 129 N. variabilis, 129, 142, 210 Namalycastis, 2 N. indica, 62, 65 Namanereis quadraticeps, 62 Neanthes arenaceodentata, 22, 49, 190, 192, 205–207, 211–212 N. japonica, 170–171, 173 N. lighti, 62 N. limnicola, 22, 29, 54, 56, 62, 70, 77–78, 80 Nematonereis unicornis, 120, 124 Neoamphitrite figulus, 25 N. robusta, 60–61, 75, 82 Neodexiospira alveolata, 62 N. brasiliensis, 62 N. foraminosa, 62 N. formosa, 62 N. kayi, 62 N. lamellosa, 62 N. pseudocorrugata, 62 N. steueri, 62 Neolepoa septochaeta, 44 Nephelopsis obscura, 71, 77, 79, 82, 172 Nephtys, 22, 111 Nephtys sp, 107 N. caeca, 60, 99, 157, 182 N. cirrosa, 182–183 N. hombergii, 60, 127, 181, 183 Nereiphylla castanea, 157, 163 Nereis, 3, 181 Nereis sp, 107 Nereis spp, 25 N. acuminata, 47, 87, 171, 173, 175 N. brevicornis, 158 N. diadroma, 158 N. diversicolor, 22, 24, 100–101, 111, 158, 171, 180–182 N. falcaria, 48, 157, 163, 166 N. fucata, 157–158, 163, 166 N. grubei, 157 N. japonica, 157–158 N. latescens, 111 N. limbata, 155, 157–158 N. limnicola, 22 N. pelagica, 25, 153, 157, 180 N. succinea, 22–23, 157, 168, 181–182 N. virens, 1, 4, 9, 22, 24, 32, 43, 58, 71, 75, 80, 82, 99, 113, 148, 153, 157, 166, 181–182, 201–202, 205–206, 211, 212, 222

Nerilla, 2 Nerillidium gracile, 63 N. macropharyngeum, 63 N. mediterraneum, 63 N. renaudae, 63 N. simplex, 63 N. troglochaetoides, 63 Nerillidopsis hyaline, 63 Nicolea upsiana, 58 N. zostericola, 44, 58, 75, 77–78, 82 Nidificarisa nidica, 62 N. palliata, 62 Notopharyngoides, 63 Notophyllum, 1

O Ocidus alvinae, 17 O. fujikuwari, 17 Ocnerodrilus, 26 O. occidentalis, 66 Octolasion, 216 O. cyaneum, 66, 68, 172, 202–204, 214–216 O. occidentalis, 66 O. lacteum, 172, 216 O. tyrtaeum, 66, 69, 172 Odontosyllis, 153, 157 O. ctenostoma, 128 O. enopla, 120, 153, 155, 157–159, 163, 166–167 O. gibba, 128 O. hyalina, 159, 166 O. luminosa, 153, 159, 163, 165–168 O. phosphorea, 120, 159, 163, 165–167 O. polycera, 120, 153, 159, 163, 166 O. prolifera, 120 O. undecimdonta, 159, 166 Olavius, 14, 16 O. algarvensis, 14, 16 O. ilvae, 14 Onuphis, 2 Onychochaeta windlei, 66 Ophidonais serpentina, 107, 129 Ophiodromus flexuosus, 155 Ophryotrocha, 27, 33–34, 71–72, 87–89 Ophryotrocha spp, 56, 61, 63, 71, 75, 181, 184 O. adherens, 29, 49, 77, 79, 81 O. bacci, 62 O. diadema, 29, 43, 45, 54, 62, 64, 72, 87–89, 177 O. gracilis, 54, 61–62, 64, 71–72, 76, 88–89 O. hartmanni, 62, 72, 89 O. labronica, 39, 57, 62, 72, 88–89, 177–178, 184–186, 191, 193 O. macrovifera, 71–72 O. maculata, 62, 71–72 O. notoglandulata, 72, 100, 120 O. puerilis puerilis, 49, 63, 80, 82, 87–89, 120, 184, 186, 219 O. robusta, 71–72

Species Index  271 

O. socialis, 29, 49, 62, 72 O. vivipara, 76 Oreochromis niloticus, 202 Oriopsis, 34 Oryzias melanostigma, 49 Osedax, 17 Owenia, 108 O. fusiformis, 99, 120 Oxydromus propinquius, 155

