Reproductive Biology and Phylogeny of Birds: Phylogeny, Morphology, Hormones, Fertilization (Reproductive Biology and Phylogeny, Vol 6A)

Reproductive Biology and Phylogeny of Birds Phylogeny Hormones

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Morphology Fertilization

Reproductive Biology and Phylogeny Series Series Editor: Barrie G. M. Jamieson

Published: Vol. 1: Reproductive Biology and Phylogeny of Urodela (Volume Editor: David M. Sever) Vol. 2: Reproductive Biology and Phylogeny of Anura (Volume Editor: Barrie G. M. Jamieson) Vol. 3: Reproductive Biology and Phylogeny of Chondrichthyes (Volume Editor: William C. Hamlett) Vol. 4: Reproductive Biology and Phylogeny of Annelida (Volume Editor: G. Rouse and F. Pleijel) Vol. 5: Reproductive Biology and Phylogeny of Gymnophiona (Caecilians) (Volume Editor: Jean-Marie Exbrayat) Vol. 6A: Reproductive Biology and Phylogeny of Birds (Volume Editor: Barrie G. M. Jamieson)

In press/under preparation: Vol. 6B: Reproductive Biology and Phylogeny of Birds (Volume Editor: Barrie G. M. Jamieson) Vol. 7: Reproductive Biology and Phylogeny of Cetacea (Volume Editor: D. Miller)

Reproductive Biology and Phylogeny of Birds Part A

Phylogeny Hormones

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Morphology Fertilization

Volume edited by BARRIE G.M. JAMIESON School of Integrative Biology University of Queensland St. Lucia, Queensland Australia

Volume 6A of Series: Reproductive Biology and Phylogeny Series edited by BARRIE G.M. JAMIESON School of Integrative Biology University of Queensland THE UNIVERSITY St. Lucia, Queensland OF QUEENSLAND AUSTRALIA Australia

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All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying or otherwise, without the prior permission. This book is sold subject to the condition that it shall not, by way of trade or otherwise be lent, re-sold, hired out, or otherwise circulated without the publisher’s prior consent in any form of binding or cover other than that in which it is published and without a similar condition including this condition being imposed on the subsequent purchaser. Published by Science Publishers, Enfield, NH, USA An imprint of Edenbridge Ltd. Printed in India

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

LE Reproductive Biology and Phylogeny of Birds ‘Natural History of Selborne’, and my good fortune in being joined by a most distinguished group of authors who need no introduction to those familiar with avian studies. A volume in preparation is on Cetacea (Debra Miller). My thanks are due to the School of Integrative Biology, University of Queensland, for facilities, and especially to the Executive Dean, Biological & Chemical Sciences, Professor Mick McManus, for his continuing encouragement. I am everlastingly indebted to Sheila Jamieson, who has supported me indirectly in so many ways in this work. I am grateful to the publishers for their friendly support and high standards in producing this series. Sincere thanks must be given to the volume editors and the authors, who have freely contributed their chapters, in very full schedules. The editors and publishers are gratified that the enthusiasm and expertise of these contributors has been reflected by the reception of the series by our readers.

THE UNIVERSITY OF QUEENSLAND AUSTRALIA

21 February 2006

Barrie G.M. Jamieson School of Integrative Biology University of Queensland

Classification and Phylogeny

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Preface to this Volume There are almost ten thousand known species of birds of which more than half are song birds. They are an ideal subject of study as they are one of the few groups of animals for which almost the total number of species is estimated to be known, they have been comprehensively catalogued and illustrated and are readily identified in the field whereas groups such as annelids (Volume 5) require detailed microscopical work for identification. Besides these practical qualifications for study and the biological questions that they pose, they have endeared themselves to humanity not least for their beauty and their song. This volume, in two parts, attempts to document most of the important aspects of the reproductive biology of birds and places them in a setting of phylogenetic relationships. Aspects of reproduction that comprise this, the first part of volume 6, are classification and phylogeny as revealed by molecular biology; anatomy of the male reproductive tract and organs; anatomy and evolution of copulatory structures; development and anatomy of the female reproductive tract; endocrinology of reproduction; ovarian dynamics and follicle development; spermatogenesis and testicular cycles; avian spermatozoa: structure and phylogeny; testis size, sperm size and sperm competition and, lastly, fertilization. We may here, albeit superficially, sample some of the many topics treated in the ten chapters. As more and more homologous DNA sequences and other genetic characters become available for more and more species, we are gradually resolving the remaining uncertain nodes of the avian phylogenetic tree, and this trend will only accelerate as DNA sequencing becomes both cheaper and more reliable. These are heady times in avian systematics and the present state of our knowledge is authoritatively reviewed. The testes of birds are intra-abdominal, and, in contrast with most mammals, they do not migrate from their site of embryological origin. They are, thus, closely related, topographically, to the kidneys. The anatomy and histology of avian testes and their ducts receive a detailed examination. In birds, unlike most other animal classes, males of some species possess an intromittent organ, whereas males of other species do not. Thus, in birds at least, the organ does not seem to be necessary for internal fertilization. This raises the question whether the avian intromittent organ has evolved as a

LEEE Reproductive Biology and Phylogeny of Birds primary sexual trait simply for the delivery of sperm or as a secondary sexual trait. Its absence in so many species is an evolutionary puzzle investigated here. Only the left genital primordia develop to functional ovaries in birds except in a few cases, particularly birds of prey. The reason for the unilateral development of female genital organs might be to reduce weight for flying. The question then arises why the falconiforms allow themselves the luxury of two genital tracts. To reduce weight their hard-shelled eggs are relatively small and are laid down at an early stage of development. This and other aspects of the female genital tract are examined. The Müllerian and Wolffian ducts receive detailed treatment and a valuable analysis of the function of the oviduct in egg laying is provided. In an innovative chapter hormonal control systems in birds are investigated in four major parts: 1) types of environmental signals that influence reproduction and how they are perceived; 2) the hypothalamus as an integrator of environmental information from the external and internal environments, biological clock, etc., that through neuroendocrine and neural secretions affects all aspects of reproduction; 3) the hypothalamo-pituitary unit that transduces environmental information processed by higher centers into endocrine secretions; 4) the functional gonads (ovary and testis) themselves. A unique morphological and functional aspect of the reproductively active avian ovary, as contrasted with the mammalian counterpart, is that follicles at all stages of development, from resting primordial and primary follicles to the fully differentiated preovulatory stage, exist simultaneously during egglaying. As a consequence, the sequential selection of one undifferentiated follicle into the final rapid growth stage of development provides for ovulation of an oocyte from a fully differentiated follicle on an approximate daily basis. The ovarian follicular hierarchy is a reflection of oviparity, and is a feature held in common with avian predecessors, the reptiles including, probably, some dinosaurs. Other aspects of ovarian function are examined. The seminiferous epithelium in avian testis is made up of germ cells at varying levels of development, and Sertoli cells. Sertoli cells have multiple functions, including formation of the blood-testis barrier by means of junctional complexes, and providing anchorage and nutrition for, as well as regulation of, germ cells during development. The most primitive or immature germ cells lie on the basement membrane and the mature germ cells, the spermatozoa, line the lumen of the seminiferous tubule. Germ cells develop in close association with one another because as they divide they maintain close linkage through intercellular bridges which are the result of incomplete cytoplasmic divisions. A detailed examination of spermatogenesis and factors affecting it is given from spermatogonia to the mature spermatozoa. The chapter on sperm structure and phylogeny is the longest of the book, in keeping with the great amount of information, though often fragmentary, which is available and the fact that the chapter represents the final scientific

Preface to this Volume

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contribution of any magnitude by the editor. Former workers recognized the ‘sauropsid’ features of non-passerine spermatozoa. However, the ratite and lower non-passerine spermatozoon, especially the former, are shown to more appropriately be termed crocodiloid. Features of Ostrich sperm which are similar to those of crocodiles are described. These features are also seen in turtles and are basic (symplesiomorphic) to amniotes, only the fibrous sheath and the long distal centriole being amniote synapomorphies. The sole spermatozoal synapomorphy of crocodiles and birds is the dense sheath investing the two central singlets within the elongate distal centriole. In birds this sheath is known only in ratites (Ostrich), galliforms and anseriforms. The literature is reviewed in a phylogenetic context. Almost since the moment that animal semen was first viewed under the microscope, in the seventeenth century, it was clear that the shape and size of spermatozoa varied from one species to the next. Early comparative biologists similarly noted striking differences in the size of the testis across different species. Although the general features of the male reproductive system are rather conservative across extant species of birds, recent work has confirmed large interspecific variation in the sizes of both testes and sperm. The chapter addresses the question why should testes mass and sperm size differ so much from one species to the next? Until recently, the cellular and molecular events comprising fertilization in birds were poorly understood. However, several recent studies in mammals as well as in birds have considerably contributed to our knowledge of the fertilization process. In mammals only a single spermatozoon enters the egg. Monospermy is ensured by a variety of mechanisms, which is collectively known as the block to polyspermy. Unlike mammals, the principal feature of fertilization in birds is physiological polyspermy, penetration of the ovum by many sperm. It has been suggested that polyspermy occurs in animals that produce large yolky (megalecithal) ova and this theme is developed. The structure of the mature ovum and the events of fertilization are discussed. There is an extensive section on assisted reproductive technologies. Many new illustrations are provided throughout the volume. The generosity of authors and publishers who have allowed illustrations to be reproduced is most gratefully acknowledged. The kind collaboration of all the authors, who have borne uncomplainingly the requests of an editor overseeing the gestation of this work, has been greatly appreciated. Finally, the courteous and efficient participation of the publishers was indispensable to production of this volume.

THE UNIVERSITY OF QUEENSLAND AUSTRALIA

21 February 2006

Barrie G.M. Jamieson School of Integrative Biology University of Queensland

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Contents Preface to the Series—Barrie G. M. Jamieson Preface to this Volume—Barrie G. M. Jamieson

v vii

1. Classification and Phylogeny of Birds John Harshman

1

2. Anatomy of the Testis and Male Reproductive Tract Tom A. Aire

37

3. Anatomy and Evolution of Copulatory Structures Robert Montgomerie and James Briskie

115

4. Developmental Anatomy of the Female Reproductive Tract Monika Jacob and Murray R. Bakst

149

5. Endocrinology of Reproduction G. E. Bentley, K. Tsutsui and J. C. Wingfield

181

6. Ovarian Dynamics and Follicle Development A. L. Johnson and Dori C. Woods

243

7. Spermatogenesis and Testicular Cycles Tom A. Aire

279

8. Avian Spermatozoa: Structure and Phylogeny Barrie G. M. Jamieson

349

9. Testis Size, Sperm Size and Sperm Competition James V. Briskie and Robert Montgomerie

513

10. Fertilization Urszula Stepinska and Murray R. Bakst Index

553

589

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About the Series This series bears the title ‘Reproductive Biology and Phylogeny’ followed by the name of the taxonomic group which is the subject of the volume. B. G. M. Jamieson is the founding series editor and each publication has one or more invited volume editors (if not the series editor) and a large number of authors of international repute. The series gives a unique coverage of reproductive biology. The level of the taxonomic group which is the subject of each volume varies according, largely, to the amount of information available, the advice of proposed volume editors, and the interest expressed by the zoological community in the proposed work. Phylogeny and classification is covered as a necessary framework for an understanding of reproductive biology. Evidence for relationships from molecular studies is an important aspect of the phylogenetic section. About the Volume There are almost ten thousand known species of birds of which more than half are song birds. They are an ideal subject of study as they are one of the few groups of animals for which almost the total number of species is estimated to be known, they have been comprehensively catalogued and illustrated and are readily identified in the field. Besides these practical qualifications for study and the biological questions that they pose, they have endeared themselves to humanity not least for their beauty and their song. This volume, in two parts, attempts to document most of the important aspects of the reproductive biology of birds and places them in a setting of phylogenetic relationships. Aspects of reproduction that comprise this, the first part of volume 6, are classification and phylogeny as revealed by molecular biology; anatomy of the male reproductive tract and organs; anatomy and evolution of copulatory structures; development and anatomy of the female reproductive tract; endocrinology of reproduction; ovarian dynamics and follicle development; spermatogenesis and testicular cycles; avian spermatozoa: structure and phylogeny; testis size, sperm size and sperm competition and, lastly, fertilization. Many new illustrations are included. About the Series and Volume Editor Dr. Barrie Jamieson is Emeritus Professor of Zoology in the School of Integrative Biology, University of Queensland. He holds a Ph.D. from the University of Bristol, England, a D.Sc. from the University of Queensland, and is a former Visiting Fellow of, and member of the Association of Corpus Christi College, Cambridge. In 1990 he was awarded the Clarke Medal for Research in Natural Sciences, early recipient of which were Thomas Henry Huxley and Richard Owen. His chief field of research is spermatozoal ultrastructure and its relevance to phylogeny but he is also an authority on taxonomy of earthworms and has published on bioluminescence, trematode taxonomy and life cycles, and DNA-based phylogenetics. He has published nearly 200 scientific papers and is the author, coauthor or editor of fourteen books.

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CHAPTER

1

Classification and Phylogeny of Birds John Harshman

1.1

INTRODUCTION

Avian phylogenetics is currently in its infancy, and paradoxically it is also nearing maturity. By infancy I mean that we do not currently know much about the relationships among birds, and by maturity I mean that within a few years we will know most of what there is to know. The reason for this apparent paradox lies in the rapidly increasing ease of gathering and analyzing molecular data. Our ignorance is largely a matter of our not yet having gathered enough DNA sequences or other molecular data for enough species, and that condition is being remedied at an increasing rate. If I had written this chapter a year or two from now, I would have been able to cite studies incorporating perhaps twice the data so far in published form—this purely from yet-unpublished studies of whose existence I am now aware— and I would have been able to say much more about the structure of the avian tree. But the editor was not anxious to follow that schedule, and so we press on with what is currently available. During more than two thousand years of avian systematics—counting from Aristotle—researchers were able to divide birds into a number of robust groups based on distinctive morphologies: the non-passerine families. (We will get to passerines in a moment.) This process was largely completed by the beginning of the last century. For reviews see Sibley and Ahlquist (1990) and Cracraft et al. (2004). Progress since then, on relationships between and within families, has been highly incremental, so much so that at least one prominent avian systematist (Stresemann 1959) announced that no further progress was possible. The cladistic revolution did little to help, though at least it focused our attention on the right questions; apparently it’s very difficult to find reliable morphological characters that will build a robust tree of birds. And the first 30 years of the molecular revolution have illuminated only a few of 4869 Pepperwood Way, San Jose, CA 95124; E-mail: [email protected]

Reproductive Biology and Phylogeny of Birds

those questions (Sheldon and Bledsoe 1993), leading another pair of systematists, once again, to wonder if there would be no further progress (Poe and Chubb 2004). However, recent discoveries suggest that there is more reason for optimism. This chapter will concentrate on what we do know confidently about avian relationships. Necessarily, it must treat mostly relationships between larger groups, since the nearly 10,000 living species are too many to be covered in detail. It is also convenient to divide the birds into two groups, only one of them monophyletic: passerines and non-passerines. One reason for separate treatment is that it splits the species roughly in half, but the main reason is that our state of knowledge differs radically between the groups. For nonpasserines, most traditional families are clearly supported as monophyletic, and we can summarize relationships as being between these entities. (The same is not true of traditional orders, as we will see.) I have used the classification of del Hoyo et al. (1992-2002) as representative of traditional familial and ordinal assignments. For passerines, however, most traditional families that have been adequately sampled are not monophyletic, and we must either speak in extremely general terms or choose individual species as our units of discussion. In both groups, genera have not fared well, and many have proven not to be monophyletic when examined closely. Non-monophyly should come as no surprise. Traditional taxonomy did not take monophyly into account. What mattered was distinctness, and whether that distinctness arose from apomorphy or plesiomorphy was not considered important. Thus if some subset of a group possessed a prominent apomorphy, that served to characterize two subgroups: one by its presence, and another by its absence. This practice has continued even to the present, e.g. the vigorous defense by Short and Horne (2002) of separating Ramphastidae from Capitonidae. One caveat should be kept in mind when assessing the results of phylogenetic analyses: they can discern the relationships only among those species actually included in the analysis. This rule may seem obvious, but it has been violated frequently in the literature. If an analysis of gruiform phylogeny includes only gruiforms and a single outgroup, the monophyly of gruiforms cannot be tested. If an analysis of bird phylogeny includes only one representative from each order, the monophyly of orders cannot be tested. If an attempt to find the sister group of hoatzins does not include the actual sister group (whatever it may be), that relationship will not be found. It would thus seem that every phylogenetic analysis, in order to be valid, must include every species of bird (and that further assumes that birds are monophyletic, and that each individual species is also), which is clearly impossible. Fortunately, chaos is not total, and uncertainties are of limited taxonomic scope: families whose ordinal relationships are unclear have themselves so far proven to be monophyletic; all members of non-monophyletic families can be constrained within single orders, and all members of non-monophyletic genera within single families. (The latter does not apply to passerines, however.) Some

Classification and Phylogeny of Birds

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groups do have clear morphological synapomorphies, and these can be easy to assay across species. Genetic distances offer some comfort too. If we find all species of Anas to be genetically similar, it’s unlikely that any of them is really a bustard, even if we do not include them all in an analysis with a bustard. But we must still be careful not to overinterpret any results. Because of their importance to avian systematics, I must discuss the DNA hybridization studies of Sibley and Ahlquist (1990) and the classification based on them (Sibley and Monroe 1990). Their final product, the “Tapestry”, is a comprehensive and nearly fully resolved tree of over 1100 species (Sibley and Ahlquist 1990, figs. 357-385). However important, this tree was severely flawed as a representation of their data, and the data themselves were insufficient to resolve objectively many of the relationships asserted by the Tapestry (Lanyon 1992; Harshman 1994). A later attempt to correct some of the flaws of the Tapestry by using rigorous methods of analysis on subsets of the data (Sibley and Ahlquist 1990, figs. 325-352)—since they used the FitchMargoliash (1967) method of distance analysis, we may call them “Fitch trees”—is an improvement but suffers from the absence of an evaluation of strength of support, which Harshman (1994) attempted to repair by reanalyzing the data. These latter analyses should be preferred to the Tapestry in the several cases of conflict. Sibley and Ahlquist’s studies contain some exciting, corroborated insights, e.g., the sister relationship of mockingbirds (Mimidae) and starlings (Sturnidae), but also some major mistakes, e.g., the placement of storks (Ciconiidae) and New World vultures (Cathartidae) as sister taxa, and even some decisions unjustified by any data, e.g., the placement of hoatzin (Opisthocomidae) with cuckoos (Cuculiformes). In the end, without corroboration, it is difficult to separate the Tapestry’s good estimates from the bad, even with access to their original data (courtesy of the late C. G. Sibley). Below, I will mention Sibley and Ahlquist’s findings only when they have been independently corroborated or if their conclusions are clearly supported by their data, properly analyzed.

1.2

NON-PASSERINES

Relationships among the non-passerine families are not well resolved at present. If we consider a tree of relationships among the 103 non-passerine families recognized by del Hoyo et al. (1992-2002) plus the order Passeriformes, that tree, if fully resolved, will have 103 internal nodes (counting the root position). We can be fairly confident of only 60 of those nodes (Figs. 1.1 and 1.2). In contrast to many other taxa, the most basal relationships are among the best known, but most other resolved nodes unite small numbers of closely related families, such as Dromaiidae (emu) and Casuariidae (cassowaries).

1.2.1

Basal Relationships

Modern birds—Aves or Neornithes depending on one’s taste in terminology (Gauthier et al. 2001)—are divided basally into two clades, Palaeognathae and

" Reproductive Biology and Phylogeny of Birds

Fig. 1.1 Relationships among non-passerine families, so far as they can be confidently asserted at present according to my perhaps biased estimate. This tree does not derive from a formal analysis, either of a combined supermatrix or by a rigorous supertree method. Branches in gray represent paraphyletic groups. This figure shows Paleognathae through Metaves. For Coronaves see Fig. 1.2. For reasons behind the topology chosen, see text.

Neognathae. Palaeognathae includes the ratites and tinamous. Neognathae is divided into Galloanserae, consisting of the sister orders Anseriformes and Galliformes, and Neoaves, consisting of all other birds. Recent analyses, both molecular and morphological, have been nearly unanimous on these four basal clades (Prager and Wilson 1978; Caspers et al. 1997; Groth and Barrowclough 1999; García-Moreno and Mindell 2000; van Tuinen et al. 2000; Paton et al. 2002; Cracraft and Clarke 2001; García-Moreno et al. 2003; Mayr and Clarke 2003; Cracraft et al. 2004). Some other studies lacked an outgroup root but are consistent with these clades (Sibley and Ahlquist 1990; Chubb 2004a; Fain and Houde 2004). A few studies using whole mitochondrial genomes or large fractions thereof have found a different topology, in which passerines are basal to all other

Classification and Phylogeny of Birds

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Fig. 1.2 Relationships among non-passerine families, continued. Coronaves only. For reasons behind the topology chosen, see text.

$ Reproductive Biology and Phylogeny of Birds birds (Mindell et al. 1997; Härlid and Arnason 1999; Mindell et al. 1999). But these have been shown to be the result of long branch attraction (Braun and Kimball 2002; Paton et al. 2002; García-Moreno et al. 2003). Also see Ericson et al. (2001), which failed to find a monophyletic Galloanserae. A note on non-monophyly: there are two forms of non-monophyly, paraphyly and polyphyly. Strictly speaking, the two cannot be distinguished on a phylogenetic tree, since the definitions require either explicit assignment of ancestral nodes to groups or optimization of diagnostic characters. In this chapter, I will define paraphyly artificially: if we code group membership as a presence/absence character, a group is paraphyletic if that state can be optimized as gained only once on the tree and lost one or more times. It is polyphyletic if group membership must be optimized as being gained at least twice. This favors paraphyly in ambiguous cases. Results may differ from those under other definitions, but at least the condition can be determined from the tree itself.

1.2.2 Palaeognathae Almost all studies using non-avian outgroups have found paleognaths to be monophyletic (García-Moreno and Mindell 2000; van Tuinen et al. 2000; Paton et al. 2002; Cracraft et al. 2004; but see Chapter 8). Under that assumption, we can add studies using neognath outgroups, and these have found the two paleognath orders, Struthioniformes (ratites) and Tinamiformes (tinamous) also to be monophyletic (Sibley and Ahlquist 1990; Lee et al. 1997; van Tuinen et al. 1998; Cooper et al. 2001; Cracraft and Clarke 2001; Haddrath and Baker 2001). Within ratites, however, there is considerable contention. It is universally agreed in all these studies that Dromaiidae (emu) and Casuariidae (cassowaries) are sister taxa. Molecular studies commonly make Apterygidae (kiwis) the sister of these two, forming an Australasian clade (Sibley and Ahlquist 1990; Lee et al. 1997; van Tuinen et al. 1998; Cooper et al. 2001; Haddrath and Baker 2001). The question of whether the Rheidae (rheas) or Struthionidae (ostrich) is the sister of all other ratites is less clear; the majority of studies have found the rhea basal (Cooper et al. 1992; Lee et al. 1997; Cooper et al. 2001; Haddrath and Baker 2001), but others have found the ostrich basal (Sibley and Ahlquist 1990; van Tuinen et al. 1998). The sole morphological study has the same unrooted topology as the molecular studies, but the root attaches to the kiwi, making the rooted topology quite different (Lee et al. 1997). Phylogenetic relations within tinamous have been little studied. There is only one published morphological phylogeny (Bertelli et al. 2002) and no molecular studies.

1.2.3 Galloanserae Monophyly of Galloanserae is affirmed by a large number of studies (Sibley and Ahlquist 1990; Caspers et al. 1997; Livezey 1997; Groth and Barrowclough 1999; García-Moreno and Mindell 2000; van Tuinen et al. 2000;

Classification and Phylogeny of Birds

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Zusi and Livezey 2000; Cracraft and Clarke 2001; Paton et al. 2002; GarcíaMoreno et al. 2003; Mayr and Clarke 2003; Sorenson et al. 2003; Cracraft et al. 2004). But see Ericson (1997) and Ericson et al. (2001). There is also strong support for monophyly of both Galliformes and Anseriformes (Livezey 1986; 1997a; Cracraft and Clarke 2001; Zusi and Livezey 2000; Ericson et al. 2001; Sorenson et al. 2003; Chubb 2004a; Cracraft et al. 2004), though molecular studies with large taxon samples of both orders have not been published. Sibley and Ahlquist (1990) found strong support for monophyly of Galliformes, but monophyly of Anseriformes could not be confirmed (Harshman 1994), and Chubb (2004a) also could not confirm anseriform monophyly.

1.2.3.1

Relationships within Galliformes

All families are monophyletic except Phasianidae (pheasants), which includes the two traditional families Meleagrididae (turkeys) and Tetraonidae (grouse) (Dimcheff et al. 2002). The monophyly of Numididae (guineafowl) is not supported by morphological characters (Dyke et al. 2003), but at least two of the four genera have been shown to be closely related by DNA hybridization (Sibley and Ahlquist 1990). Basal relationships are clear: Megapodiidae (megapodes) and Cracidae (curassows, guans, and chachalacas) are successive sister groups to the rest (Sibley and Ahlquist 1990—the Fitch tree, contradicting the Tapestry; Ericson et al. 2001; Dimcheff et al. 2002; Dyke et al. 2003; Sorenson et al. 2003; Mayr and Weidig 2004; Pereira and Baker 2006). The major puzzle is whether the sister group of Phasianidae is Odontophoridae (New World quail) (Armstrong et al. 2001; Dimcheff et al. 2002) or Numididae (Sibley and Ahlquist 1990; Kornegay et al. 1993; Kimball et al. 1999; Armstrong et al. 2001; Dimcheff et al. 2002; Pereira and Baker 2006). Some studies support both possibilities in different analyses. There are several studies of relationships within families: megapodes (Birks and Edwards 2002), cracids (Pereira et al. 2002), and phasianids (Kimball et al. 1999; Dimcheff et al. 2002). The phasianid genus Francolinus is polyphyletic (Bloomer and Crow 1998), and the tetraonid genera Bonasa and Falcipennis are paraphyletic (Ellsworth et al. 1996; Dimcheff et al. 2002; Drovetski 2002).

1.2.3.2

Relationships within Anseriformes

Monophyly of Anhimidae (screamers) is easily confirmed, as the three species are quite closely related (Livezey 1986; Sibley and Ahlquist 1990). Sibley and Ahlquist’s (1990) Tapestry makes Anseranas (magpie goose, sometimes given its own monotypic family Anseranatidae) the sister group of Anhimidae, thus making Anatidae (ducks) paraphyletic, but other analyses of the same data (Sibley and Ahlquist 1990’s Fitch tree; Harshman 1994) restore duck monophyly. In all other analyses, Anatidae is monophyletic (Livezey 1986; 1997a; Ericson 1997; Ericson et al. 2001; Cracraft et al. 2004). Relationships within Anatidae are contentious. Though the basal relationships are well established (Anseranas and Dendrocygninae as successive

& Reproductive Biology and Phylogeny of Birds sister groups to the remaining anatids), the remaining relationships differ greatly between morphological (Livezey 1997b and references therein) and molecular (Madsen et al. 1988; Sraml et al. 1996; Donne-Goussé et al. 2002; Callaghan and Harshman 2005 and references therein) analyses. Only one genus has been shown to be paraphyletic. As might be expected, this is the largest, Anas, which includes the steamer ducks (Tachyeres) as well as three monotypic genera that are sometimes merged with Anas (Amazonetta, Speculanas, Lophonetta) (Johnson and Sorenson 1999).

1.2.4

Neoaves

Monophyly of Neoaves, also called Plethornithes by Groth and Barrowclough (1999), has been confirmed in all recent studies (Groth and Barrowclough 1999; Sorenson et al. 2003; Cracraft et al. 2004; Fain and Houde 2004). It encompasses the great majority of all birds, including Passeriformes.

1.2.4.1

Monophyly of neoavian orders

There are 23 non-passerine orders within Neoaves. Of these, 9 consist of just a single family, and their monophyly is trivial. Two more orders, Psittaciformes (parrots) and Strigiformes (owls) each consist of just two closely related families, the psittaciform pair often merged into a single family, Psittacidae. Thus there are 12 orders for which monophyly is a serious question. Of these, we have good evidence of monophyly for four, good evidence against monophyly for five, and no strong evidence either way for three. The single-family orders are Sphenisciformes (penguins), Gaviiformes (loons/divers), Podicipediformes (grebes), Phoenicopteriformes (flamingos), Opisthocomiformes (hoatzin), Pterocliformes (sandgrouse), Columbiformes (pigeons), Coliiformes (colies), and Trogoniformes (trogons). The clearly monophyletic orders (other than those with only one family) are Procellariiformes (tubenoses), Apodiformes (swifts and hummingbirds), Galbuliformes (puffbirds and jacamars), and Piciformes (woodpeckers, barbets, and honeyguides). The monophyly of both Pelecaniformes and Ciconiiformes is falsified by the close relationship among the pelecaniform family Pelecanidae (pelicans) and the two ciconiiform families Scopidae (hamerkop) and Balaenicipitidae (shoebill) (Hedges and Sibley 1994; Siegel-Causey 1997; van Tuinen et al. 2001; Cracraft et al. 2004; Fain and Houde 2004; but see Mayr 2003). There remains a group that we might call “core pelecaniforms” except that it fails to include pelicans, consisting of Sulidae (boobies), Phalacrocoracidae (cormorants), Anhingidae (darters), and Fregatidae (frigatebirds) (Sibley and Ahlquist 1990; Harshman 1994; Cracraft et al. 2004). Relationships of the remaining traditional pelecaniform family, Phaethontidae (tropicbirds), are unclear, as are relationships of the remaining traditional ciconiiforms. Caprimulgiformes is at least paraphyletic, since one caprimulgiform family, Aegothelidae (owlet-nightjars) is the sister of Apodiformes (Mayr 2002a; Cracraft et al. 2004); whether the order would be monophyletic if apodiforms were included is unclear.

Classification and Phylogeny of Birds

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The monophyly of both Gruiformes and Charadriiformes is falsified by the discovery that one gruiform family, Turnicidae (buttonquails) is deeply nested within charadriiforms (Paton et al. 2003). However, charadriiforms are monophyletic with the addition of turnicids (Paton et al. 2003). But there is no evidence for monophyly of gruiforms minus turnicids, and some evidence against it (Fain and Houde 2004). There is a group we might call “core gruiforms”, consisting of Gruidae (cranes), Aramidae (limpkin), Psophiidae (trumpeters), Rallidae (rails), and Heliornithidae (finfoots) (Houde et al. 1997; Livezey 1998; Cracraft et al. 2004; Fain and Houde 2004). For another view, see Livezey (1998), in which both Turnicidae and Pedionomidae (plains wanderer) are included in a monophyletic Gruiformes. There is some morphological evidence for monophyly of Falconiformes (Griffiths 1994), but no molecular analyses have so far succeeded in putting all families together. Nor, however, is there strong evidence for any falconiform family’s relationship to any non-falconiform family. The hypothesized relationship between Cathartidae (New World vultures) and Ciconiidae (storks) (e.g., Sibley and Ahlquist 1990) has not survived rigorous analysis. There is no evidence for monophyly of Coraciiformes, though there is some support for a “core coraciiforms” including Alcedinidae (kingfishers), Todidae (todies), Momotidae (motmots), Meropidae (bee-eaters), Coraciidae (rollers), and Brachypteracidae (ground-rollers) (Johansson and Ericson 2003; Cracraft et al. 2004). There are so far only two studies that have included Leptosomidae (cuckoo-roller), and there was no support for its inclusion in or exclusion from Coraciiformes (Kirchman et al. 2001; Mayr et al. 2003). The position shown on Sibley and Ahlquist’s (1990) Tapestry is not based on valid data (C. G. Sibley, unpublished data). The remaining three coraciiform families, Upupidae (hoopoe), Phoeniculidae (woodhoopoes), and Bucerotidae (hornbills) also form a clade, but their relationship to core coraciiforms is unsupported (Cracraft et al. 2004). There is no strong evidence for monophyly of Cuculiformes, consisting of Cuculidae (cuckoos) and Musophagidae (turacos) (Sorenson et al. 2003).

1.2.4.2

Relationships between neoavian orders (or their monophyletic fragments)

Confirmed relationships between orders are few. Perhaps the most interesting published hypothesis is that of Fain and Houde (2004), who divided Neoaves into two basal clades: Metaves, consisting of Caprimulgiformes, Apodiformes, Podicipedidae, Phaethontidae, Phoenicopteridae, Opisthocomidae, Mesitornithidae, Rhynochetidae, Eurypygidae, Pteroclidae, and Columbidae; and Coronaves, consisting of the remaining Neoaves. So far, these clades are supported entirely by analyses of beta-fibrinogen, intron 7, but it is encouraging that both sequence characters and indel characters, which evolve largely by independent processes, support the same clades. In addition, no rigorous studies contradict this hypothesis. After some internal debate, I have

 Reproductive Biology and Phylogeny of Birds added these groups to Figs. 1.1 and 1.2. Note that, if accepted, these two clades make Pelecaniformes polyphyletic (through the inclusion of Phaethontidae in Metaves and the remaining families in Coronaves) and Gruiformes highly polyphyletic. One amply confirmed though surprising relationship is that between Phoenicopteridae (flamingos) and Podicipedidae (grebes). First discovered by van Tuinen et al. (2001), it has since been confirmed by many authors (Chubb 2004a; Cracraft et al. 2004; Mayr 2004). Another well confirmed relationship is that between Aegothelidae (owletnightjars) and Apodiformes (Mayr 2002a; Mayr et al. 2003; Cracraft et al. 2004). The traditional relationship of Galbuliformes and Piciformes has recently been questioned strongly enough that the two orders, generally united as Piciformes, were separated in the classification used by del Hoyo et al. (19922002). However, that relationship has subsequently been conclusively confirmed (Johansson and Ericson 2003; Mayr et al. 2003; Cracraft et al. 2004). The long-standing hypothesis that Procellariiformes and Sphenisciformes are sister groups has received some support in molecular analyses (van Tuinen et al. 2001); but see Mayr (2005) for a placement of penguins within “core pelecaniforms”. One more possible grouping, though it has ambiguous support and its boundaries are not clearly defined, is sometimes called “water birds”. It includes most of the traditional members of Pelecaniformes (except Phaethontidae), Ciconiiformes, and Procellariiformes, plus Spheniscidae (penguins) and Gaviidae (loons) (van Tuinen et al. 2001; Cracraft et al. 2004). Support in any individual analysis is not strong, however, and some analyses include additional groups within the clade (e.g., Cuculiformes, “core gruiforms”), while other analyses do not sample all groups. Because of the ambiguity surrounding this potential group, I have omitted it from the tree (Fig. 1.2). One prominent hypothesis that has no support is a clade made up of any combination of Opisthocomidae, Musophagidae, and Cuculidae. Groups uniting different pairs or all three have been proposed several times (Sibley and Ahlquist 1990; Hedges et al. 1995; Hughes 1996, 2000; Hughes and Baker 1999), but they have been plagued by data problems (Sorenson et al. 2003), and as of now no resolution is possible. Note that in the analysis by Fain and Houde (2004), the hoatzin belongs to Metaves, while turacos and cuckoos are Coronaves. There are a few other hypotheses with some support from morphological data that either contradict or are unconfirmed by molecular data. Mayr (2004b) proposed a sister group relationship between Mesitornithidae (mesites) and Cuculidae (cuckoos). Mayr (2003b) proposed a sister group relationship between Trogonidae (trogons) and Steatornithidae (oilbirds). Both relationships contradict Fain and Houde’s (2004) Metaves/Coronaves split, though they do not contradict any strongly supported clades in any other analyses.

Classification and Phylogeny of Birds

1.2.4.3



Relationships within neoavian orders (or their monophyletic fragments)

Procellariiformes. Pelecanoididae (diving-petrels) are probably nested within Procellariidae (petrels and shearwaters) (Cracraft et al. 2004), so I show them as sister taxa in Fig. 1.2. This is the only resolved node in my reanalysis (Harshman 1994) of Sibley and Ahlquist’s (1990) data. Hydrobatidae may be polyphyletic (see below) and this prevents any further resolution among families. Pelecaniformes. This order is divided into three pieces. Phaethontidae (tropicbirds) is apparently not closely related to any of the other families. The relationships of Pelecanidae (pelicans) have been noted above. This leaves a clade I will call “core pelecaniforms” despite the absence of pelicans from the group, composed of Fregatidae (frigatebirds), Sulidae (boobies), Phalacrocoracidae (cormorants), and Anhingidae (darters). The only resolved node in my reanalysis (Harshman 1994) of Sibley and Ahlquist’s (1990) data unites these four families. Siegel-Causey (1997) and Cracraft et al. (2004, fig. 27.4) also find this grouping. In other studies (van Tuinen et al. 2001; Cracraft et al. 2004, fig. 27.7) only the last three are united or Fregatidae is only weakly attached to the others (Cracraft et al. 2004, fig. 27.6). Relationships among Sulidae, Phalacrocoracidae, and Anhingidae are ambiguous, with either phalacrocoracids (van Tuinen et al. 2001; Fain and Houde 2004) or sulids (Siegel-Causey 1997; Cracraft et al. 2004) sister to the other two. Falconiformes. Though the order cannot be confidently asserted as monophyletic, relationships are well established among families of the suborder Accipitres, with Sagittariidae (secretary bird) the sister of Pandionidae (osprey) and Accipitridae (hawks) (Cracraft et al. 2004; Fain and Houde 2004); but see Griffiths (1994). Gruiformes. The relationships of Otididae (bustards), Cariamidae (seriemas), and Mesitornithidae (mesites) to any other families (gruiform or otherwise) cannot be determined at present. Turnicidae (buttonquail) belongs to Charadriiformes (see above). Eurypygidae (sunbittern) and Rhynochetidae (kagu) are sister taxa (Houde et al. 1997; Livezey 1998; Cracraft et al. 2004; Fain and Houde 2004), but their relationships to other families cannot be determined. This leaves a “core gruiforms” consisting of Gruidae (cranes), Aramidae (limpkin), Psophiidae (trumpeters), Rallidae (rails), and Heliornithidae (finfoots), for which there is strong support (Houde et al. 1997; Livezey 1998; Cracraft et al. 2004; Fain and Houde 2004). There is strong support for a sister group relationship between Gruidae and Aramidae and between Rallidae and Heliornithidae (Houde 1994; Houde et al. 1997; Livezey 1998; Cracraft et al. 2004; Fain and Houde 2004). The placement of Psophiidae is disputed, but the only strong support is for a position as sister of Gruidae plus Aramidae (Livezey 1998; Fain and Houde 2004). Charadriiformes. All molecular analyses (Sibley and Ahlquist 1990; Ericson et al. 2003; Paton et al. 2003; Fain and Houde 2004) are agreed on the



Reproductive Biology and Phylogeny of Birds

relationships of families within Charadriiformes. Paton et al. (2003) discuss incongruence between their tree and that of Sibley and Ahlquist (1990), but they are referring there to the Tapestry, while I refer here to the more rigorous Fitch tree (Sibley and Ahlquist 1990; Harshman 1994). The order is traditionally divided into three suborders, Charadrii, Lari, and Alcae (del Hoyo et al. 1996), but Alcae is nested within Lari, and the traditional Charadrii is polyphyletic. The three basal clades do consist of plover-like birds, gull-like birds, and sandpiper-like birds, which might be given the names Charadrii, Lari, and Scolopaci, respectively, with the first being sister to the other two. Ibidorhynchidae (ibisbill) has never been sampled and cannot be placed. Lacking any better notion, I have placed it at the basal node of Charadriiformes. Caprimulgiformes. There is no study strongly supporting the relationship of any caprimulgiform family to any other (Harshman 1994; Mariaux and Braun 1996; Fidler et al. 2004). The only supported relationship is that of Aegothelidae to Apodiformes, as discussed above. Apodiformes. Remarkably few studies have included representatives of all three families of apodiforms, but those that did have unsurprisingly found Apodidae (swifts) and Hemiprocnidae (treeswifts) to be sister taxa (Johansson et al. 2001; Mayr 2002a; Mayr et al. 2003; Chubb 2004a). Coraciiformes. Relationships within “core coraciiforms” are partially resolved. One clade is composed of Alcedinidae, Todidae, and Momotidae, though relationships among these families are contradictory (Sibley and Ahlquist 1990; Harshman 1994; Espinosa de los Monteros 2000; Johansson et al. 2001; Johansson and Ericson 2003), and another clade of Coraciidae and Brachypteracidae (Johansson et al. 2001; Kirchman et al. 2001; Johansson and Ericson 2003). The position of Meropidae is unresolved, though there is some evidence that it is sister to the first clade (Johansson and Ericson 2003). There is another clade of traditional coraciiforms whose relationships to core coraciiforms are unresolved; it consists of Bucerotidae, Upupidae, and Phoeniculidae, in which the last two are sister taxa (Sibley and Ahlquist 1990; Harshman 1994; Johansson et al. 2001; Cracraft et al. 2004). Leptosomidae certainly does not belong to the roller clade and probably does not belong to the core coraciiforms (Kirchman et al. 2001). Piciformes. There is a basal split between Capitonidae (which includes Ramphastidae) and the other two families, Indicatoridae and Picidae (Sibley and Ahlquist 1990; Harshman 1994; Johansson and Ericson 2003; Cracraft et al. 2004).

1.2.4.4

Monophyly of neoavian families

Most non-passerine, neoavian families have clear morphological synapomorphies, and the monophyly of many has been supported by molecular analyses. But there are a few exceptions. Charadriidae (plovers) is both paraphyletic, since it includes the two families Recurvirostridae (stilts

Classification and Phylogeny of Birds

!

and avocets) and Haematopodidae (oystercatchers) (Ericson et al. 2003; Fain and Houde 2004) and polyphyletic, since Pluvianellus (Magellanic plover) is not a close relative of the other plovers, but is instead sister to Chionidae (sheathbills) (Paton et al. 2003). Glareolidae (coursers and pratincoles) is polyphyletic, since Pluvianus (Egyptian plover), as its common name might suggest, is not a member of the family; its actual position is not clear (Fain and Houde 2004). In parrots, Psittacidae is paraphyletic to Cacatuidae (de Kloet and de Kloet 2005). There is some evidence to suggest that the procellariiform family Hydrobatidae (storm petrels) is paraphyletic (Nunn and Stanley 1998) to the rest of the order, or even polyphyletic (Cracraft et al. 2004), but topologies are contradictory. I have represented this in Fig. 1.2 by making Hydrobatidae paraphyletic and its relationships to other families unresolved. There is also strong evidence that Procellariidae (petrels and shearwaters) is paraphyletic to Pelecanoididae (diving-petrels) (Nunn and Stanley 1998, Cracraft et al. 2004). And Capitonidae (barbets) is paraphyletic to Ramphastidae (toucans) (Sibley and Ahlquist 1990; Lanyon and Hall 1994; Barker and Lanyon 2000; Johansson and Ericson 2003; Cracraft et al. 2004; Moyle 2004). There may be other exceptions that we are unaware of and that will be exposed by increased taxon sampling, but I do not expect many.

1.2.4.5

Relationships within neoavian families

Space doesn’t permit a lengthy discussion of intrafamilial relationships. Instead I have provided a list of references (Table 1.1) that investigate such relationships with a large enough taxon sample to be useful. Table 1.1

References to studies of phylogeny within individual neoavian families

Family

Reference

Spheniscidae Diomedeidae Procellariidae Hydrobatidae Phaethontidae Fregatidae Ardeidae

Bertelli and Giannini 2005 Nunn and Stanley 1998 Nunn and Stanley 1998 Nunn and Stanley 1998 Kennedy and Spencer 2004 Kennedy and Spencer 2004 Sheldon 1987 Sheldon et al. 2000 Slikas 1997 Griffiths 1999 Griffiths 1999 Griffiths et al. 2004 Wink 1995 Krajewski and King 1996 Livezey 1998 Pitra et al. 2002 Broders et al. 2003

Ciconiidae Accipitridae Falconidae Cathartidae Gruidae Rallidae Otididae

Table 1.1 Contd. ...

" Reproductive Biology and Phylogeny of Birds Table 1.1 Contd. ...

Family

Reference

Jacanidae Scolopacidae

Whittingham et al. 2000 Ericson et al. 2003 Paton et al. 2003 Thomas et al. 2004 Braun and Brumfield 1998 Thomas et al. 2004 Crochet et al. 2000 Thomas et al. 2004 Pons et al. 2005 Friesen et al. 1996 Thomas et al. 2004 Johnson and Clayton 2000 Miyaki et al. 1998 Brown and Toft 1999 de Kloet and de Kloet 2005 Hughes and Baker 1999 Veron and Winney 2000 Aragón et al. 1999 Hughes and Baker 1999 Hughes 1996, 2000 Johnson et al. 2000 Sorenson et al. 2003 Mariaux and Braun 1996 Bleiweiss et al. 1997 Chubb 2004b Espinosa de los Monteros 2000 Johansson and Ericson 2004 Moyle 2005 Sibley and Ahlquist 1990 Johansson and Ericson 2003 Kirchman et al. 2001 Barker and Lanyon 2000 Johansson and Ericson 2003 Nahum et al. 2003 Moyle 2004 Prychitko and Moore 1997 Webb and Moore 2005

Stercorariidae Laridae

Alcidae Columbidae Psittacidae/ Cacatuidae

Musophagidae Cuculidae

Nyctibiidae Trochilidae Trogonidae

Alcedinidae Brachypteracidae Capitonidae/ Ramphastidae

Picidae

1.2.4.6

Monophyly of neoavian genera

There are a limited number of families in which taxon sampling has been dense enough to determine the monophyly of genera. But in those families, paraphyly of genera has been shown to be frequent. The sparse data so far suggest that the larger a genus, and the more genera within the family, the more likely it is that non-monophyly will be found. Most often, as with families, paraphyly results from aberrant species being assigned their own genera. But there are additional cases without such clear causes.

Classification and Phylogeny of Birds

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The greatest prevalence so far is in the most heavily sampled family with large genera, Picidae, in which at least six genera are non-monophyletic: Colaptes, Piculus, Picoides, Dendropicos, Veniliornis, and Picus (Weibel and Moore 2002; Webb and Moore 2005). Other examples include Eupodotis and Ardeotis in Otididae (Pitra et al. 2002; Broders et al. 2003), Columba and Streptopelia in Columbidae (Johnson and Clayton 1999; Johnson et al. 2001), Larus in Laridae (Pons et al. 2005), Tauraco in Musophagidae (Veron and Winney 2000), and Stercorarius in Stercorariidae (Braun and Brumfield 1998). We can expect many more non-monophyletic genera to come to light as species sampling increases.

1.2.5

Classification of Non-passerines

It would be premature to offer a new classification of the non-passerines, given our great current uncertainty and the hope that it will soon be repaired, but I can at least make a few recommendations for modification of the classification I have been using, from del Hoyo et al. (1992-2002). Paraphyletic and polyphyletic groups should be reconciled so as to make them monophyletic, and though Linnean ranks are arbitrary, we may as well avoid leaving traditional family-rank names nested within other family-rank names. Therefore sevaral families should be merged into the families that enclose them: Tetraonidae and Meleagrididae into Phasianidae, Pelecanoididae into Procellariidae, Haematopodidae and Recurvirostridae into Charadriidae, Cacatuidae into Psittacidae, and Ramphastidae into Capitonidae. It may be that Hydrobatidae will need to be split into two families, but more study is needed. Turnicidae should be moved from Gruiformes to Charadriiformes. Coraciiformes should be limited to the “core coraciiforms”, and a new order established for Bucerotidae, Upupidae, and Phoeniculidae, for which Bucerotiformes is a reasonable name. Likewise, Gruiformes should be limited to the “core gruiforms”. We should probably wait a while before redefining Pelecaniformes; perhaps a clade can be found that includes both pelicans and “core pelecaniforms”, though it will surely include other families too. Galbuliformes should be returned to its traditional state as a suborder of Piciformes.