P Paradexiospira vitrea, 62–63 Paraescarpia echinospica, 17, 19 Parafabricia mazzellae, 194 Paralaeospira levinseni, 62 P. malardi, 62 P. parallela, 62 Paranais frici, 106, 123, 129 P. leidyi, 99, 148 P. litoralis, 4, 93, 98–99, 106, 121, 123, 129, 138 P. paraguayensis, 139 Paranaitis polynoides, 157, 163 Parenterodrilus taeniodes, 57 Pareurythoe californica, 22 Parougia albomaculatus, 126, 128 P. bermudensis, 107, 126, 128, 130–131, 136, 138, 150–151, 221 Pectinaria, 2 P. gouldii, 24 Pedonais crassifaucis, 129 Penaeus monodon, 201 Pentoscolex corethrurus, 215 Peregodrilus heideri, 9 Perinereis spp, 4, 199, 205 P. cultrifera, 39, 58, 75, 82, 153, 157, 181–182 P. dumerilii, 153, 180–182 P. helleri, 206 P. nuntia, 99, 113, 157, 205–206 P. nuntia brevicirrus, 157 P. rullieri, 39, 58, 205–206 P. vancaurica, 182 P. vancaurica tetradentata, 155, 157–158 Perionyx ceylanensis, 198–199, 202–204, 214 P. excavatus, 27, 96–98, 103, 113, 115, 117–118, 120–121, 123, 129, 139, 215 P. polytheca, 35 Perkinsiana spp, 60, 62, 1 P. antarctica, 60, 62 P. milae, 126, 128 P. rubra, 128 Petaloproctus socialis, 120 Phallodrilus, 14 Pheretima, 2, 66, 102, 104 Pheretima sp, 120 Pheretima (indica?), 102, 104 P. agrestis, 66 P. bicinta, 66

P. diffringens, 66 P. guillelmii, 190–191 P. hilgendorfi, 66 P. hupiensis, 66 P. levis, 66 P. loveridgei, 66 Phragmatopoma californica, 51–52 Phreodrilus, 32 Phyllochaetopterus limicolus, 11 P. prolifica, 128 P. socialis, 128 Phyllodes groenlandica, 163 Phyllodoce, 7 P. cuspidata, 157 P. groenlandica, 157 P. hartmanae, 157 P. longipes, 157, 163 Phylo nudus, 7, 10–11 Piguetiella michiganensis, 129 Pileolaris spp, 193–194 Pileolaria sp1, 62 P. sp2, 62 P. berkeleyana, 62 P. daijonesi, 62 P. darkarensis, 62 P. lateralis, 62 P. marginata, 62 P. militaris, 62 P. pseudoclavus, 62 P. spinifer, 62 P. tiarata, 62 Pillaiospira trifurcata, 62 Pionosyllis, 145, 161 P. lamelligera, 120, 159 P. neapolitana, 62, 159 P. procera, 120, 159 P. pulligera, 120, 159, 166-167 Piscicola, 3 Pisione alikunhi, 35 P. remota, 35, 76 Piutellus papillifer, 66 P. umbellulariae, 66 Placobdella, 2 P. papillifera, 172 Placostegus tridentatus, 171 Platynereis, 181 P. dumerilii, 47–48, 99, 111, 120, 148, 157, 163, 168, 193–194 P. massiliensis, 47, 49, 62, 194 Podarke pallida, 155 P. pugettensis, 22 Poecilochaetus, 47 P. johnsoni, 11 Polycera cornuta, 160–161 Polydora, 195 P. caulleryi, 99–100 P. ciliata, 99, 195 P. colonia, 128, 134–136, 151