1.3

PASSERINES

The order Passeriformes contains more than half the species of birds (Sibley and Monroe 1990). In contrast to the difficulties with non-passerines, molecular techniques have proven to be highly successful in determining relationships at all levels. The major difficulties have been the sheer number of species and the unreliability of previous taxonomies as guides to phylogeny. Thus, single species cannot be used as handy proxies for entire families, or even genera. The rules of thumb used for non-passerines—in which genera did not commonly change families, and non-monophyletic

$ Reproductive Biology and Phylogeny of Birds genera at least had their separate parts all within one family, etc.—cannot be counted on here. What that means for this review is that any summary of relationships will be difficult, as the species assigned to families and superfamilies are in flux, and new surprises are expected as more species are sampled. Further, it becomes even more important to consider the exact nature of the taxon sampling in any given study and its comparability to sampling in other studies. I will attempt only an impressionistic summary tree of higher taxa (Figs. 1.3 and 1.4), for whose suggested species membership the reader must consult the references provided. One monotypic traditional family, Hypocoliidae, has never been included in any molecular analysis or any rigorous, cladistic analysis of morphology, and it has been omitted from further discussion here. Many of the analyses used in assembling the passerine tree have used Bayesian methods (Huelsenbeck and Ronquist 2001). Recently it has become apparent that Bayesian support values can be highly inflated (Suzuki et al. 2002; Yang and Rannala 2005). Very small differences in data or choice of evolutionary model can produce contradictory but strongly supported clades (pers.obs.), and there are many contradictory clades in the sources used in this review that have high Bayesian support. Consequently, I have not accepted any relationships based solely upon Bayesian analysis of any single data set. Either there must be some other measure of support used (e.g., parsimony or likelihood bootstrap), or different data sets (including indel data if any) must agree on the topology. Neither has any contradiction in relationships supported by Bayesian analyses alone caused me to remove resolution from any clade.

1.3.1

Monophyly of Passeriformes

Passeriform monophyly is well supported by morphological apomorphies (Raikow 1982). There have been several molecular analyses with moderately large taxon samples of passerines and other orders, and passerine monophyly has always been found (Johansson et al. 2001; Johansson and Ericson 2003; Sorenson et al. 2003; Cracraft et al. 2004; Fain and Houde 2004).

1.3.2

Monophyly of Passerine Families and Genera

One reason that it’s difficult to summarize passerine relationships in the same way I did for non-passerines is that there are no convenient, easily recognized subtaxa to use in constructing a summary tree. Sibley and Ahlquist (1990) showed that many of the traditional passerine families were not monophyletic, and others since then have found that many of the groups with which Sibley and Ahlquist (1990) and Sibley and Monroe (1990) replaced those traditional groups are themselves not monophyletic (see below). Further, misplaced taxa are not limited to small movements, but may span almost the range of the passerine tree—“almost” because so far no putative suboscine has been shown to be oscine, or vice versa. However, species have moved between superfamilies. Pseudopodoces (Hume’s ground jay), supposed to be a

Classification and Phylogeny of Birds

%

member of Corvidae and thus of Corvoidea, is instead a member of Paridae (in fact within the genus Parus, which thereby becomes paraphyletic) and thus of Sylvioidea (James et al. 2003; Gill et al. 2005). Yuhina (Erpornis) zantholeuca (white-bellied yuhina) is not a yuhina (Sylvioidea) but a vireo (Corvoidea) (Barker et al. 2004). Macgregoria (Macgregor’s bird of paradise) is not a member of Paradiseidae (Corvoidea) but of Meliphagidae (Meliphagoidea) (Cracraft and Feinstein 2000). Sapayoa (broadbilled sapayoa) is not a New World suboscine but an Old World suboscine, in terms of phylogeny if not geography (Fjeldså et al. 2003; Chesser 2004). There is no reason to suppose that these examples exhaust the list. Traditional or recently defined (Sibley and Ahlquist 1990; Sibley and Monroe 1990) families that have proven non-monophyletic include Eurylaimidae (e.g., Prum 1993), Formicariidae (traditional and Sibley and Ahlquist versions), Rhinocryptidae, Furnariidae, Tyrannidae, Cotingidae, and Pipridae, in fact the large majority of suboscine families (Sibley and Ahlquist 1990; Irestedt et al. 2002; Chesser 2004). Oscines are similar in their proportion of non-monophyletic families, though I will not list them here (e.g. Sibley and Ahlquist 1990; Cibois and Cracraft 2004; Voelker and Spellman 2004; Beresford et al. 2005; Alström et al. 2006). Genera of passerines are also not reliably monophyletic. While this is true also of non-passerines, there are certainly many more visible examples within passerines, and genera are more likely to be split into more widely separated pieces. In an informal poll of bird systematists I conducted at a recent meeting, I asked them to guess what percentage of passerine genera were nonmonophyletic (excluding monotypic genera). The modal answer was “about half”. Of course the majority of genera have not been rigorously investigated so far, which requires sampling most or all the species in the genus plus many species outside it. A few examples should suffice. In addition to finding New World warblers polyphyletic, Lovette and Bermingham (2002) found six out of seven investigated genera with two or more sampled species to be nonmonophyletic; Driskell and Christidis (2003) found three non-monophyletic genera within Meliphagidae out of sixteen examined; Irestedt et al. (2004a) found three of four sampled genera of Thamnophilidae non-monophyletic, and monophyly of the fourth was not confirmed; and Irestedt et al. (2004b) found two of six sampled genera of Dendrocolaptidae non-monophyletic.

1.3.3

Basal Relationships within Passerines

Basal relationships are clear (Fig. 1.3). The family Acanthisittidae (New Zealand wrens), with only two genera and three extant species, is the sister group of all other passerines (Lovette and Bermingham 2000; Barker et al. 2002, 2004; Ericson et al. 2002a; Chesser 2004; Beresford et al. 2005). The remaining passerines are split into two clades, Suboscines and Oscines (Sibley and Ahlquist 1990; Lovette and Bermingham 2000; Barker et al. 2002, 2004; Ericson et al. 2002a; Chesser 2004; Beresford et al. 2005).

& Reproductive Biology and Phylogeny of Birds

Fig. 1.3 Relationships among passerine families and other groups, so far as they can be confidently asserted at present according to my perhaps biased estimate. This tree does not derive from a formal analysis, either of a combined supermatrix or by a rigorous supertree method. Branches in gray represent paraphyletic groups. For Passerida see Fig. 1.4. For reasons behind the topology chosen, see text.

Classification and Phylogeny of Birds

'

Within the suboscines basal relationships are also easy to describe, as there are New World and Old World clades, Tyrannides and Eurylaimides (Sibley and Ahlquist 1990; Irestedt et al. 2001; Barker et al. 2002, 2004; Chesser 2004; Beresford et al. 2005). (Recall that Eurylaimides has a single New World representative, Sapayoa, as mentioned above.) Old World suboscines are also simple: Pittidae (pittas) is either sister to or paraphyletic to Sapayoa, with a larger taxon sample necessary to determine which (Fjeldså et al. 2003; Chesser 2004), and Eurylaimidae (broadbills) is paraphyletic to Philepittidae (asities) (Prum 1993; Irestedt et al. 2001; Barker et al. 2002, 2004; Beresford et al. 2005). New World suboscines can be divided into two clades, Tyranni and Furnarii (Lovette and Bermingham 2000; Irestedt et al. 2001; Barker et al. 2002, 2004; Chesser 2004; note that Sibley and Ahlquist 1990 gave the name Tyranni to a different group), In the oscines, Sibley and Ahlquist (1990) erected two groups Corvida and Passerida. Corvida is however paraphyletic to a (mostly) monophyletic Passerida (Barker et al. 2002). Within the corvidan grade, there are three superfamilies: Menuroidea, Meliphagoidea, and Corvoidea, each very roughly corresponding to Sibley and Ahlquist’s (1990) groupings, plus a number of additional taxa unincluded in any superfamily (Barker et al. 2002, 2004; Beresford et al. 2005). Within Passerida, three superfamilies, Sylvioidea, Muscicapoidea, and Passeroidea, are also roughly similar to Sibley and Ahlquist’s (1990) groups (Sheldon and Gill 1996; Barker et al. 2002, 2004; Ericson and Johansson 2003; Beresford et al. 2005). It has however been necessary to add a fourth, Certhioidea (Barker et al. 2004), and there are also a number of groups that do not fit securely into any superfamily (Sheldon and Gill 1996; Barker et al. 2002, 2004; Ericson and Johansson 2003; Beresford et al. 2005; Fuchs et al. 2006). Ignoring for simplicity groups that do not fit into a superfamily, the relationships among those superfamilies can be resolved: Menuroidea, Meliphagoidea, and Corvoidea are successive sister groups to all other oscines (Barker et al. 2002, 2004; Beresford et al. 2005). Sylvioidea is sister to all other passeridans, and Muscicapoidea is sister to Certhioidea (Sheldon and Gill 1996; Barker et al. 2002, 2004; Beresford et al. 2005). Relationships are described in more detail below.

1.3.4 Relationships within New World Suboscines 1.3.4.1 Furnarii There is a basal polytomy among three clades: Melanopareia (crescent-chests), a genus traditionally assigned to Rhinocryptidae (Irestedt et al. 2002; Chesser 2004), Thamnophilidae (typical antbirds) plus Conopophagidae (gnateaters), and remaining Furnarii (Irestedt et al. 2001, 2002; Chesser 2004; but see Barker et al. 2002, 2004 for a different placement of conopophagids). Formicariidae (ground antbirds) is divided into two parts; one of them, ant-pittas, is the sister group of Rhinocryptidae (tapaculos) while the other, ant-thrushes, is sister to Furnariidae (ovenbirds) and Dendrocolaptidae (woodcreepers). Furnariidae is paraphyletic to Dendrocolaptidae (Irestedt et al. 2002; Chesser 2004).

 Reproductive Biology and Phylogeny of Birds

1.3.4.2

Tyranni

Several clades within Tyranni are clearly established: Pipridae (manakins), Tityridae (tityras), Cotingidae (cotingas), Oxyruncus (sharpbill), and Tyrannidae (tyrant flycatchers). But relationships among them are contradictory in different analyses, and no resolution is strongly supported in any analysis (Barker et al. 2002, 2004; Johansson et al. 2002; Chesser 2004). Relationships within Tyrannidae are likewise contentious, and no molecular study has so far accumulated a large taxon sample.

1.3.5 Relationships within the Corvidan Grade 1.3.5.1 Menuroidea and other near-basal groups Sibley and Ahlquist’s (1990) Menuroidea included Menuridae (lyrebirds), Atrichornithidae (scrub-birds), Climacteridae (Australasian treecreepers), and Ptilonorhynchidae (bowerbirds). More recent studies show that the latter two are sister taxa and are more closely related to other oscines than to Menuridae (Barker et al. 2002, 2004; Ericson et al. 2002b; Beresford et al. 2005; but see Ericson et al. 2002a). There is so far no sequence data available for Atrichornithidae, and I have retained it within a reduced Menuroidea (Sibley and Ahlquist 1990). The unnamed group including Climacteridae and Ptilonorhynchidae is sister to all oscines other than Menuroidea (Barker et al. 2002, 2004; Ericson et al. 2002b; Beresford et al. 2005; but see Ericson et al. 2002a).

1.3.5.2

Meliphagoidea and other groups outside Corvoidea

Meliphagoidea consists of Maluridae (fairy wrens), Meliphagidae (Honeyeaters), Pardalotidae (pardalotes), Acanthizidae (scrubwrens, thornbills), and Dasyornis (bristleheads) (Cracraft and Feinstein 2000; Barker et al. 2002, 2004) and is sister to the remaining oscines (Barker et al. 2002, 2004; Beresford et al. 2005). Relationships within Meliphagidae and Acanthizidae have been investigated by Driskell and Christidis (2004). Two more families, Pomatostomidae (Australian babblers) and Orthonychidae (logrunners) are successively more closely related to Corvoidea and Passerida (Barker et al. 2002, 2004). Three more clades form a polytomy with Corvoidea and Passerida: Callaeatidae (New Zealand wattlebirds), Cnemophilinae (traditionally supposed to be birds of paradise, belonging to the corvoid Paradiseidae), and Melanocharitidae (most berrypeckers—one genus, Paramythia, is corvoid) (Cracraft and Feinstein 2000; Barker et al. 2002, 2004; Beresford et al. 2005).

1.3.5.3

Corvoidea

This large clade contains most of the member of Sibley and Ahlquist’s (1990) Corvoidea (Barker et al. 2004, 2004; Beresford et al. 2005). It consists of two large, resolved subclades, a few pairs of taxa, and a large, basal polytomy. One large subclade is a collection of mostly shrike-like birds including Cracticidae (butcherbirds), Gymnorhina (Australian magpie), Strepera (currawongs), Artamidae (wood-swallows), Aegithinidae (ioras), Malaconotinae (bush-

Classification and Phylogeny of Birds



shrikes), Dryoscopus (puffbacks), Batis (batises), Lanioturdus (chatshrike), Vangidae (vangas), and Prionopidae (helmet shrikes) (Barker et al. 2002, 2004; Beresford et al. 2005; Fuchs et al. 2006). The other group consists of taxa close to Corvidae, including Laniidae (shrikes), Paradiseidae (birds of paradise), Monarchidae (monarch flycatchers), Struthidea (apostlebird), Corcorax (whitewinged chough), Grallina (mud-nest builders), Melampitta (melampittas), Dicruridae (drongos), and Rhipiduridae (fantails) (Barker 2002, 2004; Beresford et al. 2005).

1.3.6

Relationships within Passerida

These are shown in Fig. 1.4.

1.3.6.1

Groups not within any superfamily

Picathartidae (rockfowl) and Petroicidae (Australian robins) are basal members of Passerida (Barker et al. 2002, 2004; Beresford et al. 2005). Regulidae (kinglets) and Hyliota are closer to other Passerida than the previous two families, but cannot so far be linked to any of the superfamilies (Barker et al. 2002, 2004; Ericson and Johansson 2003; Beresford et al. 2005; Fuchs et al. 2006).

1.3.6.2

Sylvioidea

The most difficult problem in establishing sylvioid monophyly has been the inclusion of Paridae (titmice) and its relatives within the superfamily. Though no study has supported that conclusion strongly, several have supported it and none have strongly contradicted it (Sheldon and Gill 1996; Barker et al. 2002, 2004: Beresford et al. 2005; Alström et al. 2006). In addition to its traditional allies, Remizidae (penduline tits), the clade around Paridae has recently been shown to include a clade of refugees from other families, termed Stenostiridae by Beresford et al. (2005). This clade includes Stenostira (fairy warbler, previously considered a sylviid), Culicicapa (canary-flycatchers, usually considered muscicapid, but placed among petroicids by Sibley and Ahlquist 1990), and Elminia (blue-flycatchers, previously considered monarchids) (Barker et al. 2002, 2004; Beresford et al. 2005; Fuchs et al. 2006). Monophyly of the remainder of Sylvioidea is strongly supported, though this group differs from that of Sibley and Ahlquist (1990) by inclusion of Alaudidae (larks) (Sheldon and Gill 1996; Barker et al. 2002, 2004; Ericson and Johansson 2003; Alström et al. 2006; Fuchs et al. 2006). Alaudidae, with its unexpected sister taxon Panurus (bearded tit) (Ericson and Johansson 2003; Alström et al. 2006; Fuchs et al. 2006) is strongly supported as the sister group of all remaining sylvioids (Sheldon and Gill 1996, Barker et al. 2002, 2004; Ericson and Johansson 2003; Beresford et al. 2005; Alström et al. 2006; Fuchs et al. 2006). In the rest of Sylvioidea there is a large polytomy consisting of several genera, a few families, and one large group of families. There are still a great many sylvioid (or supposed sylvioid) genera and species yet to be sampled;

Reproductive Biology and Phylogeny of Birds

Fig. 1.4 Relationships among passerine families, continued. Passerida only. For reasons behind the topology chosen, see text.

Classification and Phylogeny of Birds

!

more sampling of taxa and genes may resolve this polytomy and illuminate unknown clades. Beresford (2005) proposed a “Sphenoeacus group” consisting of Sphenoeacus, Sylvietta, Achaetops, Bradypterus victorini (the genus being polyphyletic), and Macrosphenus; but this was supported solely by Bayesian analysis. Other studies, with only partly overlapping taxon samples, have produced contradictory results (Alström et al. 2006; Fuchs et al. 2006). Alström et al. (2006) found Macrosphenus to belong to Pycnonotidae (bulbuls) with strong support; however, the two studies sequenced different species, and it’s quite possible that both are right and the genus is polyphyletic. The sylvioid polytomy includes four large clades (Alström et al. 2006). Two are the families Cisticolidae (cisticolas) and Timaliidae (babblers). Note that Timaliidae as used here includes the genera Sylvia and Zosterops; there are no families Sylviidae or Zosteropidae, following the terminology of Alström et al. (in press). A third clade includes the families Megaluridae (grassbirds) and Acrocephalidae (acrocephaline warblers) plus the monotypic Donacobius, once considered a troglodytid (Barker 2004; Alström et al. 2006). The final clade consists of a number of families: Hirundinidae (swallows), Pycnonotidae (bulbuls), Phylloscopidae (leaf warblers), Cettiidae (bush warblers), and Aegithalidae (long-tailed tits), plus the genus Hylia (Alström et al. 2006), and the last three form a subclade (Beresford et al. 2005).

1.3.6.3

Certhioidea

This superfamily was erected by Cracraft et al. (2004) to cover a clade of four families removed from Sibley and Ahlquist’s (1990) Sylvioidea. It includes Troglodytidae (wrens), Polioptilidae (gnatcatchers), Certhiidae (treecreepers), and Sittidae (nuthatches) (Barker et al. 2002, 2004; Ericson and Johansson 2003; Beresford et al. 2005). Within this clade, Sittidae is basal and Troglodytidae and Polioptilidae are sister taxa (Sibley and Ahlquist 1990; Harshman 1994; Sheldon and Gill 1996; Ericson and Johansson 2003; Barker 2004; Alström et al. 2006; but see Barker et al. 2002, 2004).

1.3.6.4

Muscicapoidea

The most difficult question is whether Bombycillidae (waxwings, silky flycatchers, palmchat) belongs to this superfamily. No analysis has been conclusive, but several have given weak support (Sibley and Ahlquist 1990; Barker et al. 2002, 2004; Cibois and Cracraft 2004) and I consider the combination to offer strong support. Monophyly of the rest of Muscicapoidea is clear (Sibley and Ahlquist 1990; Harshman 1994; Barker et al. 2002, 2004; Ericson and Johansson 2003; Cibois and Cracraft 2004; Voelker and Spellman 2004; Alström et al. 2006). Relationships among the five families Sturnidae (starlings), Mimidae (mimic thrushes) Cinclidae (dippers), Turdidae (thrushes), and Muscicapidae (sensu stricto—Old World flycatchers) are also contentious, but a strong case can be made for the topology I have chosen, in which Sturnidae and Mimidae are sisters, Turdidae and Muscicapidae are sisters, and Cinclidae is sister to

" Reproductive Biology and Phylogeny of Birds the turdid-muscicapid clade (Sibley and Ahlquist 1990; Barker et al. 2002, 2004; Cibois and Cracraft 2004; Beresford et al. 2005; Fuchs et al. 2006). An alternative arrangement in which Cinclidae is sister to the sturnid-mimid clade is supported only by Bayesian analysis, though the relationship between Turdidae and Muscicapidae is also supported by parsimony jackknifing (Ericson and Johansson 2003). Voelker and Spellman (2004) show a substantially different topology, in which Muscicapidae is basal, but contradictory nodes again have weak support. Two additional genera form a polytomy with Sturnidae and Mimidae (Cibois and Cracraft 2004). Buphagus (oxpeckers) is traditionally considered part of Sturnidae. Rhabdornis is a genus of previously uncertain relationships, sometimes placed in Certhiidae or Timaliidae (Sibley and Monroe 1990).

1.3.6.5

Passeroidea

It may be that Promeropidae (sugarbirds) is the basal family in Passeroidea, but that conclusion is supported by Bayesian analyses alone, and only by different versions of one data set (Barker et al. 2002, 2004; Beresford et al. 2005). What may be partial confirmation is offered by Fuchs et al. (2006). The two genera Modulatrix (spot-throat) and Arcanator (dapple-throat, sometimes merged into Modulatrix) are strongly supported as the sister group of Promerops (Barker et al. 2002, 2004; Beresford et al. 2005). Fuchs et al. (2006), again supported only by Bayesian analysis, show Modulatrix and Arcanator as the sister group of Passeroidea, but Promerops appears in a polytomy at the base of Passerida, and in light of the ambiguity of this information, that is where I have left the entire family. However, there does seem to be good evidence for inclusion of those two species, whose relationships have previously been highly uncertain (Sibley and Monroe 1990), as sister taxa within Promeropidae. The remainder of Passeroidea is clearly monophyletic (Barker et al. 2002, 2004; Ericson and Johansson 2003; Beresford et al. 2005). Nectariniidae (sunbirds) and Dicaeidae (flowerpeckers) are sisters (Barker et al. 2002, 2004; Ericson and Johansson 2003; Beresford et al. 2005), and form a trichotomy with Irenidae (fairy bluebirds) and remaining passeroids (Barker et al. 2002, 2004; Beresford et al. 2005). Peucedramus (olive warbler), previously thought to be either a parulid or basal within 9-primaried oscines (Sibley and Monroe 1990) is instead sister to Prunellidae (accentors) (Ericson and Johansson 2003), and these are sister to the remaining passeroids (Barker et al. 2002, 2004; Ericson and Johansson 2003; Beresford et al. 2005). Passeridae (Old World sparrows) and Motacillidae (pipits and wagtails) are successive sister groups to the nine-primaried oscines (Barker et al. 2002, 2004; Ericson and Johansson 2003). Though this is weakly supported in each analysis, agreement between analyses and a three-codon insertion in the c-myc sequences (Ericson et al. 2000) are conclusive. The nine-primaried oscines form a strongly supported group (Klicka et al. 2000; Ericson and Johansson 2003; Barker et al. 2004), including the families

Classification and Phylogeny of Birds

#

Fringillidae (finches), Emberizidae (buntings and New World sparrows), Icteridae (New World blackbirds), Parulidae (New World warblers), Thraupidae (tanagers), and Cardinalidae (grosbeaks). Fringillidae is the sister of the others (Klicka et al. 2000; Yuri and Mindell 2002; Ericson and Johansson 2003; Barker 2004). Two genera previously considered emberizids, Calcarius (longspurs) and Plectrophenax (snow buntings) are sister taxa of each other and together the sister of the remaining nine-primaried oscines (Klicka et al. 2000; Yuri and Mindell 2002; Ericson and Johansson 2003). Relationships among the remaining five families are contentious. Of analyses with representatives of all five families, most unite Cardinalidae and Thraupidae (Bledsoe 1988; Barker et al. 2002, 2004; Yuri and Mindell 2002), though two (Sibley and Ahlquist 1990; Harshman 1994; Klicka et al. 2000) do not. Again, most unite Icteridae and Parulidae (Bledsoe 1988; Barker et al. 2002, 2004), though two (Klicka et al. 2000; Yuri and Mindell 2002) do not. But support for the minority arrangements are weak. The position of Emberizidae is unclear; Barker et al. (2002, 2004) have strong support for relationship to the cardinalidthraupid clade; Bledsoe (1988) found a relationship to the parulid-icterid clade; Yuri and Mindell (2002) and Klicka et al. (2000), though failing to find the parulid-icterid clade, did find a clade consisting of Emberizidae, Icteridae, and Parulidae. For the present, relationships of Emberizidae are best treated as unresolved.

1.4

CONCLUSION

Progress in avian systematics during the past few years has finally, after a long period of frustration, become worthy of optimism. As more and more homologous DNA sequences and other genetic characters become available for more and more species, We will gradually chip away at the remaining uncertain nodes of the tree, and this trend will only accelerate as DNA sequencing becomes both cheaper and more reliable. These are heady times in avian systematics.

1.5

ACKNOWLEDGMENTS

I would like to thank Keith Barker and Fred Sheldon for their comments on the manuscript, and the other members of the Early Bird project, particularly Kathy Miglia, for sending me copies of many of the referenced articles.

1.6

LITERATURE CITED

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n n

CHAPTER

2

Anatomy of the Testis and Male Reproductive Tract Tom A. Aire

2.1

INTRODUCTION

The testes of birds are intra-abdominal, and, unlike in most mammals, they do not migrate from their site of embryological origin. They are, thus, closely related, topographically, to the kidneys. As in mammals, birds have two testes, one on either side of the midline, bordering the aorta and caudal vena cava, laterally. They are attached from their dorso-medial borders to the dorsal abdominal wall by a short mesorchium, and as the kidneys with which they are related embryologically, they are largely retroperitoneal, and ventral to the vertebral column. Topographically, the cranial poles (extremitates craniales) of the testes lie close to the ventral border of the lungs, while their caudal poles (extremitates caudales) lie cranio-ventral to the cranial divisions of the kidneys (Nickel et al. 1977). The dorsomedial aspect of the testis attaches to the relatively small epididymis. The ductus deferens runs distally from the caudal border of the epididymis, toward the cloaca, into which it opens. There are no known accessory sex organs or glands in birds that are either homologous or analogous to those found in mammals. Mammalian terminologies have often been erroneously applied to certain structural modifications in birds, such as seminal vesicle in passerine birds, and the ampulla of the ductus deferens. These have to be understood for what they are, structurally and functionally, as segmental convolutions or enlargements, respectively, of the ductus deferens. The testis is surrounded by the cranial and caudal thoracic as well as abdominal air sacs, but contrary to the opinion of Cowles and Nordstrom (1946) that this relationship helps to cool the testes, as the mammalian scrotum does, Williams (1958) has observed no differences in the temperature Department of Anatomy and Physiology, Faculty of Veterinary Science, University of Pretoria, Onderstepoort, Republic of South Africa. E-mail: [email protected]

!& Reproductive Biology and Phylogeny of Birds in the area of the viscera and at the testicular surface in the domestic fowl. Herrin et al. (1960) have also disproved the assertion by Cowles and Nordstrom (1946), that adjacent air sacs cool the testis. Testicular color is nearly white in sexually mature and active birds, but it may be grey or black, or a mixture of black and white patches or may be entirely greyish black due to the presence of melanin pigments in melanoblasts, in the testicular connective tissues. The testes of seasonally resting birds are very small, being functionally atrophic. Organs of sexually immature or resting birds have yellowish white to black or grey colour, being quite black in regressed testes that already contain melanoblasts, as a result of the concentration of these pigment cells in the connective tissues of much smaller, regressed organs.

2.1.1

Testis Shape, Asymmetry and Size

The avian testes are usually bean- or oval-shaped (Fig. 2.1), but they are normally vermiform in Cypseloides spp., swifts (Marshall 1961). Although both left and right testes are symmetrically situated on either side of the median plane, they are, however, often dissimilar in size. The left testis is often larger than the right one in most species of birds. Riddle (1918, 1925) observed that the left testes were larger than the right in pigeons (Columba species), but in 36 of 39 in the dove (Streptopelia risorsia), the right was larger than the left. In rooster (Gallus domesticus), the left testis is larger than the right in 57% (Mimura 1928) and 65.3% (Marvan 1969) of the birds studied. The right was larger in only 26.5% while both testes were equal in 8.2% of the birds (Marvan 1969). In 169 genera of birds he studied, Friedmann (1927) concluded that the right testis was not larger than the left in any genus, but in 104 genera, both testes were of the same size and the left was larger than the right in 60 genera. Five genera were inconstant, varying between the first two positions. The left organ is, also, larger than the right organ in the ratite, Emu (Dromaius novaehollandiae), although each gram of testis tissue contributes equally to the production of androgen and, possibly, also, spermatozoa (Malecki et al., 1998). It seems that the right testis diminishes in size with age, as the left testis was larger than the right in 90.9% of day-old Turkey toms (Meleagris gallopavo), 94.3% in toms 53-55 wk old and 100% in toms 63-67 wk of age. Care is needed in removing and identifying each testis especially when the carcass of the bird is placed on dorsal recumbency, with the left side of the specimen being on the right of the operator. That is probably why Law and Kosin (1958) found that all Turkey toms they investigated had larger right testes than the left, whereas Burke (1973) reported that the left testis surpassed the right testis in size, with increasing age in their own Turkey toms. The reason for the generally observed difference in testis size, in favor of the left, is not clearly understood, especially given the variations that have been reported by different authors. However, the embryological study by Witschi (1935) shows that the cortex of the right indifferent gonad loses its chemotactic attraction for primordial germ cells, in favor of the cortex of the left gonad, from the beginning of the 3rd day of embryogenesis. Nevertheless, there are no

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Anatomy of the Testis and Male Reproductive Tract

Fig. 2.1 The topography of the reproductive organs of the quail (Coturnix japonica), from a ventral view. T, testis; E, epididymis; D, ductus deferens; P, pars recta ductus deferentis; R, receptacle of the ductus deferens; L, lung; K, kidney. Original.

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" Reproductive Biology and Phylogeny of Birds differences in sizes between the left and right testes in the Marsh hawk (Circus cyaneus) (Witschi 1935; Stanley and Witschi 1940). The size of the avian testis, except that of several breeds of Domestic Fowl and Turkey developed for agricultural production, varies considerably between phases of the reproductive cycle. Thus, the testis reaches maximum size for the species or breed during the peak of the breeding season, and becomes considerably smaller, following regression, during the sexually inactive or resting period of the reproductive cycle, in most wild birds. This variation in size may be as much as 400- to 500-fold in seasonally breeding species (Lofts and Murton 1973). During the active breeding season, testis size is positively related to sperm production rate (Møller 1991), as has been observed in mammals (Møller 1989). Testis sizes and body weights of 247 species of birds, from 152 genera, 37 families, and 16 orders, have been compiled by Møller (1991). Testis size in birds is further discussed in Chapter 9.

2.1.2

Body Temperature and Testis Function in Birds

Whereas in most mammals the testes migrate from the site of their embryogenesis distally and ventrally into the scrotum, those of birds retain their position of origin, close to the kidneys. Mammalian male germ cells degenerate at the body’s internal temperature of 37∞C, and even slight and temporary elevations of scrotal temperature may cause male infertility (Plöen 1972, 1973a,b). On the contrary, spermatogenesis occurs normally at core body temperature (of 40-41∞C) in birds (Béaupre et al. 1997). Mezquita et al. (1998) recently demonstrated a number of apparently genetically controlled biochemical effects that may contribute to ensure the synthesis of essential proteins which achieve thermotolerance during avian, but not mammalian, spermatogenesis.

2.1.3

Consistency and Structure of the Testis

The testis of sexually active birds is soft to touch and quite fragile compared to the mammalian testis. The main exception is the Ostrich (Struthio camelus), whose testicular substance is enclosed in a firm testicular capsule which is relatively very thick (vide infra); in other investigated birds it is thin. An incision through the testicular capsule into the testis causes a mild protrusion of the testicular substance from which there is a dripping of a relatively copious amount of milky fluid, even in the Emu whose testis is very dark in color. Sexually active avian testes have a considerable amount of fluid content (Lake 1957; Aire 1979a; Clulow and Jones 1988).

2.1.4

The Excurrent Ducts of the Testis

The avian epididymis is a small spindle-shaped structure that is attached intimately to the medial border of the testis (Fig. 2.1). A thin cranial extension of this structure is incorporated into the adrenal gland capsule, as the appendix paradidymidis (Gray 1937; Budras and Sauer 1975a; Budras and Meier 1981).

Anatomy of the Testis and Male Reproductive Tract

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The caudal end of the epididymis also thins out, and is continued by the ductus deferens. The epididymis of the rooster is 3-5 mm thick at its base, by which it is attached to the testis (Hodges 1974; Pers. obs.). The epididymis is relatively large in the ostrich, being between 21 and 31 g in weight, 12 and 15.4 cm in length and 3-7 cm in height, at its highest point at the caudal onethird of the organ (Pers. obs.). It is about 2.4 cm wide (Soley 1992). The epididymis, as the testis, is also firm to touch in the Ostrich. In all birds, the ductus deferens leaves the caudal end of the epididymis in a slightly wavy manner, but becomes considerably convoluted, increases in diameter, cranio-caudally, and is situated lateral to the relatively firm and regular ureter. A short segment of the duct straightens out to form the pars recta ductus deferentis, just before it enters the cloaca. The ductus deferens terminates thereafter, in a spindle- or barrel-shaped enlargement, the receptaculum ductus deferentis, which is embedded in the cloacal wall, but opens into the urodeum through its protruding and pointed distal end, the papilla ductus deferentis. Birds do not have accessory sex organs that are to be found in mammals.

2.1.5

Blood Supply to the Reproductive Organs

The blood supply to the male reproductive organs of birds has been studied fully only in the rooster (Nishida 1964) and Pigeon (Bhaduri et al.1957), and partly by Siller and Hindle (1969) and Kurihara and Yasuda (1973) in their reports of the vasculature of the kidney in the rooster. Blood supply to the male organs in the ostrich has also been studied grossly to complement these reports (Elias M, Aire, T. A. and Soley, J. T. unpublished results), and the pattern of arterial supply as well as venous drainage of the male organs is generally similar in these three species of birds. The following account is based on these reports. The avian male reproductive organs are intra-abdominal and retain their embryological positions, and therefore their blood supply is relatively simpler than in mammals. A short testicular artery (Arteria testicularis) arises from the A. cranialis renalis, and runs into the testis, along its dorso-medial border, supplying the testis, and sending thin branches to the epididymis (Figs. 2.2, 2.3 and 2.4). One or two short accessory testicular arteries (Aa. testiculares accessoria) may originate directly from the aorta, unilaterally or bilaterally, and similarly run into the hilus of the testis. The testicular arteries run short courses into the hilus of the testis from where they divide into intratesticular loops (Fig. 2.3). Branches ramify in the interstitial tissue, from these loops. The cranial part of the ductus deferens is supplied by the R. ureterodeferentialis cranialis that arises from the A. cranialis renalis. The A. renalis communis originates from A. sciatica in the Ostrich (Elias et al., unpublished observations) and immediately divides into the A. renalis medius and A. renalis caudalis. The former sends one or two twigs to the middle part of the ductus deferens. The A. pudenda sends several branches (Rr. ureterodeferentiales caudales) to the caudal part of the ductus deferens, and several other branches (Rr. cloacales) into the cloaca and root of the phallus. The general disposition of the blood

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Reproductive Biology and Phylogeny of Birds

Fig. 2.2 The arterial supply to the reproductive organs of male Gallus domesticus. Ara, A. renalis cranialis; At, A. testicularis; Ae, Aa. epididymicae; Auda, Aa. ureterodeferentiales craniales; Audm, Aa. uretero-deferentiales mediae; Audp, Aa. uretero-deferentiales caudales; Apc, A. pudenda communis. From Nishida, T. 1964 Japanese Journal of Veterinary Science 26: 211-221. Figure 1. Reproduced with the permission of Japanese Society of Veterinary Medical Science.

Anatomy of the Testis and Male Reproductive Tract

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Fig. 2.3

Fig. 2.4 Figs. 2.3 and 2.4 A semi-diagrammatic illustration of blood vascular supply in the right testis of Gallus domesticus. Aoa, aorta; At, A. testicularis; Ata, A. testicularis accesorius; Vic, V. iliaca communis; Vcp, V. cava caudalis. From Nishida, T. (1964) Japanese Journal of Veterinary Science 26: 211-221. Figure 4. Reproduced with the permission of Japanese Society of Veterinary Medical Science.

"" Reproductive Biology and Phylogeny of Birds vessels in the rooster, pigeon and ostrich are similar, with only minor intraand inter-species variations. The veins draining the testicular substance are tributaries of larger veins which run peripherally into the testicular capsule (Figs. 2.3 and 2.4). The latter are large, wide and clearly visible as they run radially toward the hilus from which they emerge and open into the caudal vena cava as numerous short veins (Nishida 1964). The veins draining the ductus deferens are mostly satellites of the arteries that supply the duct (Nishida 1964).

2.2 2.2.1

THE TESTIS The Testicular Capsule

The avian testicular capsule is generally very thin (Lake 1971). In a preliminary study, figures for testicular capsule thickness in certain species of birds are, on the average, 578.1 mm in the ostrich, 81.5 mm in the rooster, 91.7 mm in the Japanese quail (Coturnix japonica) and 91.8 mm in the drake (Anas platyrhynchos) (Pers. obs.). However, at the interface between the testis and epididymis, the capsule is much thicker, and has been found to be, on the average, 1215.9 mm thick in the ostrich, 515.0 mm in the rooster, 255.4 mm in the Japanese quail, and 233.7 mm in the drake. There are no readily available data on testicular thickness in mammals, but Davis et al. (1970) indicate that the thickness of the testicular capsule varies from one testicular region to another. The tunica albuginea in man is approximately 1000 mm thick, at 75 years of age (Yoshimura and Fukunishi 1965, cited by Davis et al. 1970). The testicular capsule in the bird divides in the epididymal region, and sends a thick branch around the epididymis. The other branch, devoid of the tunica serosa, runs on the orchido-epididymal border, and is pierced at a number of points, in most birds, by the intracapsular portion of the rete testis (RT) ductules, to enter the epididymis. Histologically, the testicular capsule is composed of three main tissue layers: an outer, thin tunica serosa, a thick tunica albuginea and the innermost, very thin tunica vasculosa (Fig. 2.5). A basement membrane separates the thin mesothelial cells of the t. serosa from the bulky connective tissue of the t. albuginea. The latter forms the bulk of the capsule, and is composed of collagen, elastic fibers and abundant fibroblasts (Hodges 1974). Blood vessels, especially veins of varying sizes run within these bundles of fibroblasts and collagen, and are easily observed running radially on the surface of the testis, grossly. Smooth muscle actin intermediate filaments have been demonstrated, immunohistochemically, in the testicular capsule of the Japanese quail, Drake, Turkey, rooster and Ostrich (Aire, T.A. unpublished observations). This indicates that there are smooth muscles in the testicular capsule of birds, as has been demonstrated in some mammals (Davis et al. 1970; Hargrove et al. 1977). Smooth muscle cells in the testis capsule are probably responsible for the spontaneous and drug-induced contractions of the capsule of mammals. These contractions may assist in moving immobile testicular spermatozoa

Anatomy of the Testis and Male Reproductive Tract

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Fig. 2.5 Histological section of testicular capsule of the ostrich (A, C) and drake (B). A. The capsule is thick and shows the tunica serosa (arrowheads), tunica adventitia (T) and tunica vasculosa (arrow). V, large intracapsular blood vessels; S, seminiferous epithelium. B. A thinner capsule, displaying the three tissue layers, as in A. Nu, an elongated nucleus of a myofibroblasts. C. testicular capsule (T) of a juvenile ostrich. There are no obvious septa originating from the testicular capsule, rather, connective tissue containing oval, euchromatic nuclei aggregate and pass between the seminiferous cords (Sc) of the testis. Bars: All figures = 100 µm. Original.

"$ Reproductive Biology and Phylogeny of Birds toward the epididymis (Davis et al. 1970). A similar function may be expected to occur in birds. There are no septa running from the testicular capsule into the testicular substance, as occurs in most mammals (Fig. 2.5A, C). Loose connective tissue, which appears better formed in the ostrich than in other birds, may be seen conducting blood vessels from the subcapsular region into the testis substance, and sending slivers of connective tissue between the seminiferous tubules (Fig. 2.5A, C).

2.2.2 Seminiferous Tubules The seminiferous tubules of birds are dissimilar to those of mammals by forming highly and complexly anastomotic, non-blind-ending network of tubules (Huber 1916; Bailey 1953; Lake 1957; Marvan 1969). Evidence of this structure is commonly found in histological sections (Fig. 2.6), and is probably responsible for the lack of connective tissue septa, as well as the nonlobulation of the avian testis. The seminiferous epithelium is several cells thick, and contains two cell types: fixed, somatic cells, represented by the Sertoli cells, and temporary and mobile germ cells, comprising a series of differentiating cells (spermatogonia, primary and secondary spermatocytes and spermatids) (Fig. 2.6 B, inset). The basal stem cell is the spermatogonium that divides repeatedly to form larger, upward-mobile, cells in successive stages of meiotic division and maturity. Spermatogenesis in birds, as in mammals, involves a series of divisions of spermatogonia, resulting in primary spermatocytes and secondary spermatocytes, both of which undergo meiotic divisions, culminating in the evolution of spermatids. The latter differentiate to form motile, itinerant spermatozoa. Spermatogenesis is discussed fully in Chapter 7, but the Sertoli cell is discussed below.

2.2.3

Sertoli Cells

The Sertoli (sustentacular) cell is the fixed or permanent and non-germinal cell in the epithelium of the seminiferous tubule. It is known as a “nurse” cell because it supports and provides nutrition for the developing germ cells. Avian Sertoli cells are generally similar to those described for the mammal (Cooksey and Rothwell 1973). Sertoli cells are tall, columnar and extend from the basal lamina to the luminal border of the seminiferous epithelium (Fig. 2.7A, B). Sertoli cells in the rooster have fewer broad base contacts with the basement membrane than in mammals (Osman et al. 1980). A similar observation has been made in the Japanese quail (Fig. 2.7A, B). The cytoplasm of this cell is generally more electron-dense than that of attaching or adjacent germ cells (Aire, T. A. personal observations), and arborizes between the germ cells that attach to, or embed in, it in an irregular manner. The nucleus lies at the level immediately above the spermatogonia, and is relatively large and euchromatic. It may therefore be relatively of high activity. It is usually irregular, has a prominent nucleolus, and is situated close to the basal lamina in most birds except in the ostrich where it lies in the middle part of the cell

Anatomy of the Testis and Male Reproductive Tract

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Fig. 2.6 Coturnix japonica. A. A survey photomicrograph of the seminiferous tubules showing complex branching of the seminiferous tubules. B. Higher power view of seminiferous tubules displaying various germ cells in the epithelium. Inset: shows Spermatogonia (arrows); Sertoli cell nucleus (arrowhead); round spermatids (Sr); elongated spermatids (Sel). Bars: A = 100 µm, B, including inset = 200 µm. Original.

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Fig. 2.7 Coturnix japonica. A. A resin section of seminiferous tubules stained with toluidine blue, showing positions of concentrated Sertoli cell cytoplasm (arrowheads). Dark particles and elongated profiles in the Sertoli cell cytoplasm are sections of embedded elongated spermatids. B. A TEM micrograph of the basal half of the seminiferous epithelium showing a Sertoli cell (S) with its characteristic elongated and irregular nucleus (N), electron-dense cytoplasm (stars), spermatogonium (Sg) and spermatocyte (Sp). T, peritubular and interstitial tissue. Bar: A = 200 µm, B = 2 µm. Original.

Anatomy of the Testis and Male Reproductive Tract

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and hence epithelium (Aire, T. A. personal observations). The cytoplasm contains abundant, small, smooth endoplasmic reticulum (SER), numerous free ribosomes and polyribosomes, a well-developed Golgi complex and micropinocytotic vesicles, both basally and between adjacent cells (Fig. 2.8). Microtubules are conspicuous in the cell body and processes that surround the germ cells. Lipid droplets are dense and small in size, and are few in the rooster (Cooksey and Rothwell 1973) unlike in the Budgerigar (Melopsittacus undulatus) (Humphreys 1975), where they are numerous. Sertoli cells are absolutely necessary for the development of germ cells, from spermatogonia to spermatozoa, not only because they provide physical support and nutrition for the germ cells, but also because they provide the avenue for substances to pass between the blood and germ cells. In mammals, there is a close temporal relationship between the development of the secretion of many Sertoli cell products and the appearance and increase in number of primary spermatocytes and early spermatids (Jégou 1991). The same is probably true in birds because Jégou (1991) considers that the interaction between late spermatids and the Sertoli cell has survived throughout evolution and therefore constitutes a major aspect of the paracrine regulation of spermatogenesis. The passage of substances from Sertoli cells to germ cells is regulated by a peculiar structural feature of Sertoli cells (Mann and LutwakMann 1981), which involves cell-cell contacts in the seminiferous epithelium. These contacts are necessary because there is an intimate interdependence of germ cells and Sertoli cells. Elongating and elongated spermatids are difficult to isolate from Sertoli cells without their being considerably damaged. Besides, early spermatids have not survived well in cell culture (Jutte et al. 1981; Le Magueresse and Jégou 1988). Indeed, Jégou (1991) emphasises that spermatids are major regulators of Sertoli cell function. Details of the cellular contacts between the Sertoli and germ cells have been reported in mammals (Fawcett 1973; Dym 1973; Dym and Fawcett 1970; Fawcett 1975) and birds (Cooksey and Rothwell 1973; Osman et al. 1980; Bergmann and Schindelmeiser 1987; Sprando and Russel 1987; Pelletier 1990; Pfeiffer and Vogl 1993). The main cell contacts in the seminiferous epithelium of the rooster (Osman et al. 1980; Bergmann and Schindelmeiser 1987; Pelletier 1990) are similar to those described in mammals, and are classified as Sertoligerm cell junctions and Sertoli-Sertoli cell junctions (Fig. 2.9). Sertoli-germ cell contacts consist of gap and adhering junctions that have subsurface condensations of 7 nm filaments that are lined internally by cisternae of endoplasmic reticulum (Pelletier 1990). Where two germ cells are apposed, contacts are made by means of gap junctions that are similar to those of Sertoli-germ cell adhesions (Fig. 2.9). The Sertoli-Sertoli cell junctions comprise occluding, gap and adhering junctions, with the occluding junction being most basal, but situated above spermatogonia as well as early primary spermatocytes (Pelletier 1990). The gap junction separates the occluding junction from the adhering junction. The inter-Sertoli cell-occluding junction is the morphological basis of the intra-testicular and extremely effective blood-

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Fig. 2.8 Coturnix japonica. High power view of the cytoplasm of the Sertoli cell of the quail exhibiting an abundance of smooth endoplasmic reticulum (SER), a few lipid droplets (L), some mitochondria (M), Golgi complex (G), a few microtubules (upper arrowheads), and a lysosome (Y). Sp, spermatogonium; RER (lower arrowheads) B, basal lamina. Bar = 1 µm. Original.

Anatomy of the Testis and Male Reproductive Tract

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testis barrier. The avian occluding junction is different from that of mammals in only a few details, such as the absence, in birds, of subsurface filaments, disposed of as bundles, that lie between the adjacent tight junctions and the subsurface cisternae (Fig. 2.9). This structural barrier, as in mammals, separates the seminiferous epithelium into the basal and adluminal compartments, and is able to prevent tracer compounds in the basal compartment from entering the adluminal compartment of the seminiferous epithelium in the rooster (Osman et al. 1980; Bergmann and Schindelmeiser 1987; Pelletier 1990). The occluding Sertoli-Sertoli cell junctions are present and relatively constant in position, above the spermatogonia, in both regressed testes (Pelletier 1990), and active testes (Osman et al. 1980; Bergmann and Schindelmeiser 1987; Pelletier 1990) of birds. Thus, in the birds investigated, the Sertoli cell junction is constantly present and active, irrespective of the phase in the reproductive cycle.