272  Reproduction and Development in Annelida

P. commensalis, 62 P. cornuta, 46, 92, 99, 103–104 P. curiosa, 171, 173 P. elegantissima, 128 P. flava, 99 P. gigardi, 62 P. hermaphroditica, 62 P. ligni, 35, 41, 52, 58–59, 62, 92, 99, 104–105 P. websteri, 195 Polydorella spp, 103 P. dawydoffi, 128, 145, 150–151 P. kamakamai, 126, 128, 131, 134, 144–146 P. prolifera, 128 P. schizogenica, 130 P. smurovi, 128 P. stolinifera, 128, 134 P. tetrabranchia, 128, 145 Polygordius, 3 P. triestinus, 62 Polyophthalmus pictus, 120, 123 Polynoe, 1 Polypheretima elongata, 215 Pomatoceros, 57 Pomatoceros lamarckii, 62–63 P. triqueter, 63, 171 Pomatoleios kraussi, 58 Pontobdella, 2 P. muricata, 172 Pontoscolex corethrurus, 66, 72, 215 Potamilla sp, 60 P. antarctica, 83 P. torelli, 60, 116, 128, 136, 151, 199 Potamothrix bedoti, 126 P. hammoniensis, 12, 28 P. moldaviensis, 27–28 P. vejdoskyyaneum, 126 Pristina, 146 Pristina sp, 197 P. aequiseta, 129, 143, 170–171, 210 P. breviseta, 129, 143 P. evelinae, 129, 143 P. foreli, 143 P. leidyi, 99, 129, 220 P. longiseta, 118, 129, 132, 145, 210 P. longiseta longiseta, 143 P. macrochaeta, 129, 143 P. proboscidea, 129, 143 P. sperberae, 129, 143 P. synchites, 129, 143 Pristinella acuminata, 129, 143 P. jenkinae, 129, 143 P. menoni, 129, 143 P. minuta, 129, 143 Proceraea cornuta, 159 P. okadai, 159 P. picta, 159-160 Procerastea, 145, 161

P. halleziana, 114–115, 120–121, 128, 150–151, 160 Propappus glandulosus, 11 Proscoloplos cygnochaetus, 124, 128 Proteobacteria, 14 Protocirrineris antarctica, 128 P. chrysoderma, 128 Protodrilus albicans, 73, 76 Protolaeospira eximia, 62 P. pedalis, 62 P. striata, 62 P. translucens, 62–63 P. tricostalis, 62 P. triflabellis, 62 P. stalagmia, 62 Protomystides confusa, 157 Protomyzostomum lingua, 63 P. roseus, 63 Psammoryctides barbatus, 28 Pseudobranchiomma emersoni, 128 P. minima, 128 P. perkinsi, 128 P. punctata, 128 P. schizogenica, 128, 130–131, 134, 136, 151 Pseudochaetopterus prolifica, 136 Pseudopolydora kempi japonica, 20, 92, 99, 103, 205 P. paucibranchiata, 41 Pseudopotamilla reniformis, 128, 130, 132–137, 151 Pterocirrus foliosus, 157 Pulvinomyzostomum inaki, 63 P. messingi, 63 Pygospio californica, 128, 131 P. elegans, 20, 84, 98–101, 103, 116, 120–121, 126, 128, 131–136, 150–151, 221

R Ramex californiensis, 60–61, 77–78, 80 Raphidrilus nemasoma, 62, 128 Raricirrus arcticus, 128 R. beryli, 128 R. maculatus, 128 R. variabilis, 11, 156, 163 R. variegatus, 153 Rhodopsis pusilla, 128 R. simplex, 128 Rhyacodrilus, 12 R. hiemalis, 20, 96 Rhynchelmis limosella, 28 R. vagensis, 120 Rhynchospio glutaeus, 20, 98–99, 103 Ridgeia piscesae, 17–19, 33 Riftia, 3 R. pachyptila, 17–19, 33, 43 Ripistes parasita, 129 Romanchella pustulata, 62