2.2.3.1

Functions of the Sertoli cell

Since avian Sertoli cells are similar to those of mammals, structurally, it is tempting to assume that the main functions in both classes of animals are generally similar. The abundance of SER in the mammalian Sertoli cell indicates a hormone-secreting cell (Fawcett 1975). Sertoli cells are unable to significantly synthesise steroids de novo, per se, but promote interconversion of steroids e.g. progesterone and androstenedione to testosterone and reduced 5 a-androgens (Mann and Lutwak-Mann 1981). Whereas the presence of steroids was demonstrated in extracts of the fowl testis (Delrio et al. 1967), it was Tingari’s (1973) histochemistry study that demonstrated steroidogenic activities in the seminiferous epithelium, and particularly in those parts that are consistent with Sertoli cell locations, in the rooster. Androgen binding protein (ABP), inhibin and activin are endocrine regulators of follicularstimulating hormone (FSH) production and of testicular steroidogenesis and spermatogenesis, not only in mammals but also in birds (Johnson and Brooks 1996; Lovell et al. 2000; Onagbesan et al. 2004). It is likely that the avian Sertoli cell functions as its mammalian counterpart, in secreting ABP, inhibin, activin, and a part of the testicular fluid (Hagenäs et al. 1975) that have the same types of functions, as in mammals. The phagocytic ability of the Sertoli cell is well known in both mammals and birds. These cells have been observed to remove residual bodies in the rooster and Japanese quail (Cooksey and Rothwell 1973; Sprando and Russel 1987; Lin and Jones 1992) and India ink particles (Fig. 2.10) that were introduced into the seminiferous tubular lumen in the rooster (Pers. obs.). Perhaps one of the most important functions of the Sertoli cell is the establishment of the most durable component of the blood-testis barrier, which creates two fluid compartments within the seminiferous epithelium. This barrier, already described above, is similar to the mammalian barrier, in major particulars. It is present and functional in both sexually mature and active as well as regressed testes in birds, by preventing tracers from entering the adluminal compartment, from the basal compartment (Osman et al. 1980;

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Reproductive Biology and Phylogeny of Birds

Fig. 2.9. A. A diagrammatic representation of Sertoli-germ cell and Sertoli-Sertoli cell ectoplasmic specializations and junctions in an active testis. Sert, Sertoli cell; Fig. 2.9 Contd. ...

Anatomy of the Testis and Male Reproductive Tract

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Pelletier 1990). It has an important protective role in safeguarding the germ line from noxious influences originating both from within and external to the individual, “…including both isolation of sperm-specific autoantibodies and autoimmunocytes from the tubule in the event that sensitization does occur” (Neaves 1977). In addition to these functions, there are peritubular-Sertoli cell interactions. Peritubular cells constitute an important extra-tubular part of the blood-testis barrier, and they also have important regulatory interactions with Sertoli cells in the seminiferous tubule, via the production of paracrine factors that enhance Sertoli cell functions, including the production of ABP and transferrin (see Skinner et al. 1991). Sertoli cell-Leydig cell interactions also exist, in which both cells influence each other with regard to steroidogenic activities, as well as inhibin and activin secretions (see Skinner et al. 1991).

2.2.4

Interstitial Tissue of the Testis

The interstitial tissue of the testis consists of two main components. The first is that compact layer of myofibroblasts and connective tissue which closely surrounds the seminiferous tubule, and known as the boundary tissue. The other is the loose connective tissue, the interstitium, which lies between seminiferous tubules, with full expression in the angular areas or wedges between 3 or more adjacent seminiferous tubules (Fig. 2.11). The boundary or peritubular tissue consists of subepithelial tissue and layers of alternating elongated cells and their interconnecting amorphous tissue. The interstitium, on the other hand, consists of single, or groups of a few, Leydig cells (Cellulae interstitiales), blood vessels, fibroblasts and cells of the macrophage system, such as macrophages, lymphocytes and monocytes. The interstitial tissue of Fig. 2.9 Contd. ...

Sg, spermatogonium; Sp, spermatocyte; Sd, elongating spermatids; B, basal lamina; L, lumen of seminiferous tubule. Arrowheads indicate a junctional complex between adjacent Sertoli cells (Sertoli-Sertoli junctional complex) situated above the spermatogonium, and consisting of a varying number of tight junctions and parallel-running subsurface cisternae of endoplasmic reticulum (highlighted in Figure B). Other less frequently encountered inter-Sertoli cell junctions, lying apical to the tight junctions, are adhering junctions (Ad) (comprising intracytoplasmic condensation of material, on both sides of the Sertoli cell plasma membranes, and a line of dense material between the two cell membranes, and gap junctions (Gp). The former is also found between adjoining Sertoli and germ cells. A special type of Sertoli cell-elongating spermatid cell junction (arrows) displays a subsurface condensed material in the Sertoli cell cytoplasm, but none in the spermatid cytoplasm (hence also known as hemi-ectoplasmic specialization). B. A higher power representation of tight junctions between adjacent Sertoli cells. Arrowheads, focal tight junctions; Sert, Sertoli cell; er, cisternae of endoplasmic reticulum. Unlike in mammals, there are no layers of filaments between the cisternae and the cell membranes, on either side. Original.

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Fig. 2.10 Coturnix japonica. India ink particles (arrowheads) occur in the apical half of the Sertoli cytoplasm after 15 m of retrograde infusion of India ink through the rete testis into the seminiferous tubules. Bar = 50 µm. Original.

the testis has been described in a number of mammals (Christensen 1965; Fawcett et al. 1969, 1970; Dym 1973; Fawcett et al. 1973; Weaker 1977; Skinner et al. 1991), but reports on this tissue in birds are rather scanty, and are, in the main, on domestic species of birds, such as the rooster (Rothwell and Tingari 1973, 1974; Rothwell 1975; Aire 1997), Duck (Marchand 1973; Aire 1997), Guineafowl (Numida meleagris) and Japanese quail (Aire 1997). Although a very important component of the interstitium, Leydig cells have been described in only a few species of birds (Connel 1972; Garnier et al. 1973; Marchand 1973; Rothwell 1973; Scheib 1973; Aire 1997). The following review of the testicular interstitial tissue is based largely on these reports, and a number of unpublished observations from our laboratory.

2.2.4.1

The boundary (peritubular) tissue

Figures 2.11A and B show interstitial tissue located between three seminiferous tubules in the drake. Figure 2.11C illustrates the relationship between the seminiferous tubules, the boundary tissue, and the interstitium containing blood vessels and Leydig cells. The boundary tissue in the rooster (Rothwell 1975) and Ostrich (Soley, J. T., van Wilpe, E. and Aire, T. A. unpublished observations) consists of an inner fibrous layer, subjacent to the seminiferous epithelium, and an outer cellular layer of myofibroblasts. The inner fibrous layer or lamella exhibits a homogeneous, moderately dense basal lamina, and an adjacent layer of multi-directional collagen fibrils. The latter layer, of collagen fibrils, is absent in the birds studied by Aire (1997) (see Fig. 2.12A). It is not known why this structural difference occurs between the bird

Anatomy of the Testis and Male Reproductive Tract

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Fig. 2.11 Anas platyrhynchos (A, B), Coturnix japonica (C). A. Wedges of interstitial tissue between three seminiferous tubules shows a peripheral lymphatic vessel (arrows). V, blood vessels; L, Leydig cells. B. The lymphatic vessel (arrow) in this histological section appears to be centrally located in the interstitium. L, Leydig cell. C. An electron micrograph displays a lymphatic vessel (star) situated between the boundary tissue (B) of a seminiferous tubule (S), a blood vessel (V) and a Leydig cell (L). The blood vessel and Leydig cell are within the interstitium. Bars: A = 60 µm; B = 60 µm; C = 2 µm. Original.

#$ Reproductive Biology and Phylogeny of Birds tissues that were fixed by immersion fixation (Rothwell and Tingari 1973; Rothwell 1975; Soley, J. T., van Wilpe, E. and Aire, T. A. unpublished observations) and those fixed by vascular perfusion (Aire 1997). The basal lamina rests on a thin layer of microfibrils and amorphous, moderately electron-dense material in perfused birds (Aire 1997). However, Humphreys (1975) reports that the basement membrane of the budgerigar testis exhibits “irregular collagen content, and that many birds do not show collagen”, at all. Microfilament rearrangement and re-configuration in the Leydig cells of the dog has been described in tissues that were re-fixed by immersion in glutaraldehyde after a previous perfusion fixation (Connel and Christensen 1975). Collagen fibrils have been observed, recently, in unpublished micrographs, below the basal lamina in a patchy manner, in some, but not all, of the seminiferous tubules, in intravascularly perfused adult sexually active Japanese quail, in our laboratory (Fig. 2.12B). This is in accord with the findings by Humphreys, referred to above. Further studies are necessary, using various fixatives and buffers, as well as fixation methods in several species of birds in order to clarify this concern. The mesenchymal layer of myofibroblast cells varies in thickness, based upon the number of concentric, alternating or overlapping cells. There are up to 5, and occasionally, more, cellular lamellae (Aire 1997; Soley, J. T., van Wilpe, E and Aire, T. A. unpublished observations). The myofibroblasts contain highly elongated, uniformly granular nuclei displaying a few foci of marginated chromatin and eccentric nucleoli. Short profiles of moderately distended RER, a few oval mitochondria, a well-developed Golgi complex and a number of micropinocytotic vesicles are found in the cytoplasm, which, also, abounds in typical 5 nm-thick intermediate filaments and associated focal intracytoplasmic densities. In the rooster and ostrich tissues fixed by immersion, bundles of collagen fibres lie in the intercellular spaces between myofibroblast cells (Fig. 2.13). Myofibroblasts that exhibit features which are found in either fibroblasts or smooth muscle cells are not uncommonly seen. In the quail and ostrich, the peritubular cells are invested by what appears to be an incomplete layer of basal lamina (Fig. 3.12B). The endothelium of blood or lymphatic capillary forms the peripheral limit between the boundary tissue and the interstitium (Fig. 2.11C). Birds are similar to the rat, chinchilla, guinea pig and mouse, in possessing a relatively small volume of Leydig cells and in the location of the lymphatic vessels, but differ from these mammals in lacking an extensive peritubular lymphatic system (Fawcett et al. 1973). However, the arrangement of the myofibroblasts is similar to that in human and cat (Burgos et al. 1970), in being multilayered. The peritubular tissue of birds therefore varies in certain particulars between species (Rothwell 1975; Aire 1997; Soley, J. T., van Wilpe, E. and Aire, T. A. unpublished observations), and combines structural features of the interstitium found variably in mammals.

2.2.4.2

The interstitium

The interstitium is relatively compact in birds (Aire 1997) except in the ostrich in which it forms a loose and ‘oedematous’ connective tissue (Soley, J. T., van

Anatomy of the Testis and Male Reproductive Tract

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Fig. 2.12 Anas platyrhynchos (A) and Coturnix japonica (B). A. The basal lamina (B) of the seminiferous tubule does not rest on an internal lamella of collagen fibers, but directly on an amorphous material adjacent to myofibroblast cells (M) of the boundary tissue. Extensions of the basal lamina (arrows) into the seminiferous epithelium occurs frequently. B. The internal (fibroreticular) lamella of the boundary tissue displays a patchy presence of collagen fibers (C) below the basal lamina (B); Arrow indicates a part of the internal lamella devoid of collagen fibers. Arrowheads, dense material, similar to the lamina densa of the basal lamina, interposed between myofibroblasts (M); S, seminiferous epithelium. Bars: A = 2 µm; B = 1 µm. Figure A is from Aire, T. A. 1997 Onderstepoort Journal of Veterinary Research 64: 291-299, with permission of the Editor. Figure B is original.

#& Reproductive Biology and Phylogeny of Birds

Fig. 2.13 Struthio camelus. A. The boundary tissue at the base of the germinal epithelium (GE) displays a regular basal lamina (arrowheads), the inner fibrous lamellum (F) and the outer peritubular layer consisting of alternating cellular (1-4) and acellular lamellae. Collagen fibrils (cf) are the most conspicuous element of the acellular lamellae. B, Dilated profiles of rough endoplasmic reticulum (RER), bundles of microfilaments (arrow), focal densities (squat arrow) and coated pit (arrowhead) are present in the myofibroblasts. Cross-sections of collagen fibers occur in the acellular lamellae. Basal lamina (double arrowhead). Bars for both figures = 1 µm. (From Soley, J. T., van Wilpe, E. and Aire, T. A. unpublished micrographs).

Anatomy of the Testis and Male Reproductive Tract

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Wilpe, E. and Aire, T. A. unpublished observations). The component tissue and cell types include a number of centrally situated blood vessels, varying in size from capillaries to large vessels (Fig. 2.11). Lymphatic vessels are sparse and usually peripherally, but occasionally, centrally, located in the interstitium. When peripheral, they meander between the boundary tissue and the central components of the interstitium (Aire 1997). They are commonly located centrally in the Ostrich (Soley, J. T., van Wilpe, E. and Aire, T. A. unpublished observations). Other component cells include very few macrophages that are closely associated with a few, single or groups of a small number of Leydig cells. It has not been determined in birds if there are structural and functional relationships between these two cell types, as in mammals in which they are morphologically and functionally coupled (Bergh 1985, 1987; Gayton et al. 1994). Other blood-derived cells, such as lymphocytes, plasma and mast cells may occur in the interstitium. The central role of the Leydig cell in the secretion of the primary male sex hormone, and possibly also testicular estrogens in man, the stallion and boar (Akingbemi et al. 1999) sets this cell apart for special treatment in this review.

2.2.4.3

The Leydig cell

Leydig cells of birds (Connel 1972; Nicholls and Graham 1972; Garnier et al. 1973; Marchand 1973; Rothwell 1975; Aire 1997; Soley, J. T., van Wilpe, E. and Aire, T. A. unpublished observations) are generally similar, structurally, to those of mammals (Neaves 1975; Russel 1996; Akingbemi et al. 1999). The number of Leydig cells that are present in the interstitial wedges between multiple seminiferous tubules is small (Fig. 2.11C). Leydig cells seem to form columns of cells in the interstices in the turkey (personal observations). References to information on the ostrich are taken from unpublished results of studies done on the interstitial tissue of this bird by Soley, J. T., van Wilpe, E. and Aire, T. A. Ultrastructurally, Leydig cells of gonadally active birds are large, relatively electron-dense, and contain euchromatic, oval, polygonal or elongated nuclei, depending on location in the interstitial tissue (Fig. 2.14). The Golgi complex is generally moderately developed in birds (Aire 1997) except in the ostrich, in which it is well developed and displays numerous Golgi fields (Fig. 2.15B). Prominent, relatively numerous, oval or elongate mitochondria contain tubular cristae that are embedded in an electron-dense matrix (Fig.2.14). The rough endoplasmic reticulum (RER) is poorly developed, relative to the smooth endoplasmic reticulum (SER) which is more developed (Fig. 2.14), but not nearly as well as in mammals (Neaves 1975; Aire 1997). The SER profiles are short, moderately dilated, or, in the ostrich, may form branched or anastomotic strands. Whorls of SER are present only in the guineafowl (Aire 1997), and, in the ostrich, there are concentric collections of thickened membranes which develop within existing cisternae of SER, resulting in arrays of parallel-oriented, long, rod-shaped structures of greater electron density than normal strands of SER with which they are continuous (Fig. 2.15). Their functions are not known. Scattered ribosomes, polyribosomes,

$ Reproductive Biology and Phylogeny of Birds

Fig. 2.14 Coturnix japonica. Part of an electron micrograph of a Leydig cell showing mitochondria (M) with tubular cristae within a dense matrix. Abundant profiles of smooth endoplasmic reticulum (SER), several lipid droplets (D) and only a few, short profiles of rough endoplasmic reticulum (arrowhead) occur in the cytoplasm. The nucleus (N) is oval in shape. F, microfilaments in peritubular myofibroblasts (P); S, seminiferous tubule. Bar = 1 µm. Original.

Anatomy of the Testis and Male Reproductive Tract

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dense bodies and intermediate filaments of variable prominence and location within the cell are present in the ostrich. Lipid droplets, often partially extracted, are fewer and larger in the drake, guineafowl and rooster, than in the Japanese quail (Fig. 2.14) and ostrich. In several cells, a solitary cilium, of undetermined axoneme structure, originates deeply in the cytoplasm and, therefore, traverses an intracytoplasmic canal to reach the cell surface, in the ostrich. Non-myelinated nerve cell processes, as well as occasional naked axons are to be found throughout the interstitium. These nerves are transmitted into the substance of the testis by septa-like strands of connective tissue that surround the seminiferous tubules. The boundary tissue, however, does not display any neural elements, but some nerve fibers seem to lie close to the basal lamina of the seminiferous tubules in the rooster (Tingari and Lake 1972a). Adrenergic nerves appear to innervate interstitial cells in Mute swan (Cygnus olor) (Baumgarten and Holstein 1968).

2.2.4.4

Functions of the intertubular tissue

Peritubular cells are involved not only in environmental interactions, that is structural interactions, but also regulatory interactions, in which they produce a paracrine or an autocrine agent to elicit a signal transduction event that influences cellular functions on a molecular level (Skinner 1987). Considerable and varying interactions occur between the cells of the testis (Skinner et al. 1991). These include peritubular cell-Sertoli interactions, all of which are vital to the proper structural and functional roles of the various cells in the testis. For example, the peritubular cells secrete PModS which influences Sertoli cell functions that are vital for the maintenance and control of spermatogenesis (Skinner 1987).

2.3

THE EXCURRENT DUCTS OF THE TESTIS

The excurrent ducts of the testis which comprise various ducts that transport spermatozoa and fluid produced by the seminiferous epithelium, in birds, have, surprisingly, been described in only a few species: the rooster (Gray 1937; Lake 1957; Marvan 1969; Tingari 1971, 1972; Budras and Sauer 1975a,b), Turkey (Hess and Thurston 1977; Hess et al. 1976; Aire 2000a; Aire and Josling 2000), Guineafowl (Aire et al. 1979; Aire 1980, 1982a), Japanese quail (Aire 1979a, 1980, 1982b, 2000a; Clulow and Jones 1982, 1988), Ostrich (Budras and Meier 1981; Aire and Soley 2000, 2003), passerine birds (Bailey 1953; Traciuc 1967, 1969; Middleton 1972; Barker and Kendall 1984), Pigeon (Stefanini et al. 1999), Mute Swan (Mehrotra 1962, 1964) and drake (Marchand and Gomot 1973; Aire 1982a,b; 2000a). The excurrent duct system of birds, as in mammals, comprises (a) the rete testis (RT) unit, (b) the efferent duct unit and (c) the epididymal duct unit. The epididymal duct unit, in turn, comprises the ductus conjugens or connecting duct, the ductus epididymidis or epididymal duct and the ductus deferens or deferent duct. The epididymis of birds is not divided into the

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Reproductive Biology and Phylogeny of Birds

Fig. 2.15 Struthio camelus. A. A Leydig cell displays an array of rod-shaped structures (R) in longitudinal profiles. These structures, some of which appear branched (arrows), are more electron dense than the smooth endoplasmic reticulum (SER) (arrowheads). Only a few profiles of rough endoplasmic reticulum or sparsely granulated endoplasmic reticulum (squat arrows) occur in the Fig. 2.15 Contd. ...

Anatomy of the Testis and Male Reproductive Tract

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customary gross segments of caput, corpus and cauda epididymides, seen in mammals, but it is a complex structure (Fig. 2.16) consisting of several duct units (Lake 1957; Tingari 1971; Budras and Sauer 1975; Hess et al. 1976; Aire 1979a; Aire et al. 1979; Budras and Meier 1981). This organ incorporates the extratesticular portion of the RT, the efferent ducts, the connecting and epididymal ducts (Fig. 2.17). A description of the histological and ultrastructural features of the various duct units of the epididymis will be based, in the main, on the various reports referred to, above.

2.3.1

The Rete Testis

The rete testis (RT) of birds is connected to the seminiferous tubules whose products it transports to the succeeding segment of the excurrent duct system. The occurrence of divisions, based on their location, of the RT has been a controversial subject. Tingari (1971) considers that the rete is located entirely outside the testis but it is generally agreed that there are intracapsular (intratunical) and extratesticular portions of this duct unit. In addition, Bailey (1953), Budras and Meier (1981), Aire (1982a) and Stefanini et al. (1999) have described an intratesticular portion which is made up of rete channels that are similar in epithelial lining to those in the intracapsular and extratesticular segments, or appear in the form of tubuli recti in various species of birds (Gray, 1937; Mehrotra 1964; Aire 1979a; Aire et al. 1979; Budras and Meier 1981; Barker and Kendall 1984). Osman (1980), however, reports that the seminiferous tubules of the testis of the domestic fowl are joined to the RT in three ways: by way of a terminal segment and a tubulus rectus, a terminal segment only, or by direct opening into the rete lacunae and Lake (1957) describes a transitional tubule, lined by modified Sertoli cells, at the junction between the seminiferous tubule and RT. The intratesticular portion of the RT is rarely observed in birds, but in both birds and mammals it constitutes a minor part of the duct unit (Amann et al. 1977; Roosen-Runge and Holstein 1978; Budras and Meier 1981). They are more distinctly evident in regressed testes (Fig. 2.17B). However, in birds, most of the seminiferous tubules terminate by opening into the RT directly (Aire 1982b). The main part of the RT of birds is extratesticular, forming an integral part of the epididymis. Both the intracapsular and extratesticular parts of the RT of the Ostrich (Budras and Meier 1981) are even more complex in organisation than in most of the other birds studied. They consist of a series of connected Fig. 2.15 Contd. ...

cytoplasm. The perimeter of the cell is demarcated by a basal lamina-like material (double arrowheads). B. Rod-shaped structures, as in Fig. 2.15A, are sectioned transversely. Strands of SER and the rods seem to be continuous or closely associated (arrowheads). Some SER profiles display thickened membranes in the lumen (arrows), probably indicating the formation of rod-shaped structures from normal elements of the SER. An extensive Golgi complex (G), a dense body (D) and a vesicular nucleus (Nu) are present in the cell. Bars: A = 1 µm; B = 2 µm. (From Soley J. T., van Wilpe, E. and Aire, T. A. unpublished micrographs)

$" Reproductive Biology and Phylogeny of Birds

Fig. 2.16 Gallus domesticus. A diagrammatic representation of the epididymis and its duct units, based partly on casts. T, testis; RT, rete testis; P, proximal efferent ductule; D, distal efferent ductule; CD, connecting ductule; DE, ductus epididymidis; DD, ductus deferens. From Nasu, T., Nakai, M., Murakami, T., Saito, I., and Takahara, H. 1985 Japanese Journal of Zootechnical Science 56: 81-85. Fig. 2.1-7. Reproduced with the kind permission of Japanese Society of Animal Science.

longitudinal cisterns and ducts that open ultimately into the efferent duct unit (Budras and Meier 1981). The distribution of the RT in mammals also varies between species, being septal and mediastinal in the rat, Rattus norvegicus (Roosen-Runge 1961; Dym 1976; Hermo and Dworkin 1988), Guinea pig, Cavia porcellus (Fawcett and Dym 1974), Goat, Capra hircus (Ezeasor 1986; Goyal et al. 1992), bull, Bos taurus (Hees et al. 1987) and man, Homo sapiens (Roosen-Runge and Holstein 1978). An extratesticular portion appears to be found in only a few mammalian species, e.g. in man (Roosen-Runge and Holstein 1978) and goat (Goyal et al. 1992). In birds, the RT is the smallest duct unit, by volume, in both the epididymis (Table 2.1), and the entire excurrent duct system, constituting only 2.3% of the extra-testicular ducts in the Japanese quail (Clulow and Jones 1988). But, the ratio of surface area of luminal border:luminal volume for this duct unit (43.4:1) is quite large in the Japanese quail (Clulow and Jones 1988).

2.3.1.1

Surface features of the rete testis tubules/lacunae

Scanning electron microscopical features of the RT of birds, have been reported in the drake (Aire 1982a), Ostrich (Aire and Soley 2000) and certain domestic galliform birds—Turkey, rooster, Guineafowl and Japanese quail (Bakst 1980; Aire and Josling 2000). The RT spaces are, not infrequently, broken up into interconnecting cavernous spaces by lamellae or sheets of connective tissue which are, themselves, lined by the rete epithelium. Narrow cylindrical struts of connective tissue or chordae retis, lined by rete epithelium,

Anatomy of the Testis and Male Reproductive Tract

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Fig. 2.17 Coturnix japonica. A. Low power histological view of the epididymis showing profiles of the rete testis (RT) opening into ductuli efferentes proximales (PED). DED, ductuli efferentes distales; DC, ductus conjugens and DE, ductus epididymidis. B. A histological section of the para-epididymal region of an involuting testis showing exaggerated profiles of the intratesticular duct system (asterisks), which are portions of the rete testis. TC, testicular capsule; IR, capsular portion of the rete testis. Bar: A = 200 µm, B = 100 µm. Original.

$$ Reproductive Biology and Phylogeny of Birds Table 2.1

Species variation in volumetric proportions (%) of epididymal ducts and structures Species and number of birds

Structures

Chicken (4) Japanese quail (5) Guinea-fowl (3) Turkey (3) Ostrich (4)

Rete testis Proximal efferent duct Distal efferent duct Connecting duct Ductus epididymidis Connective tissue Blood vessels Aberrant ducts

13.3 ± 1.5* 27.6 ± 3.9

9.9 ± 0.7 40.8 ± 3.5

10.7 ± 1.8 45.7 ± 4.2

7.7 ± 1.3 2.3 ± 0.4

15.2 ± 2.5 1.7 ± 0.4

16.2 ± 2.1 0.7 ± 0.0

7.6 ± 0.4 38.7 ± 3.7 2.5 ± 0.4 0.3 ± 0.0

2.4 ± 0.6 27.3 ± 5.4 2.7 ± 0.5 –

1.8 ± 0.2 22.6 ± 1.5 2.3 ± 0.3 –

14.0 ± 6.8 28.1 ± 3.9

2.4 ± 1.8

38.8 ± 4.1 4.4 ±1.6 –

58.2 ± 4.9 1.8 ± 1.4 –

78 11.8 ± 1.8 6.2 ± 2.8 9 5.2 ± 2.2 7 8 26.1 ± 4.1 3.3 ± 1.6 9

* ± Standard error. Adapted from Aire, T.A. 1979. Journal of Anatomy 129: 703 – 706, Table 1, with permission of Blackwell Publishing Ltd.

may be seen to traverse the lacuna, linking opposite walls. Chordae retis, named by Roosen-Runge and Holstein (1978), may be a common feature of the RT in many species, and probably acts as transluminal coupling device for the contractile system of myofibroblasts (Hees et al. 1989), as do the trabeculae septomarginales of the heart. The surface of the epithelial lining of the RT ductules, except for a few minor details, is generally regular, but may bear a few shallow grooves in the ostrich (Aire and Soley 2000). The apical cell outlines are elongated or polygonal in shape. The cell surfaces extend into short, stubby regular microvilli which vary from very few and sparsely distributed to very numerous and evenly distributed (Fig. 2.18). A single, central cilium projects from most cells into the duct lumen in all birds studied (Aire 1982a,b; Aire and Soley 2000; Aire and Josling 2000). The solitary cilium, that exhibits the 9+2 axonemal structure in the bull (Hees et al. 1989), and is probably sensory in function, appears to be a common feature of the rete cells in animals, as it has also been reported in other mammals, including man (Dym 1976; RoosenRunge and Holstein 1978; Goto 1981; Hees et al. 1989). The solitary cilia of the non-ciliated cells of the human oviduct, which are similar to those described in the male reproductive organs, have been shown, using phase- or videomicroscopy, to have vortical or funnel-like movement (Odor and Blandau 1985; Nonaka et al. 1998), contrary to Ghadially’s (1997) assertion that they are immotile. In most birds studied, especially in the rooster, macrophages are seen in the RT lumen, resting on the epithelium (Aire and Malmqvist 1979a; Osman 1980; Budras and Meier 1981; Aire 1982b; Aire and Josling 2000). Only a few spermatozoa and earlier germ cell series occur in the lumen, and each spermatozoon in the RT of the ostrich bears a single, spindle-shaped, distal cytoplasmic droplet (Aire and Soley 2000), a phenomenon that appears to be unique to this bird, and perhaps other ratites. The motility and fertilizing ability of the rete spermatozoa, and, indeed, of spermatozoa in other segments of the excurrent ducts in the ostrich need to be investigated, as has been done

Anatomy of the Testis and Male Reproductive Tract



%$Fig. 2.18 Coturnix japonica (A), Gallus domesticus (B, C). Surface morphology of the rete testis epithelium. A. Short, stubby microvilli are concentrated centrally and around the edge of the cell in the quail, or B. are evenly and sparsely distributed throughout the surface. In C. the microvilli are restricted to the edges of the cell surfaces. In B. and C., solitary cilia project from most cells. Bars: A = 10 µm; B and C = 2 µm. Original.

$& Reproductive Biology and Phylogeny of Birds for some other birds (Munro 1938; Bedford 1979; Howarth 1983, 1995), for purposes of comparability.

2.3.1.2

The histology and ultrastructure of the rete testis cells

The rete epithelium varies from simple, low cuboidal to squamous. Because the apical portions of some of the rete cells overlap sections of adjacent cells, the histological sections of the epithelium often display a pseudostratified appearance (Fig. 2.19). This appears to be a common feature in all birds except the ostrich in which the epithelium is almost always simple (Aire and Soley 2003). In mammals, the rete epithelium is simple squamous to low columnar (Leeson 1962; Dym 1976; Bustos-Obregon and Holstein 1976; Osman 1978; Hees et al. 1989). The rete epithelium contains only one, non-ciliated cell type. Other cell types that may be found in the epithelium are intraepithelial lymphocytes (Fig. 2.19A) and an occasional ciliated cell. Ciliated cells have been described by Barker and Kendall (1984) as being a usual component of the epithelium in some wild birds, but Aire (2002a) is of the opinion that a few scattered ciliated cells occur in the rete epithelium of gonadally-resting birds. Most of these cells, obviously, are lost during recrudescence, preparatory to resumption of active gonadal function. The surfaces of the rete cells display short, straight and regular microvilli, in well-fixed tissue. Adjacent lateral cell membranes, or parts thereof, are either straight and regular (Tingari 1972) or form complex interdigitations (Aire 1982b; Aire and Soley 2003) that may extend to the basal part of the cell. Extensive, intricate cell membrane interdigitation is probably associated with active transport of substances (Morales et al. 1984) but there is little net fluid reabsorption in the RT of the Japanese quail (Clulow and Jones 1982). The apical tight junctional complexes between the rete cells contain only one or two punctate fusions which, however, do not permit tracer compounds to reach the duct lumen via the paracellular space (Nakai and Nasu 1991). This type of junctional complex has also been reported in the mammalian rete epithelium (Dym 1976) but Claude and Goodenough (1973) regard it as a ‘leaky’ junctional complex. Desmosomes are also present, both in the apical and basal zones of the plasma membrane (Barker and Kendall 1984). In the ostrich, a unique lateral cell membrane modification, similar, in some respects, Fig. 2.19 Coturnix japonica (A) and Anas platyrhynchos (B). Transmission electron micrographs of rete testis epithelial cells. A. The epithelium often appears pseudostratified. The nuclei (N) are deeply indented, euchromatic and irregular in shape. The cytoplasm contains sparse organelles. L, an intraepithelial lymphocyte; P, periductal tissue. B. Part of the rete cell exhibiting short, stubby microvilli (arrowheads), a number of multivesicular bodies (arrows), a small Golgi complex (G), a few subapical vesicles (V) and round/oval profiles of mitochondria (M). Bars: A = 20 µm; B = 1 µm. A is adapted from Aire, T. A. 2002 Anatomy Histology Embryology 31: 113-118, Fig. 2, with permission of Blackwell Publishing Ltd., and B is adapted from Aire, T. A. 1982 Journal of Anatomy 135: 97-110, Fig. 15, with permission of Blackwell Publishing Ltd.

Anatomy of the Testis and Male Reproductive Tract

Fig. 2.19

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% Reproductive Biology and Phylogeny of Birds to a hemi-desmosome, occurs frequently along the length of this membrane. Its function is unknown (Aire and Soley 2003). Rete cell nuclei are typically irregular in outline, being often deeply indented and exhibiting marginated chromatin as well as a single, central nucleolus (Fig. 2.19A). However, in the ostrich, the nuclei have regular, vertically elongated profiles, and are more euchromatic than in other birds. Oval to elongate mitochondria occur above and below the nucleus in moderate numbers, and are more numerous in the ostrich rete cells (Aire and Soley 2003) than has been reported for other birds (Tingari 1972; Aire 1982b; Barker and Kendall 1984). The Golgi complex is moderately large and usually lies in the perinuclear region (Fig. 2.19B). Other organelles worthy of note are multivesicular bodies and strands of rough endoplasmic reticulum, both of which may be numerous in the drake (Aire 1982b). A few, small, lipid droplets are a constant feature of this cell. However, in the ostrich, a solitary, large heterogeneous lipid droplet is a consistent feature in the immediate supranuclear region (Aire and Soley 2003). Intermediate filaments commonly surround the nuclei and support the basal part of the cell, especially in the ostrich (Aire 1982b; Aire and Soley 2003). The chordae retis is lined by an epithelium that is similar to that of the rest of the RT, but its core substance is composed of loose spongy connective tissue, as has also been observed in the bull (Hees et al. 1989). Phagocytised sperm fragments are found occasionally in the rete cell of the rooster (Tingari 1972), and in certain mammalian species (Sinowatz et al. 1979; Holstein 1978). Luminal macrophages contain ingested fragments of spermatozoa only occasionally (Aire and Malmqvist 1979a; Aire and Josling 2000), apparently, removing by ingestion, only damaged or abnormal spermatozoa, but they become avidly spermiophagic in vasectomized or vasoligated birds (Tingari and Lake 1972b; Aire and Heath 1977; Nakai et al. 1989b; Aire 2002a). The subepithelial tissue is composed of dense connective tissue and myofibroblasts or smooth muscle cells (Tingari 1972; Aire 1982b; Aire and Soley 2003). This tissue is richly innervated (Tingari 1972), but poorly vascularized (Barker and Kendall 1984; Nakai et al. 1988) by a non-fenestrated capillary network that is irregularly arranged (Fig. 2.20) (Nakai et al. 1988). Aire (1982b) has observed a rich lymphatic drainage system in the subepithelial tissue in the vicinity of the RT in the drake. Intraepithelial lymphocytes, exhibiting irregular outlines, light-staining cytoplasm and considerably heterochromatic nuclei (Fig. 2.19A), are not uncommon components of the rete epithelium in birds (Aire and Malmqvist 1979b; Aire 1982b; Osman 1980; Stefanini et al. 1999) and mammals (Dym and Romrell 1975; Hees et al. 1989). They have sparse organelles, as in mammals, and there are no junctional complexes between them and epithelial cells. They occur at all levels in the epithelium. These cells probably monitor the epithelium with a view to sequestering germ cell antigens and preventing their escape into the blood or periductal connective tissue (Dym and Romrell 1975; Osman and Plöen 1978).

Anatomy of the Testis and Male Reproductive Tract

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Fig. 2.20 Gallus domesticus. Three types of microvasculature of the rete testis ductule (a), efferent ductule (b) and epididymal duct and ductus deferens (c) are shown diagrammatically. The rete testis has a sparse and irregular capillary network, while both the efferent ductule and epididymal duct unit have a dense capillary network. The meshwork of capillaries is polygonal in (b) and elongated in (c). From Nakai, M., Hashimoto, Y., Kitagawa, H., Kon, Y. and Kudo, N. 1988 Japanese Journal of Veterinary Science, with the kind permission of Japanese Society of Veterinary Medical Science.

2.3.1.3

Functions of the rete testis

The functions of the avian RT are still quite conjectural. The RT constitutes only 2.4% in a preliminary study in the ostrich, and between 10% and 13% of the entire volume of the epididymis in galliform birds (see Table 2.1). This does not take into account the small intratesticular and intracapsular portions of this duct unit. The RT is, understandably, the smallest component (2.3%) of the units of the extratesticular genital ducts of the Japanese quail (Clulow and Jones 1988). The histology and ultrastructure of the RT epithelium in birds (Tingari 1971 1972; Budras and Sauer 1975; Hess and Thurston 1977; Hess et al.1976; Aire 1979a, 1982b; Budras and Meier 1981; Barker and Kendall 1984; Aire and Soley 2003) is similar, in most respects, to those of mammals (Dym 1976; Roosen-Runge and Holstein 1978; Goto 1981; Hees et al. 1989), and because of the ‘leaky type’ apicolateral junctional complex between the epithelial cells (Claude and Goodenough 1973; Lopez et al. 1997), this segment of the excurrent duct system may be a particularly weak one, with regard to the blood-epididymal barrier. In mammals, the RT epithelium secretes about 65% of total testicular fluid (Tuck et al. 1970), but the fluid entering the rete testis is the primary secretion of the seminiferous tubules in the quail (Clulow and Jones 2004). However, even though there is a net reabsorption of testicular fluid by all of the epididymal ducts in the Japanese quail, there is little net fluid transport across the RT epithelium (Clulow and Jones 1988).

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Reproductive Biology and Phylogeny of Birds

Spermatozoa spend only 25 sec in traversing the RT in the Japanese quail (Clulow and Jones 1988), and therefore the through-flow that enters the proximal efferent duct is highly diluted fluid which contains only a few germ cells (Aire 1979a, 1982b). The RT cells are capable of endocytic activity by both fluid-(bulk)-phase and adsorptive endocytosis in the rat (Morales et al. 1984). Tracers interiorised from the lumen by rete cells are disposed of by the lysosomal system in a similar fashion, irrespective of mode of endocytosis. Morales et al. (1984) have concluded that the RT cells may play a role in determining the composition of the RT fluid. These cells in the Japanese quail are also capable of interiorizing intraluminally-introduced India ink particles which are then conveyed to the lysosomal system of the cells (Aire T.A. unpublished observations). However, RT cells fail to phagocytize intraluminally-introduced avirulent strain of Salmonella gallinarium organisms in the Japanese quail (Aire et al. 2004). The myofibroblast layer surrounding the RT lacunae is well developed, and probably acts to move testicular fluid forward into the efferent ducts. The relaxation of the contractile layer, including the chordae retis, could suck fluid from the seminiferous tubules, by means of vis-a-fronte forces, into the cavernous spaces of the RT. Actin, desmin and vimentin filament staining are strongly positive in the myofibroblast layer in the Japanese quail and drake but only slightly positive in the ostrich (Pers. obs.).

2.3.2

The Efferent Ducts (Ductuli Efferentes)

In birds, the RT lacunae are continued by the efferent duct unit (Bailey 1953; Lake 1957; Traciuc 1969; Tingari 1971; Budras and Sauer 1975a; Hess et al. 1976; Aire 1979a, 1980; Aire et al. 1979; Budras and Meier 1981; Bellamy and Kendall 1985), as in mammals (Reid and Cleland 1957; Ladman and Young 1958; Ilio and Hess 1994). The volume proportion of the efferent ducts in the epididymis varies between 35% and 62% in domestic Galliform species of birds and 12% in the ostrich (Table 2.1), and about 19% of the total volume of the genital ducts in the Japanese quail (Clulow and Jones 1988). Two different but serially arranged segments of the efferent duct occur in birds, viz., proximal efferent duct [ductuli efferentes proximalis] (PED) and the distal efferent duct [ductuli efferentes distalis] (DED). The latter is the distal continuation of the PED, and opens into the connecting duct [ductulus conjugens] (Budras and Sauer 1975a; Aire 1979a). The ductuli efferentes, according to Ilio and Hess (1994), are unique in being the only duct unit of the male reproductive tract that is lined by a ciliated epithelium. Whereas in birds there is a distinct histological division of the efferent ducts into the two segments, PED and DED, such a division is not clearly marked in mammals, among which are species-based differences in the non-ciliated cell types present along the length of the duct. Thus, in the laboratory animals, there is a single type of non-ciliated (NC) cell (Hamilton 1975; Robaire and Hermo 1988) and three types in the cane rat, Threonomys swinderianus (Aire and van der Merwe 2003), man, goat and bull (Goyal et al.1992; Morita 1966; Goyal and Hrudka 1981; Gray et al. 1983; Goyal and Williams 1988). Ciliated (C) cells in

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birds, as in mammals, do not vary structurally along the efferent duct, but the non-ciliated (NC) cell types (types I and II, in the PED and DED, respectively) exhibit different and characteristic cytological features (Aire et al. 1979; Aire 1980). Tingari (1971,1972), Marchand and Gomot (1973), Hess et al. (1976), Hess and Thurston (1977) regarded the distal efferent ductule to be the connecting duct, in error, and, consequently, described only a single nonciliated cell type in the efferent duct system.

2.3.2.1

Surface morphology of the efferent ducts

The surface features of the efferent ducts have been described, using scanning electron microscopy, in the rooster and Turkey (Bakst 1980; Aire and Josling 2000), drake (Aire 1982a), rooster, drake and Japanese quail (Aire and Josling 2000) and Ostrich (Aire and Soley 2000). The following account is derived from these reports. Proximal Efferent Ducts (PED). The epithelial lining, and, in some instances, the mucosa of the PED is highly folded, presenting an irregular, festoon appearance (Fig. 2.21A,B). The folds project prominently into the ductal lumen and are lined by cells whose apical surfaces extend into a lush brush border of microvilli (non-ciliated cells) or numerous cilia (ciliated cells). In all investigated birds, the non-ciliated cells (NC) are more numerous than the ciliated (C) cells, and their microvilli are closely packed, long and regularly cylindrical in shape. The microvilli of the C cells are fewer, shorter and thinner than those of the NC cells, and are scattered between the cilia which usually overshadow adjacent NC cells. The microvilli of the NC type I cell of the PED are shorter in the Turkey and very much so in the ostrich than in the rooster and Japanese quail (Aire and Josling, 2000; Aire and Soley 2000). A single cilium projects into the duct lumen from the central region of most NC cells (Aire 1982b; Aire and Soley 2000; Aire and Josling 2000). In vascularly perfused birds (Aire 1982a; Aire and Soley 2000; Aire and Josling 2000), the NC cells do not exhibit apical blebbing, as reported by Bakst (1980), whose specimens were fixed by immersion fixation. Aire (1980; 2000a) regards apical blebbing in this cell type to be a fixation artifact. Distal Efferent Ducts (DED). The epithelium of this round or oval duct is regular and exhibits no folds that are very prominent in the PED. The cilia of the predominant cell type, C cell, overhang the fewer NC type II cells whose apical surface features are similar to those of the NC type I cell of the PED (Fig. 2.21C).

2.3.2.2

Histology and ultrastructure of the efferent duct epithelia

The histology of the efferent ducts in birds has been reported in only a small number of species: in the rooster (Lake 1957; Tingari 1971; Budras and Sauer 1975a, b), turkey (Hess et al. 1976), Japanese quail (Aire 1979a), Guineafowl (Aire et al. 1979), duck (Marchand and Gomot 1973), Pigeon (Stefanini et al. 1999) and the Common Starling, Sturnus vulgaris (Bellamy and Kendall 1985). Histology of Efferent Ducts. The epithelial lining of the ductuli efferentes proximalis (proximal efferent duct) (PED) or occasionally, the mucosa, projects

%" Reproductive Biology and Phylogeny of Birds

Fig. 2.21 Gallus domesticus. Scanning electron microscopy of the surface features of the efferent ducts. A. The duct wall of the proximal efferent duct (PED) displays variably-sized epithelial and mucosal folds. Note that the epithelial surface is devoid of apical cytoplasmic blebs. B. A higher power view of the epithelial surface of PED. C, cilia of ciliated cells; V, microvilli of non-ciliated cells. Cilia of ciliated cells frequently over-shadow the non-ciliated cells which are more numerous that the former cells. C shows an overwhelming preponderance of ciliabearing ciliated cells over the non-ciliated type II cell of the DED. Bars: A = 50 µm; B and C = 5 µm. Original.

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into the duct lumen as folds of varying length and thickness, thus conferring a ‘festoon’ appearance on transverse profiles of the PED (Fig. 3.17A). The epithelium is columnar and pseudostratified because the C cells usually appear truncated between NC cells, and their nuclei are usually situated in the apical half of the cells, while those of the NC type I cells are located in the basal half (Fig. 2.22A). Both cell types make contact with the basement membrane. The NC cells are predominant over the C cells, and, in the common starling in the ratio of 5:1 (Bellamy and Kendall 1985). The luminal content is mainly proteinaceous fluid, in which there is a suspension of sparse spermatozoa and earlier germ cell series. The apical surface of the NC cell is extended by long, closely bunched, regularly cylindrical microvilli that project into the duct lumen. In plastic sections, the subapical region of the cell displays a few vacuoles of varying sizes, below which are rows of round dense bodies extending to the level of the nucleus. The nucleus is round or, more commonly, oval in shape, and contains one or two nucleoli. The C cell is generally of a lighter stain than the NC cell. The nuclei of the C cells are also round or oval, but may be irregular in shape. Even in plastic sections, profiles of organelles in the C cell, other than the nuclei, are hardly discernible. The epithelium rests on a distinct basement membrane that is supported by periductal tissue of fibroblasts, collagen fibers and myofibroblasts. The ductuli efferentis distalis (DED) has a regular profile, both externally and internally (Fig. 2.22B). It has a columnar or high cuboidal, pseudostratified epithelium consisting of NC cells, and a preponderance of C cells. The apical morphological features of these cells are similar to those of the PED. In plastic sections, there are no obvious subapical vacuoles or dense bodies in the NC type II cells of this duct. The C cell is similar, structurally, to that in the PED. The microvasculature of the avian epididymis is derived from branches of testicular artery, including the cranial ureterodeferential ramus, that supply blood to the epididymis in the rooster (Nishida 1964; Nakai et al. 1988) and Ostrich (Elias M, Aire T.A. and Soley J.T. unpublished observations). Smaller branches of these arteries run along the length of the efferent duct and produce a rich periductal network of fenestrated capillaries (Fig. 2.20B). Nerves that supply the epididymis of the rooster are derived from the testicular plexus, and reach the organ by accompanying the testicular arteries (Nishida 1964; Tingari 1971). Both cholinergic and adrenergic components, which have a similar distribution, have been demonstrated in the efferent ducts. These nerve fibers run around the ducts and are associated with the walls of blood vessels and muscle fibers (Tingari and Lake 1972a). Ultrastructure of the Efferent Ducts. Detailed reports of the fine structure of the epithelium in both segments of the efferent ducts (PED and DED) have been published mainly in Galliformes (Tingari 1972; Budras and Sauer 1975a; Hess and Thurston 1977; Aire 1980), drake (Aire 1982b, 2002b) and Pigeon (Stefanini et al. 1999).

%$ Reproductive Biology and Phylogeny of Birds

Fig. 2.22 Gallus domesticus. Histological sections of the PED (A) and DED (B). In A., the NC type I cell (N) contains numerous supranuclear, dense bodies; C, ciliated cell. In B., the NC type II cell (N) lacks the dense bodies found in the corresponding cell type of the PED; C, ciliated cell, and S, spermatozoa in the DED lumen. Bars: A = 200 µm; B = 10 µm. Original.