Species Index  273 

R. quadricostalis, 62 R. scoresbyi, 62 R. solea, 62 Rubifabriciola tonerella, 194

S Sabella, 2 Sabella spp, 120 S. alveolata, 51 S. discifera, 128 S. melanostigma, 99, 107 S. microphthalma, 63 S. pavonina, 99, 114, 116, 121, 128, 136, 150–151, 199 S. spallanzanii, 49, 58, 196 S. spectabilis, 49 Sabellastarte sp, 128, 135–136, 151, 199 S. magnifica, 63 S. spectabilis, 63–65 Saccocirrus eroticus, 44 Sagitella kowalewskii, 62 Salix gigantea, 196 Salmacina, 140 S. amphidentata, 128, 131 S. australis, 63, 128, 131 S. dysteri, 128, 131, 140, 150–151 S. incrustans, 63, 128 Scolelepis squamata, 99 Scoloplos armiger, 124 Seephiophila jonesi, 17 Serpula polycerus, 62 S. vermicularis, 170–171 Siboglinum ekmani, 33 Simplaria postwaldi, 62 S. pseudomilitaris, 62 Slavina appendiculata, 129, 143 S. evelinae, 129 S. sawayai, 129 Spargnophilus, 32 Spartina alterniflora, 24–25 Specaria josinae, 129 Sphaerosyllis corruscans, 62 S. hermaphrodita, 62 S. hystrix, 159 Spiochaetopterus costarum costarum, 128 S. oculatus, 99 S. solitaries, 128 Spio benedicti, 99 S. decoratus, 194 S. filicornis, 33, 62 S. setosa, 99–100 Spiophanes anoculata, 11 S. kroeyeri, 11 Spiraserpula snelli, 128 Spirobranchus cariniferus, 62 S. polycerus, 51, 62, 174 S. giganteus, 52

Spirochaeta, 14 Spirorbis bifurcanis, 62, 171 S. borealis, 52 S. corallinae, 62, 171 S. cuneatus, 62 S. gesae, 62 S. incrustans, 63, 128 S. infundibulum, 62 S. inornatus, 62 S. rothilisbergi, 62 S. rupestris, 52, 62 S. spatulatus, 62 S. spirorbis, 53–54, 62, 70, 73–74, 76–77, 80–81, 83, 171 S. strigatus, 62 S. tridentatus, 62, 171 Spirosperma ferox, 27–28 Stauronereis rudolphi, 22 Stephenosoniana sp, 129 S. trivandrana, 129, 143 Stratiodrilus novaehollandiae, 44 Streblospio spp, 73 S. benedicti, 29, 41, 46, 48, 70, 78, 81, 83–85, 106, 120, 124, 127, 219 S. shrubsolii, 78 Streptosyllis verrilli, 159–160 S. websteri, 120, 159 Stylaria, 146 Stylaria spp, 120 S. fossularis, 115, 118, 121–122, 129, 143 S. lacustris, 27, 102, 104, 126, 129, 131–133, 144–145, 148, 150–151, 221 Stylodrilus asiaticus, 11 S. heringianus, 27–28 Syllides japonica, 159 Syllis spp, 120 S. amica, 61, 63, 89, 159, 165–166 S. gracilis, 114–115, 128, 159, 163 S. prolifera, 63, 65, 181, 186–187, 194 S. ramosa, 159–161, 163 S. variegata, 63 S. vittata, 159, 162, 164

T Terebella, 2 Tevnia jerichonana, 17, 19 Tharyx kirkegaardi, 11 T. perbranchiata, 156 Thelepus cincinnatus, 73 T. crispus, 60–61, 75, 77–78, 80 Theromyzon rude, 172 Timarete filigera, 128 T. punctata, 128 T. tessulatum, 50, 172 Tomopteris spp, 1 T. kils, 3 Trichodrilus, 12

274  Reproduction and Development in Annelida

Trilobodrilus, 62 Trocheta bykowskii, 172 T. subviridis, 172 Troglochaetus beranecki, 63 Tubifex, 3–4, 32 T. benedii, 4 T. blanchardi, 171 T. costatus, 30, 171 T. rivulorum, 111, 123 T. tubifex, 8, 12, 27–28, 30, 66–67, 77, 79, 80–81, 83, 99, 102, 104, 107, 112–113, 115, 118, 120, 123, 171, 190–191, 194, 196–198, 200, 205, 208, 210–211, 216–217, 219, 223 Trypanosyllis, 145, 161 T. asterobia, 120, 159–161 T. coeliaca, 159 163 T. crosslandi, 159 T. gemmipara, 159–160 T. ingens, 159 T. zebra, 63, 89, 114–115, 159, 163 Tylorhynchus heterochaetus, 157

Typosyllis antoni, 121, 186, 188 T. hyalina, 159–160 T. prolifera, 59, 63, 90, 120, 159–160, 165–166, 188–189 T. pulchra, 101–102, 120, 159, 188 T. variegata, 62, 159–160, 163 T. vittala, 62

U Uncinais uncinata, 129, 200

V Veneriserva pygoclava meridionalis, 13, 73 Vejdovskyella sp, 129 Vinearia zibrowii, 62