Anatomy of the Testis and Male Reproductive Tract

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The classification of the NC cell into types I and II is based upon important characteristic organelle differences and disposition between the two nonciliated cells lining the PED and DED, respectively (Aire 1980). In mammals, NC cells have also been known to vary structurally along the length of the efferent ducts: thus there is only one type in rodents (Hoffer and Greenberg 1978; Ilio and Hess 1994), but there are three types in the Great cane rat, Threonomys swinderianus (Aire and van der Merwe 2003), and 3 in man, bull, Goat and Dog, Canis canis (Morita 1966; Goyal and Hrudka 1980, 1981; Gray et al. 1983; Goyal and Williams 1988). Variations in the disposition and number of vacuoles and/or granules have been used as criteria for classifying the non-ciliated cells in the efferent ducts of animals. It is not clearly understood whether or not these organelle differences influence the functions of these cells, individually in mammals, but there appears to be an obvious dichotomy in function(s) between the NC type I and NC type II, in birds (Aire 1979a, 1980, 2000b, 2002b; Aire et al. 2004; Clulow and Jones 1988). The NC Type I Cell. The apical surface extends into a microvillous brush border of closely bunched, long, and uniformly cylindrical microvilli (Fig. 2.23). The apical surface may also invaginate as fuzzy-lined, tubular coated pits into the subapical cytoplasm which is remarkably endowed with an elaborate endocytic or tubulovacuolar system. The tubular coated pits are continued by straight or coiled apical tubules, which together with endosomes made up of large dilated membranous vacuoles, as well as a few multivesicular bodies (MVBs) occupy the apical one-fourth of the cell (Fig. 2.24). The apical tubules frequently contain an amorphous inspissated material, probably protein, taken in from the duct lumen. Distal to the endocytic system there occurs a large number of round, variably-sized homogeneous or heterogeneous dense bodies. The dense bodies may become heterogeneous as a result of endocytosis and lysosomal activity by the cell. This type of dense body is probably a telolysosome. Numerous, elongated or oval mitochondria occur in both the supranuclear, and to a greater extent, in the subnuclear regions of the cell. They may measure up to 0.6 mm in breadth (Aire 2002b). The Golgi complex is moderately developed in the supranuclear region of the cell. The oval nucleus is situated in the basal half of the cell, contains a central or eccentric nucleolus, and is generally euchromatic. Lipid droplets are uncommonly encountered in the cytoplasm. Strands of RER and short, small profiles of SER are scattered in the cytoplasm. The apical junctional complexes between the NC cells of the efferent ducts in G. domesticus show, apico-basally, a series of punctate fusions (zonula occludentes), adhering or intermediate junctions (zonula adherens), and desmosomes (macula adherens), in that order, along the complex. A continuous line of fusion between the outer leaflets of adjacent cell membranes in the apical junctional complex is also evident (Nakai and Nasu 1991). But Claude and Goodenough (1973) and Suzuki and Nagano (1978) regard these junctional complexes as being of the ‘leaky type’, in the rat. However, Nakai and Nasu (1991) have shown that the tight junctions in both the rete and

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Fig. 2.23 Gallus domesticus. Transmission electron micrograph of epithelial cells of the PED. N, non-ciliated type I cell; C, ciliated cell. Note the large number of dense bodies (D) and long mitochondria (arrowheads) in the supranuclear region of the NC cells. The mitochondria of the C cells (arrow) are thinner than those of the NC cells. The lush microvilli (m) of the NC cells are regularly cylindrical and longer than the inter-cilial microvilli of C cells. Bar: 1 µm. Adapted from Aire, T. A. and Josling, D. 2000 Onderstepoort Journal of Veterinary Research 67: 191-199, Figure 12, with permission of the Editor.

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Fig. 2.24 Anas platyrhynchos (A), Gallus domesticus (B, C, D). Non-ciliated type I cells of the PED. A. Supranuclear region of NC type I cells showing an elaborate subapical tubulovacuolar or endocytic system, comprising coated pits (arrowheads), coated apical tubules (arrows) and vacuole (V). D, dense bodies; N, nucleus. B. A coated pit (arrowhead) leads into a non-coiled apical tubule (T); M, microvilli; J, junctional complex. C. Transverse sections of apical tubules contain inspissated material (arrowhead). D. Heterogeneous dense bodies (H) and mitochondria in a cell that has experienced phagocytosis. Bars: A = 10 µm; B = 2 µm; C and D = 1 µm. Adapted from Aire, T. A. 2002 Journal of Morphology 253: 64-75, Fig. 4., with permission of Wiley-Liss, Inc.

& Reproductive Biology and Phylogeny of Birds efferent duct epithelia are able to exclude lanthanum nitrate from the duct lumen in the rooster. It is not known if this complex can exclude compounds of smaller molecular weights from passing into or out of the lumen. Large and extensive intercellular spaces occur between adjacent NC cells in the PED in the Ostrich. These spaces usually occur in the basal two-thirds of the epithelium, and are frequently seen to extend to the basal lamina in electron micrographs. Dilated intercellular spaces are usually associated with enhanced fluid absorption by epithelia (Pudney and Fawcett 1984), and in this case, paracellular fluid movement, apparently by active solute transport (Suzuki and Nagano 1978). The NC Type II cell. The microvillous brush border is similar to that of the NC type I cell. The type II cell lacks the characteristic, elaborate, subapical endocytic apparatus and numerous dense bodies in the supranuclear zone of the type I cell (Fig. 2.25). Instead, only a few, sparsely-distributed, subapical coated pits and apical tubules, containing inspissated material, are scattered between the bundles of microfilaments which project from the microvilli into the subapical cytoplasm. Vacuoles are usually few, small and scattered in the apical half of the cytoplasm. Mitochondria are fewer than in the NC type I, and are scattered within the supranuclear, and to a lesser extent, in the subnuclear cytoplasm. The nuclei are similar to those of NC type I cells in location, size, shape and configuration. Strands of RER and a few quite small dense bodies and lysosome-like bodies may be seen in the cytoplasm. Ciliated Cells. Readily discernible differences are not to be found in the ultrastructure of the C cells in both segments of the efferent ducts, but the number of C cells relative to NC cells increases markedly in the DED, with a ratio of 4:5, respectively, in the Common starling (Bellamy and Kendall 1985) and 9:1 in the ostrich (Budras and Meier 1981) than in the PED that has a respective ratio of C to NC cells of 1:5 in the common starling (Bellamy and Kendall 1985) and 3:7 in the Ostrich (Budras and Meier 1981). Uniformly-spaced cilia, interspersed with a few, short and thinner microvilli than in the NC cells, project into the lumen (Figs. 2.23 and 2.25). A few coated apical pits are also present. Mitochondria are mainly in the supranuclear region of the cell; they are only about 30% as broad as those of the NC cells (Aire 2002b). The Golgi complex is of moderate size. Numerous bundles of microfibrils, possibly assisting in stabilizing the cell whose role includes movement of the luminal through-flow, may be seen running in different directions, in the supranuclear and perinuclear zones of the cell (Aire 1980). A euchromatic nucleus, which is generally oval in profile, but is often indented or invaginated, is situated in the apical half of the cell. Profiles of RER, polyribosomes, a few small dense bodies and lysosomes may be seen in the cytoplasm. The C cell is thought to assist in moving the seminal content of the efferent ducts, in addition to a limited endocytic activity, and, thus, also influencing the composition of the luminal content. Intraepithelial lymphocytes are not uncommonly present, in varying numbers and at various levels, in the epithelium of both segments of the

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Fig. 2.25 Gallus domesticus. A. A survey transmission electron micrograph of the DED epithelium. C, ciliated cell; N, non-ciliated type II cell showing one or two subapical vacuoles; L, intraepithelial lymphocyte. B. High power of the supranuclear region of ciliated (C) and non-ciliated type II (N) cells. The N cell lacks the numerous dense bodies found in the supranuclear region of the corresponding cell in the PED, and only a few apical tubules (arrowheads) are present in the DED. Arrow, microfibrillar bundles in the supranuclear region of a C cell. Bars: A = 200 µm, B = 2 µm. Original.

efferent ducts (Fig. 2.25A), as in the RT epithelium (Aire and Malmqvist 1979a). Function of the Efferent Ducts. The structure of the efferent ducts is generally similar in mammals and birds (Burgos 1960; Ladman 1967; Tingari 1972; Ramos and Dym 1977; Jones et al. 1979; Aire 1980; Hermo et al. 1988; Robaire and Hermo 1988). It is therefore very tempting to ascribe similar functions to the ducts in most, if not all, animals. In birds, however, unlike in mammals, the proximal segment (PED) of the duct possesses a highly folded epithelium

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Reproductive Biology and Phylogeny of Birds

which increases considerably the surface area that is available and exposed to the luminal content. Besides, the PED constitutes a greater proportion of the epididymal volume (about 300% more) than the distal segment (DED), but, together, both of them constitute between 35% and 62% of the entire epididymal volume in various species of birds (Aire 1979b). This is significant because the avian testis has a high fluid content and sperm production, the latter being also very rapid. The structure of the PED epithelium is consistent with active uptake of luminal macromolecules and considerable luminal fluid reabsorption, and transport across the epithelium. The PED must therefore play a major role in modifying luminal content, and, in general, the functioning of the avian excurrent duct system, as in mammals. However, little is known about endocrine regulation of the avian male reproductive tract and its role in production of fertile spermatozoa (Janssen et al. 1998). The efferent ducts are probable sites of steroidogenesis in the rooster (Tingari 1973), but androgens do not prolong sperm viability in the ductus deferens of the rooster (Munro 1938). Estrogen receptors are strongly expressed in its efferent ducts (Kwon et al. 1997) and in several mammals (Goyal et al. 1997, 1998; Fisher et al. 1997). Estrogens seem to have a profound effect on the ability of the efferent ducts in Mouse (Mus musculus) to reabsorb luminal fluid (Hess et al. 1997). In mammals, micropuncture studies indicate that the efferent ducts reabsorb most of the testicular fluid entering the excurrent ducts of the testis (Crabo 1965; Jones 1980; Jones and Clulow 1987; Clulow and Jones 1988; Clulow et al. 1994; Man et al. 1997). In their excellent studies in the Japanese quail, Clulow and Jones (1988) show that even though spermatozoa (in their fluid medium) spend only 3 min traversing the PED, yet about 86% of the fluid leaving the testis is reabsorbed there, and another 6.5% in the DED. Important ion transporters, such as sodium-potassium ATPase [Na+, K+-ATPase] carbonic anhydrase II (CA II) and sodium hydrogen exchanger isoform 3(NHE3) have been immunolocalized in the efferent ducts of the rooster (Bahr et al. 2006). Transmembrane water channel proteins (aquaporins –2, –3, and –9) that are responsible for water flow, have similarly been localized in efferent ducts of the large white turkey (Zamboni et al. 2004). The arrangement of the efferent duct unit into a large number of narrow ducts in a parallel array provides a large ratio of luminal surface area:luminal volume in the Japanese quail (Clulow and Jones 1988). Spermatozoa traverse the entire efferent duct unit in 8 min in the Japanese quail (Clulow and Jones 1988) in contrast with about 45 min in the rat (English and Dym 1981). It is clear that the rate of fluid reabsorption of testicular fluid by the efferent ducts of the testis is much higher in birds than in mammals (Jones 1998). The inspissated material in the lumen of apical tubules of the NC type I cell in birds (Fig. 2.24C) is probably proteinous, and, if so, it indicates that this type of cell, and therefore the entire efferent duct unit, is capable of absorption of testicular proteins secreted into the testicular fluid, as has been established for mammals (Koskimies and Kormano 1975; Olson and Hinton 1985; Jones and Jurd 1987; Veeramachaneni and Amann 1991; Clulow et al. 1994).

Anatomy of the Testis and Male Reproductive Tract

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Morales and Hermo (1983) and Hermo and Morales (1984) have demonstrated that the non-ciliated cells of efferent ducts are capable of internalizing specific substances from the duct lumen by both adsorptive and fluid-phase endocytosis in the rat. Nakai et al. (1989a) and Aire (2000a) have also demonstrated the respective ability of NC cells in the PED of birds to interiorize testicular proteins and India ink particles (Fig. 2.26). The absorbed materials are destined for the lysosomal system of the cell. The NC cells in the PED are able to recognise, by an unknown mechanism, and remove cationic ferritin, even in bicameral chambers (Janssen et al. 1998), as well as ‘designated’ spermatozoa (Hess et al. 1982; Aire 2000b, 2002a) and intraluminally introduced avirulent strain of Salmonella gallinarium (Aire et al. 2004) from the luminal through-flow in the rooster and Japanese quail. In vasectomized birds, desquamated germ cells that are transported from the seminiferous tubules to the excurrent ducts are sequestered in the PED where they degenerate and their fragments are removed by phagocytic acitivities of the NC type I cells (Tingari and Lake 1972b; Aire and Heath 1977; Nakai et al. 1989b; Aire 2002a). The DED seems to be screened, by an unknown mechanism, from such germ cell debris by the PED. Remarkably, the NC type I cells are also capable of proliferating and forming new adluminal sheets of cells which are highly spermiophagic on both their free surfaces, in the process of removing an overwhelming accumulation of sperm debris, following vasectomy in the Japanese quail and rooster (Aire 2000b, 2002a) or carbendazim exposure in Coturnix (Aire 2004). These new sheets of cells subsequently sequester small ducts from the original duct lumen, during the process of microrecanalization in the efferent ducts (specifically the PED segment) in birds (Aire 2004). Microrecanalization, specifically of the transected end of the vas deferens, has been described previously only in vasectomized men, and was probably responsible for unexpected fathering of babies by such men (Cruickshank et al. 1987; Freund et al. 1989).

2.3.3 The Epididymal Duct Unit 2.3.3.1 General organization and features From ontogenetical, structural and functional perspectives, the connecting duct (ductus conjugens), epididymal duct (ductus epididymidis) and deferent duct (ductus deferens) are essentially similar and constitute a functional unit (Gray 1937; Lake 1957; Tingari 1971, 1972; Budras and Sauer 1975a; Budras and Meier 1981; Aire 1979a; Aire et al. 1979; Aire 2000a). This is also in accord with the observations made by Tingari (1971, 1972) and Aire et al. (1979), that the NC type III cell is identical in the epithelia of these ducts, which shows that the avian ductus epididymidis and ductus deferens are merely different segments of the same organ, an organ equivalent to, but not grossly structured as, the epididymis in mammals. This functional unit may therefore be referred to, for convenience, as the “epididymal duct unit”. This duct unit forms the greater proportion (about 74%) of the entire excurrent duct volume than all other duct units put together, in the Japanese quail (Clulow and Jones 1988).

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Fig. 2.26 Coturnix japonica. A. Rete testis-infused India ink particles are present in the non-ciliated, but not the ciliated cells of the PED epithelium, and are absent in the epithelial cells of the DED and ductus epididymidis (DE). B. India ink particlefilled lysosomes are present in transverse sections of non-ciliated (N) cells but not ciliated (C) cells of the PED. Bars: A = 200 µm; B = 1 µm. Original.

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Similarly, the epididymis and vas deferens in mammals, together, constitute, a priori, the greatest proportion by volume of the excurrent duct system. The efferent ducts are continued distally by the epididymal duct unit that is derived from the Wolffian duct (Budras and Sauer 1975a). This duct is lined by a non-ciliated epithelium. The first portion of this unit is the short connecting duct (CD) that joins the efferent duct, specifically, the DED segment of the efferent duct unit, to the epididymal duct (ED). The CD opens into the ED which is a slightly wavy duct (Fig. 2.16), but may be complexly tangled in the Common Tern (Sterna hirundo) and Jackdaw (Corvus monedula) (Traciuc 1967, 1969). It is situated longitudinally on the dorsomedial border of the epididymis, and runs caudally into the ductus deferens (DD). Neither the ED nor DD is therefore similar to those of mammals in length, gross features or configuration.

2.3.3.2

The ductus deferens and its modifications

In non-passerine birds, the ductus deferens is highly wavy and increases in diameter cranio-caudally, in sexually active birds (Fig. 2.1). Close to the cloaca, the ductus deferens straightens out for a variable length, depending on the species and size of bird, to form the pars recta ductus deferentis (Marvan 1969; Lake 1971; Marchand and Gomot 1973). This part subsequently enlarges into a thick-walled, barrel-or spindle-shaped structure, the receptaculum ductus deferentis that pierces the wall of the cloaca to open into the urodeum of the cloaca by an opening in its pointed end, the papilla ductus deferentis. In passerine birds, the ductus deferens is complexly thrown into coils that form a compact ball, the seminal glomus or sac, caudally (Bailey 1953) (Fig. 2.27). The seminal glomus is a tubular and highly coiled structure that is encapsulated in loose connective tissue (Bailey 1953; Salt 1954 and Wolfson 1954). It lies beneath the skin, dorso-lateral to the cloaca (Salt 1954), where its coiled ducts may be visible through the skin of the cloacal protuberance (Mulder and Cockburn 1993). The latter, akin to the scrotum in scrotal mammals, is a swelling of the skin, that accommodates the seminal glomus. The seminal glomus is a characteristic feature of passerine birds (though also reported for the psittacid Melopsittacus undulatus, see Chapter 8), and is regarded as an anatomical adaptation that ensures a storage and probable maturation site for spermatozoa (Lake 1981) that are necessary for sperm competition (Birkhead et al. 1993). It is tempting to regard the lower temperature in the seminal glomus as beneficial to spermatozoa in passerine birds, as it is in scrotal mammals. Wolfson (1954) believes that the lower temperature is necessary for sperm maturation. But, avian sperm do not require maturation and capacitation in order to be fertile (Munro 1938; Bedford 1979; Howarth 1983). The seminal glomus may therefore serve as a daily sperm store since, it appears, sperm production in the testis as well as sperm transport to the glomus seminalis occurs more rapidly at night, while its sperm content declines significantly during the day (Birkhead et al. 1994). The seminal glomus may, in addition to acting as a storage area, provide a

&$ Reproductive Biology and Phylogeny of Birds

Fig. 2.27 A diagrammatic representation of a ventral view of the reproductive organs and tract of a sexually mature and active passerine bird, Passer domesticus (the House sparrow). T, testis; E, epididymis; D, ductus deferens; SG, seminal glomus; CL, cloaca; R, rectum; V, vent; Arrowhead, skin of the cloacal protuberance. Not drawn to scale. Original.

lower temperature for spermatozoa, in order to conserve their energy or perhaps permit a biochemical or physiological reaction or adjustment in the spermatozoa, before insemination into the female tract. The precise function of the seminal glomus remains speculative. This structure is neither homologous nor analogous to the seminal vesicle of mammals. The cloacal protuberance is a very useful structure for sexing passerine birds, and in determining the prevailing phase of their reproductive cycle. The size of the protuberance is strongly and positively correlated both with the mass of the seminal glomus and its sperm content (Birkhead et al. 1993)

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and, therefore, a guide to the sperm production capacity of the bird under examination. In the Superb Fairy-Wren (Malurus cyaneus), a pointed anterior ‘tip’ of the cloacal protuberance, not yet described in other passerine birds, probably facilitates effective sperm transfer, during short periods of copulation (Mulder and Cockburn 1993).

2.3.3.3

Surface features of the epididymal duct unit

The epididymal epithelium, at low power magnification, appears smooth and presents no folds except at sharp angulations and at the entry of the CD into the ED where several longitudinal ridges and grooves occur (Aire and Soley 2000). A number of irregularly distributed invaginations or ‘craters’ (Fig. 2.28) occur on the epithelial surface in the drake (Aire 1982b). In all birds studied (the ostrich by Aire and Soley 2000; Turkey, rooster and Japanese quail by Aire and Josling 2000), except the drake (Aire 1982b), the luminal surfaces of the CD, ED and DD appear cobbled, with distinct intercellular grooves. Close-up views show numerous, evenly distributed, regular microvillar extensions of the apical surfaces of the principal epithelial cells (Fig. 2.28B). A single, central cilium projects from several cell surfaces into the duct lumen.

2.3.3.4

Histology of the epididymal duct unit

In non-passerine birds, the histological features of the epididymal duct unit have been reported in several species. Discrepancies or errors in the nomenclature of the ducts, as well as the interpretation of normal structure, are to be found in the literature (Gray 1937; Lake 1957; Tingari 1971, 1972; Budras and Sauer 1975a; Hess et al. 1976; Aire 1979a, 2000a). These will be highlighted in this review. The CD, ED and DD are lined by cuboidal to columnar, non-ciliated epithelium, simple or pseudostratified, depending on the angle of section (Fig. 2.28). The epithelium consists of the non-ciliated type III cell (Aire et al. 1979) which is distinctly different, ultrastructurally, from the non-ciliated types I and II cells of the PED and DED, respectively. Basal cells, not present in the more proximal duct units, are wedged between the NC type III cells, and their long axes typically lie parallel to the basal membrane. Basal cells increase in number, cranio-caudally, i.e., they are least numerous in the CD and ED, but quite numerous and virtually form a distinct layer of cells in the caudal part of the DD. The ductal lumen is regular in outline and oval in cross-section, save at the entry of the CD into the ED where epithelial ridges and grooves occur (Aire and Soley 2000). A short microvillous brush border, varying in height from 0.7 mm in the drake to 1.6 mm in the Japanese quail (Aire 2000a) projects from the NC type III cells into the lumen. Nuclei of the NC type II cells are round or oval in shape, and display single, central nucleoli. Intraepithelial lymphocytes are also present in the epithelium (Aire and Malmqvist 1979b). The epithelium rests on a compact, richly vascular peritubular boundary tissue composed of fibroblasts, collagen fibers and several layers of smooth muscle cells (Aire 2000a). In passerine birds, not much has been reported on the normal structure of the excurrent ducts of the testis, even though passerines constitute a

&& Reproductive Biology and Phylogeny of Birds

Fig. 2.28 Anas platyrhynchos (A), Meleagris gallopavo (B) and Struthio camelus (C). In A., A SEM view of the surface epithelium of the epididymal duct, showing a regular surface interspersed with crater-like depressions. Several solitary cilia Fig. 2.28 Contd. ...

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significantly large proportion of all birds. In fringillids, the epithelium of the epididymal duct is reported to be columnar and ciliated (Bailey 1953) but that of the ductus deferens is columnar or pseudostratified and non-ciliated. Bedford (1979) considers that the DD has a ciliated epithelium. The apical cytoplasm of NC cells blebs into the duct lumen (Bailey 1953), in a manner that is reminiscent of apocrine secretion. Care must be taken in regarding these blebs as representing apocrine secretion, because histological sections of this tissue, fixed by immersion, but not by vacular perfusion, in non-passerine birds, display such apical blebs that are regarded as artifacts of fixation (Aire 1980, 1982b, 1997, 2000a). Basal cells that lie on the basement membrane are present between non-ciliated cells. The epithelial lining of the seminal glomus is cuboidal to low columnar, and is, also, composed by non-ciliated and basal cells (Bailey 1953; Salt 1954; Wolfson 1954; Middleton 1972). However, Bullough (1942) and Bhat and Maiti (2000) have described a ciliated columnar epithelium. Middleton (1972) considers that the epithelium contains secretory cells whose product is PASpositive, and probably glycogen. The duct is filled with spermatozoa the heads of which are pointed toward, and make contact with, the epithelium, which in some cases is penetrated and eroded by the spermatozoa, thus creating the impression of holocrine secretion by this epithelium. The epithelium is invested by a well-defined fibromuscular layer of boundary tissue. There is a great deal of discrepancy in the nomenclature of the various ducts of the epididymis in these reports. Further and more detailed studies of the structure of the epithelial lining of the various excurrent ducts of the testis of passerine birds is necessary in order to minimise the confusion that exists in the literature. The Receptaculum Ductus Deferentis. This structure displays a large number of epithelial and mucosal folds that are not obliterated, even when full of spermatozoa. The folds are longer and more complex toward the papilla of the receptacle. The epithelium is columnar, may be pseudostratified and comprises non-ciliated cells and numerous basal cells (Fig. 2.29). Several deep grooves or crypts, and their cross-sections occur in histological sections. In the ostrich, the cross-sections of the crypts, especially toward the papilla, Fig. 2.28 Contd. ...

(arrowheads) project from most cells. S, spermatozoon. B. Higher power view of the surface epithelium of the epididymal duct, showing uniformly distributed, regular microvilli and solitary cilia projecting from several cells. C. A histological section of the epididymal duct of the ostrich, exhibiting a columnar profile; basal cells (arrowheads) are present, and increase considerably in number craniocaudally. Bars: A = 200µm; B = 2 µm; C = 200 µm. Figure A is adapted from Aire, T. A. 2000 Anatomy Histology Embryology 29: 179-191, Figure 2. Reproduced with permission; Figure B is from Aire, T. A. and Josling, D. 2000 Onderstepoort Journal of Veterinary Research 69: 191-199, reproduced by permission of the Editor. Figure C is original.