Z Zeppelina monostyla, 126, 128

Subject Index

A Acrosome, 32–34, 48, 175, 191, 222 Adelpophagy, 84, 86–87 Aestivation, 20, 96 Amputation, 92, 94–97, 100–108, 110, 113, 117–118, 121–122, 151, 169, 185 Analysis Allozyme, 67 Electrophoretic, 67, 84

B Bathymetric distribution, 10, 12, 17 Bindin, 15, 48, 175, 222 BrdU labeling retention assay, 108 BrdU pulse chasing assay, 107

C Calvin-Bensen cycle, 15 Catastrophic metamorphism, 47 Chemical cue, 44 Chloragogue, 71, 96, 102, 104, 113, 121, 138, 219 Cloning Unidirectional, 20, 94–96, 199–200 Bidirectional, 1, 96, 98, 132, 150, 199 Multidirectional, 1, 200 Cyclotrophism, 14

D Denitrification, 222 Diapause, 96, 133, 144 Doubling time, 209–210

E Eleocytes, 32, 102, 104, 106, 180–182 Emergency adaptation, 90 Endemism, 12

Endomitosis, 67 Epimorphosis, 92

F Fertilization Cross, 45, 57, 174 Self, 45, 54, 87

G Germline markers, 148 Gut load, 23

H Hibernation, 20, 96, 104 Hirudin, 4 Hirudotherapy, 4 Hybridization, 173, 177 Hypodermic injection, 44, 76

I Inbreeding depression, 57 Interstitial fauna, 15, 75, 218 Iteroparity, 74, 154, 158, 180, 182

J Jelly coat, 48 Juvenile hormone, 31, 52, 101, 102, 162, 180–182, 184

K Krebs cycle, 26

M Michael-Menten kinetics, 21 Microsatellite markers, 87 Moisture potential, 12

276  Reproduction and Development in Annelida

Morphallaxis, 92, 94, 116–117, 146 Muller’s ratchet, 125

N Neoblasts, 96, 107, 110, 113, 115–119, 121–123, 125, 149–150, 152, 221 Nereidin, 162, 180 Nitrification, 197, 222 Nuptial dance, 44, 167–168, 173

O Ocean acidification, 190–191, 193–194 Osmotrophism, 1, 14, 20–22, 218 Oxygen demand, 25

P Pheromones, 90, 153, 167–168, 221 Photoperiod, 70, 113, 131–132, 135, 165–166, 180, 182, 221 Polygenic genetic system, 57 Polyspermy, 42 Pre-meiotic doubling, 67 Primordial Germ Cells, 56, 148, 184, 220 Pseudocolony, 140–141 Pseudocopulation, 34, 44, 76

R Receptor-mediated endocytosis, 32 Redox discontinuity, 15

S Schizoparity, 135–137 Self-assemblage, 112, 200 Seminal segments, 114–116, 119, 121, 150, 219, 221 Semelparity, 74, 76, 144, 154, 158, 180 Sex allocation theory, 87, 90 Sperm Ect aquasperm, 34 Ent aquasperm, 34 Eusperm, 33 Introsperm, 34 Parasperm, 33 Spermatophore, 33–35, 43–44, 67 Spermatozeugmata, 33 Stigobiont, 12, 218 Stolonization, 101–102, 144, 160, 162, 165, 168, 181, 186–189, 194, 222

T Transplantation, 184, 186–187, 192–193 Troglomorphic features, 12 Trophosome, 14, 18, 95

V Vestimentiferan, 1, 13–19, 33, 95

W Wet funnel method, 200

Author’s Biography

Recipient of the S.S. Bhatnagar Prize, the highest Indian award for scientists, one of the ten National Professorships, T.J. Pandian has served as editor/ member of editorial boards of many international journals. His books on Animal Energetics (Academic Press) identify him as a prolific but precise writer. His five volumes on Sexuality, Sex Determination and Differentiation in Fishes, published by CRC Press, are ranked with five stars. He is presently authoring a multi-volume series on Reproduction and Development of Aquatic Invertebrates, of which the volumes on Crustacea, Mollusca, and Echinodermata and Prochordata, have already been published. The present book is on Annelida. The next one, Platyhelminthes is being prepared.