' Reproductive Biology and Phylogeny of Birds

Fig. 2.29 Struthio camelus. Histological sections of the receptacle of the ductus deferens. A. The epithelium projects finger-like processes into the duct lumen. Eosinophilic and PAS-positive, but not glycogen, secretory material (arrowheads) occurs within the epithelium, particularly in the region of the crypts between epithelial folds. B. Basal cells (arrows) are extremely numerous, and line the basal part of the epithelium. Bar: A = 100 µm, B = 200 µm. Original.

often display spermatozoa and/or an eosinophilic content that is PASpositive, non-glycogen polysaccharide material (Fig. 2.29), of which the mode of secretion and function are unknown (pers. obs.). The epithelium of the receptaculum ductus deferentis therefore appears to be secretory and capable of influencing or modulating sperm viablility or nutrition. Its sperm content is highly concentrated, and therefore can be rapidly regulated (Clulow and Jones 1988). In the drake, the epithelial grooves contain only spermatozoa, if any (Aire et al. 1979). The epithelium is enclosed by a very thick coat of fibromuscular tissue. In the rooster, this tissue contains subepithelial sinuses and tortuous arteries, akin to the situation in the mammalian penis (Lake 1957), but the function and mechanism of action of this structural peculiarity, with regard to ejaculation in birds, is unknown. Nevertheless, the receptacle appears to be a temporary storage compartment for a relatively large number of spermatozoa, close the point of their emission.

2.3.3.5

Ultrastructure of the epididymal duct unit

Reports of ultrastructural studies on the avian epididymal duct unit are scarce, and are mainly on the rooster (Tingari 1972; Aire 2000a), turkey (Hess and Thurston 1977; Aire 2000a), drake and Japanese quail (Aire 2000a). Non-ciliated Type III Cells. The apical surface of the type III cells extend into a microvillous brush border (Figs. 2.30 and 2.31), which is only about 65% as

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Fig. 2.30 Anas platyrhynchos. A survey electron micrograph of epithelial cells lining the epididymal duct unit, taken from the ductus epididymidis. Short microvilli, a well developed Golgi complex (G), complexly folded lateral plasmalemma (open arrows); nucleus (N) and an investing layer of intermediate filaments (IF). Numerous mitochondria occupy the supranuclear region of the cell. Bar = 1 µm. Original.

'

Reproductive Biology and Phylogeny of Birds

Fig. 2.31 Anas platyrhynchos. Magnified portions of the non-ciliated type III cell of the epididymal duct unit. A. The microvilli may branch (small arrowhead), the Golgi Fig. 2.31 Contd. ...

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long as those of NC types I and II cells in efferent ducts of the Japanese quail (Aire 2000b). The microvilli are regularly cylindrical in profile, and blebbing of the apical cytoplasm is rarely seen in tissues that are well fixed, either by very good immersion fixation or, more importantly, by good intravascular perfusion fixation. Apical blebbing in NC types I and III cells has been described by various authors (Budras and Sauer 1975a; Hess and Thurston 1977; Bakst 1980; Stefanini et al. 1999). However, Aire (1979a, 1980, 1982b), Aire and Josling (2000) and Aire et al. (1979) have found that these blebs are absent in very well fixed tissues. Nicander (1970) and Hamilton (1975) have also regarded these structures, in mammalian epididymal tissues, as artifacts of fixation. Ericsson (1964) has made similar observations in poorly fixed homologous cells of the kidney. Adjacent lateral plasma membranes are intricately folded (Fig. 2.30), especially in the basal two-thirds of the cell length. These foldings may serve a mechanical purpose of stabilizing intercellular attachments, or in water transport (Pease 1956). Apicolateral junctional complexes are well formed (Friend and Gilula 1972) and both the tight junction (zonula occludens) and adhering intermediate junction (zonula adherens) are composed of multiple punctate fusions which exclude tracer compounds from the duct lumen, in the rooster (Nakai and Nasu 1991). Desmosomes on the lateral membranes and hemi-desmosomes in the basal plasma membrane are also present. The nucleus of the NC type III cell is round or oval in shape in the drake (Aire 2000a) and Ostrich (pers. obs.) or vertically elongated in the rooster, Turkey and Japanese quail (Tingari 1972; Hess and Thurston 1977; Aire 2000a). It is moderately heterochromatic, contains a usually eccentric nucleolus, and is basally situated in the cell. Bundles of intermediate filaments (IFs), up to 640 nm wide, may surround the nucleus, partially or wholly (Fig. 2.30), particularly in the drake (Aire 2000a) and may be seen to attach to other organelles as well as the plasma membrane. The function of this distinct assemblage of IFs in the drake is not clearly understood, but IFs are known to form the cytoskeleton component that connects different parts of the cell into an organic network, act as organizers that control the distribution of different subcellular structures, and as integrators of the cellular space (Lazarides 1980; Geiger 1987 and Zhu et al. 1997). A moderately abundant endoplasmic reticulum occurs in the cell, and are mainly of the sparsely granulated (SGER) type in the drake, in particular, while in other birds, it is represented mainly Fig. 2.31 Contd. ...

complex (G) is well developed but consists of only a few saccules; the cytoplasm contains abundant, distended profiles of SER or sparsely granulated endoplasmic reticulum (SGER) (large arrowheads); the subapical region shows numerous secretory vesicles with a dense content (arrows). B. Apical half of part of the NC type III cell displaying profiles of microtubules (arrowheads) extending to the apical membrane of the cell. Arrows, clathrin-coated vesicles. Bars for both A and B = 2 µm. Original.

'" Reproductive Biology and Phylogeny of Birds by profiles of SER and RER. In Turkey, a whorl of RER occurs invariably in the immediate supranuclear zone (Aire 2000a). Abundant free ribosomes or rossettes of ribosomes occur in the cells. Lipid droplets are uncommon in all birds (Tingari 1972; Aire 2000a) except the turkey in which they are quite abundant, as lipid aggregates, in the supranuclear, and to a lesser extent, infranuclear regions of the NC type III cell (Hess and Thurston 1977; Aire 2000a). The lipid content in the cells, particularly in the Turkey, may be related to some form of steroidogenesis, as Tingari (1973) has shown that 3b-and 17b-hydroxysteroid dehydrogenases are moderately localized in the epididymal duct of the rooster. Mitochondria are dispersed evenly in the cytoplasm, and are each usually surrounded by a strand of RER. A well developed, supranuclear Golgi apparatus occurs in the drake and Japanese quail. Numerous smooth-surfaced vesicles and clathrincoated vesicles are observed in the region of the Golgi complex. The Golgi complex, in this duct unit, is very well developed (Fig. 2.31), but is not as elaborate as in the epididymal duct of mammals (Nicander and Glover 1973; Ramos and Dym 1977; Hermo et al. 1991; Stoffel and Friess 1994). Its function in birds appears quite significant (Aire 2000a). Noteworthy features include numerous secretory vesicles, with increasing condensation of their content, as they move from the peri- and supra-nuclear zone toward the apical plasmalemma (Fig. 2.31B), in the rooster (Tingari 1972) and drake (Aire 2000a). Microtubules extend from the region of the Golgi complex to the apical zone in the Japanese quail (Aire 2000a) and drake. Movement of secretions by means of vesicles and/or microtubules is well established in, possibly, all eukaryotic cells (Palade 1975; Cooper et al. 1990; Darnell et al. 1990). The secretory vesicles contain dense, amorphous material, which is probably discharged in a merocrine manner of secretion into the duct lumen (Tingari 1972; Aire 2000a). The nature of this secretion is unknown. However, about four Wolffian duct proteins bind to spermatozoa as they pass through the epididymis and the ductus deferens in the rooster (Esponda and Bedford 1985; Morris et al. 1987). Only one androgen-dependent protein of 17kDa has been identified in the Japanese quail (Kidd 1982, cited by Jones 1998). These proteins seem to have a specificity that is confined to the same order of birds, in this case, Galliformes (Esponda and Bedford 1985; Morris et al. 1987). Coated pits invaginate into the subapical cytoplasm, and lead to only a few, scattered subapical coated vesicles. These indicate that the NC type III cell is capable of micropinocytosis, probably of luminal proteins, along with a very small quantity of fluid, because Clulow and Jones (1988) have shown that the epididymal duct unit (CD, ED and DD) reabsorbs only about 0.8% of the total testicular plasma output in the Japanese quail. The NC type III cells appear to be incapable of endocytosis of India ink particles infused into the excurrent duct system through the RT (Fig. 2.26A), whereas numerous rete cells and almost all of efferent duct NC type I cells, but not NC type II or ciliated cells, avidly take up the ink particles (Aire 2000a, and unpublished observations). Similarly, the epididymal duct unit fails to interiorize

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horseradish peroxidase, while both the RT cells and NC type I contain large amounts of the substance (Nakai et al. 1989b). On the whole, the NC type III cell, in birds, appears to be similar, functionally, to the principal cell of the rat epididymis, in being involved primarily, but not exclusively, in secretion, while the clear cells, not described in birds, are primarily involved in absorption (Hamilton 1975; Robaire and Hermo 1988; Hermo et al. 1988). Basal Cells. Basal cells are found between the bases of NC type III cells only in the epididymal duct unit, and they increase in frequency, cranio-caudally (Figs. 2.26, 2.28 and 2.29C). They are cuboidal or pyramidal in shape, containing nuclei of varying shape, from elongated to triangular or irregular (Fig. 2.32). The nucleus contains a central nucleolus and heterochromatin aggregations attached to the inner nuclear membrane. The organelle content of the cell is sparse, and includes a small Golgi apparatus, a few mitochondria and strands of RER (Fig. 2.32). The nucleus is encircled by bundles of fibrillar material that are best developed in the rooster and Japanese quail, but are also present, to a lesser extent, in the drake and Turkey (Aire 2000a). Tingari (1972) therefore speculates that basal cells subserve a similar function as myoepithelial cells, in assisting the contraction of the muscular coat, during ejaculation. Croisille et al. (1978) suspect that basal cells serve as stem cells for the regeneration of the periodically exfoliating epithelial lining of the epididymal duct unit. Intraepithelial lymphocytes are also seen in the epithelium of the epididymal duct unit, as in other duct units, described above. Periductal Tissue. One or two layers of fibroblasts and up to ten concentric layers of smooth muscle cells constitute the periductal tissue which progressively thickens, cranio-caudally. The smooth muscle cells contain highly elongated, euchromatic, regular nuclei, surrounded by longitudinally orientated filaments, and organelles that are typical of smooth muscle cells (Fig. 2.33). This tissue, in the epididymal duct unit, is richly vascularized, being penetrated by a dense peritubular blood capillary network (Nakai et al. 1988; Aire 2000a), contrary to Tingari’s (1971) observation. The blood capillaries are fenestrated and close to the epithelium in the domestic fowl (Nakai et al. 1988) but not as close to the epithelium as in the mammalian epididymis (Abe et al. 1984). Encapsulated nerve endings are found between the blood capillaries and the ductal epithelium (Nakai et al. 1988), and both cholinergic and adrenergic nerve components have been demonstrated (Tingari, and Lake 1972a). The adrenergic nerve component is particularly closely associated with the epithelium, and ramifies abundantly in the walls of the ducts as well as the receptacle and papilla of the ductus deferens. The abundance of fine intrinsic nerves in the epididymal duct unit may be related to the higher level of development of smooth muscle cells in the boundary tissue of this duct unit than in the more cranial duct units, particularly the RT and efferent ducts. Therefore this unit may have a greater contractile force for onward movement of spermatozoa toward the cloaca than the more cranial duct units (Tingari and Lake 1972a).

'$ Reproductive Biology and Phylogeny of Birds

Fig. 2.32 Gallus domesticus. A basal cell rests on the basal lamina of the epithelium of the epididymal duct, and its nucleus (N) is oval in shape and surrounded by bundles of microfilaments (arrows) which may attach to the cell membrane (arrowheads). Organelles are sparse. Bar: 1 µm. Adapted from Aire, T. A. 2000 Anatomy Histology Embryology 29: 179-191, Figure 15. Reproduced with permission of Blackwell Publishing Ltd.

2.4

APPENDIX EPIDIDYMIDIS

The appendix epididymidis is made up of blind-ending tubules in the rooster (Budras and Sauer 1975a) and lies close to the adrenal gland, into which its cranial portion may be incorporated, for which reason, the appendix epididymidis is usually separated from the epididymis, on the removal of the testis from the body cavity. In the ostrich, the appendix epididymidis is very large and forms the cranial 40% of the entire length of the epididymis in the

Anatomy of the Testis and Male Reproductive Tract

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Fig. 2.33 Gallus domesticus. The epithelium of the ductus epididymidis rests on a relatively thick periductal layer of smooth muscle cells (S), displaying typical organelle features. Bar: 2 µm. Adapted from Aire, T. A. 2000 Anatomy Histology Embryology 29: 179-191, Figure 6. Reproduced with the permission of Blackwell Publishing Ltd.

sexually mature and active bird (Budras and Meir 1981). It is attached to the dorsal body wall by the mesepididymis, to the testis by the epiorchium and to the adrenal gland, cranially, by connective tissue as well as by some of its ductules and duct, whose free ends embed in the adrenal gland. The appendix epididymidis contains two vestigial duct components: (i) the ductus aberrans, that represents the straight, cranial blind end of the ductus epididymidis (Wolffian duct), and into which open (ii) the ductuli aberrantes. The latter are vestiges of the nephrons of the mesonephros that are farther away than others from the testis, and, instead, lie close to the adrenal gland. Ontogenetically, the ductuli aberrantes fail to make contact with the RT ducts because of the distance between them. They have lost their Bowman’s capsules, and become blind-ended (Budras and Sauer 1975b). The ductulus aberrans therefore constitutes the distal end of the nephron that normally gives rise to the DED and CD, developmentally (Budras and Sauer 1975b; Croissile et al. 1978). Other categories of ductules, are ductuli aberrantes which are connected to the RT but not to the epididymal duct, as well as tubuli paradidymidis, which are blind at both ends, and are seldom seen in histological sections along the length of the epididymis (Budras and Sauer 1975b).

'& Reproductive Biology and Phylogeny of Birds Transverse sections of these ducts exhibit regular outlines, simple cuboidal or low columnar epithelium, and a homogeneous, non-cellular luminal content (Tingari 1971), or small clumps of cellular debris in the lumen (Fig. 2.34). Budras and Sauer (1975b) have demonstrated, by histochemical and ultrastructural methods, moderate steroid hormone synthesis in the ductuli efferentes, ductus conjugens, and ductus epididymidis, and, in particular, high synthesis in the ductuli efferentes proximales segment of the efferent ductule, ductus aberrans and ductuli aberrantes in the rooster. The noduli epididymidis, in the Ostrich, is situated within the substance of the appendix epididymidis rostrally, and is derived from the swollen ends of the ductuli aberrantes. It is involved in steroidogenesis (Budras and Meier 1981).

2.5

THE PHALLUS

In most birds there is no organ that is homologous to the mammalian penis structurally. Closely related folds of tissue, derived from the ventral wall of the proctodeum of male birds, collectively form the phallus. The phallus is analogous to the penis in mammals because it is the organ of transfer of ejaculated spermatozoa from the male into the female reproductive tract. The phallus in most birds, for example, the domestic fowl, turkey and quail, is small and not intromittent. An engorgement of the vascular folds by lymph in the phallus causes an ‘erection’ that permits contact between this spermbearing organ and the slightly everted vagina of the female, during copulation.

Fig. 2.34 Coturnix japonica. A histological section of the rostral part of the epididymis, containing profiles of proximal efferent ductules (PED), ductus epididymidis (ED) and ductuli aberrantes (DA) which are round/oval and display cuboidal, ciliated epithelium and relatively empty lumina. Bar: 100 µm. Original.

Anatomy of the Testis and Male Reproductive Tract

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On the other hand, in Anseriforms and Ratites, the phallus is long, may be coiled, and intromittent (see King 1981 for an excellent review of the structure of avian phallus). This better developed phallus consists of paired fibrous bodies that constitute the bulk of the organ, a phallic sulcus that bears and transports the ejaculated semen, and an elastic vascular body which probably inclines the phallus cranioventrally, during erection. The avian phallus is in the subject of Chapter 3.

2.6

HISTOCHEMISTRY OF THE MALE REPRODUCTIVE ORGANS

A number of enzymes and other chemical compounds have been studied in specific regions of the male genital tract in mammals (Nicander 1954, 1957; Dawson and Rowlands 1959; Allen and Slater 1961; White et al. 1961; Mann 1964; Risley and Skrepetos 1964a,b; Stallcup and Roussel 1965; Blackshaw and Samisoni 1967). In order to ascertain homologies between parts of the male reproductive tract in mammals and birds, and to relate structure to function, it is necessary to evaluate a number of relevant enzymes and compounds in the male reproductive tract of birds. Glycoproteins, but not glycogen, are present in small amounts in the epithelium of the reproductive tract of the rooster (Tingari 1972), and Esponda and Bedford (1985) have shown that most of the sperm-binding secretions in the excurrent ducts of the rooster, save for one glycoprotein, are proteins. Four Wolffian duct proteins have thus been identified in the ducts of the rooster (Morris et al. 1987). Efferent duct non-ciliated cells seem capable of many metabolic functions, including the probable production of glutamic acid by mitochondrial-rich NC type I cells. Most of the amino acid content of the semen of the rooster is glutamate (Lake and McIndoe 1959). Lactic dehydrogenase (LDH) exhibits strong activity in the seminiferous tubules, but lipid reactivity and GDH, SDH and G-6-DH activities are very weakly expressed in the seminiferous tubules and RT (Tingari and Lake 1972c). The more distal excurrent ducts, however, have shown the presence of lipids (phopholipids, neutral lipids and free fatty acids) in both the epithelial cells and lumen (Tingari and Lake 1972c). Of all of the excurrent ducts, the PED exhibits the highest level of lipid content specifically in the NC type I cells. The function of the lipids may be related to estrogen production and/or utilization by efferent ducts of the rooster (Hess et al. 1997). Tingari (1973) concludes rightly that there exists a mechanism for the metabolism of steroids in the male fowl tract. The presence of both 3b- and 17b-hydroxysteroid dehydrogenases, and 3b-ol-steroid dehydrogenase (Budras and Sauer 1975b) in the various epithelia, and the presence of estrogen receptors (ER) in the germ cells and efferent ducts of the rooster (Kwon et al. 1995, 1997) confirm this assumption. Similar findings have been reported in the mouse (Janulis et al. 1996). It has been established that the absence of estrogen receptors in the efferent ducts of mice compromises considerably the indispensable fluid-resorptive ability of these ducts (Hess et al. 1997). The non-ciliated cells in efferent ducts appear to have a common function

 Reproductive Biology and Phylogeny of Birds in all species studied (Hess 2000). Thus, it is assumed that estrogen receptors in the homologous avian ducts have a similar function, perhaps, among others, of regulating fluid reabsorption in the efferent ducts of these animals, as in mammals. Strong cholinesterase activity is expressed in the epididymal duct, and this may be related to ionic movements, as found in homologous mammalian ducts, but it is absent in the seminiferous tubules, RT and efferent ducts (Tingari 1972). Recent reports show that the efferent ductule epithelium immunohistochemically expressed sodium-potassium ATPase (Na+, K+ ATPase), carbonic anhydrase II (CAII) and sodium hydrogen exchanger isoform 3 (NHE3), and that the connecting ductule and epididymal duct epithelia immunoexpressed Na+, K+ -ATPase and CA II (Bahr et al. 2006). Similarly, Zamboni et al. (2004) have shown that transmembrane water channel proteins (aquaporins –2, –3, and –9), that are responsible for water flow, are present in the epithelia of efferent ducts, collecting ducts and ductus epididymidis. Acid phosphatase activity is present only in the luminal macrophages in the RT, as well as in the dense bodies in NC type I cells of the PED, indicating that these bodies are lysosomal (Nakai et al. 1989a). The microvillous border, as also the lateral plasma membrane of the epididymal duct unit, shows intense acid phosphatase activity. Alkaline phosphatase activity is present only on the outer covering of the microvilli of the efferent ducts, and is completely absent in the epididymal duct unit. The functions of these enzymes in the male tract are only speculative. However, according to Aitken (1971), a significant concentration of acid phosphatase is characteristic of all spermstorage areas although its exact significance in storage is not clearly understood. Sperm are stored in the ductus deferens in birds, albeit for only short periods of time.

2.7

ACKNOWLEDGMENTS

The University of Pretoria kindly provided library support for this review, as well as research grants for new material contained in the text. I also acknowledge the assistance of Mrs. Wilma Olivier who made all the line diagrams. The assistance of both Dr. Peter Ozegbe and Dr. Wahab Kimaro in computerization and composition of the text and figures is gratefully acknowledged.

2.8 LITERATURE CITED Abe, K., Takano, H. and Ito, T. 1984. Microvasculature of the mouse epididymis, with special reference to fenestrated capillaries localized in the initial segment. The Anatomical Record 209: 209-218. Aire, T. A. 1979a. The epididymal region of the Japanese quail (Coturnix coturnix japonica). Acta Anatomica 103: 305-312. Aire, T. A. 1979b. Microstereological study of the avian epididymal region. Journal of Anatomy 129: 703-706. Aire, T. A. 1980. The ductuli efferentes of the epididymal region of birds. Journal of Anatomy 130: 707-723.

Anatomy of the Testis and Male Reproductive Tract



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Madekurozwa, M.-C., Chabvepi, T. S., Matema, S. and Teerds, K. J. 2002. Relationship between seasonal changes in spermatogenesis in the juvenile ostrich (Struthio camelus) and the presence of the LH receptor and 3b-hydroxysteroid dehydrogenase. Reproduction 123: 735-742. Malecki, I. A., Martin, G. B., O’Malley, P. J., Meyer, G. T., Talbot, R. T. and Sharp, P. J. 1998. Endocrine and testicular changes in a short-day seasonally breeding bird, the emu (Dromaius novahollandiae), in southwestern Australia. Animal Reproduction Science 53: 143-155. Man, S. Y., Clulow, J., Hansen, L. A. and Jones, R. C. 1997. Adrenal independence of fluid and electrolyte reabsorption in the ductuli efferentes testis of the rat. Experimental Physiology 82: 283-290. Mann, T. and Lutwak-Mann, C. 1981. Testis and testicular semen. Pp. 83-138. In Male Reproductive Function and Semen. Themes and Trends in Physiology, Biochemistry and Investigative Andrology. Springer-Verlag, Berlin, Heidelberg and New York. Marchand, C.-R. and Gomot, L. 1973. Étude histologique et cytologique des testicules et des voies génitales du canard de Barberie (Carina moschata L.) en activité sexuelle. Journées de Recherches Avicoles et Cunicoles A5: 127-134. Marchand, C.-R. 1973. Ultrastructure des cellules de Leydig et des cellules de Sertoli du testicule Canard de Barberie (Cairina moschata L.) en activite sexuelle. Comptes Rendus des Seances, Societe de Biologie 167: 933-1004. Marshall, A. J. 1961. Reproduction. Pp. 164-213. In A. J. Marshall (ed.), Biology and Comparative Physiology of Birds. Volume 2. Academic Press, New York. Marvan, Fr. 1969. Postnatal development of the male genital tract of the Gallus domesticus. Anatomischer Anzeiger 124: 443-462. Mehrotra, P. N. 1962. Cyclical changes in the epididymis of the goose, Anser melanotus. Quarterly Journal of Microscopical Science 103: 377-383. Mehrotra, P. N. 1964. On the microscopic anatomy of the epididymis of Anser melanotus L. Transactions of the American Microscopical Society 83: 456-460. Mezquita, B., Mezquita, C. and Mezquita, J. 1998. Marked differences between avian and mammalian testicular cells in the heat shock induction and polyadenylation of Hsp70 and ubiquitin transcripts. FEBS Letters 436: 382-386. Middleton, A. L. 1972. The structure and possible function of the seminal sac. Condor 74: 185-190. Mimura, H. 1928. On the bilateral asymmetry of testes in the domestic fowl. Japanese Journal of Zoology 2: 24. Møller, A. P. 1989. Ejaculate quality, size and sperm production in mammals. Functional Ecology 3: 91-96. Møller, A. P. 1991. Sperm competition, sperm depletion, paternal care, and relative testis size in birds. The American Naturalist 137: 882-906. Morales, C. and Hermo, L. 1983. Demonstration of fluid-phase endocytosis in epithelial cells of the male reproductive system by means of horseradish peroxidase-colloidal gold complex. Cell and Tissue Research 230: 503-510. Morales, C., Hermo, L and Clermont, Y. 1984. Endocytosis in epithelial cells lining the rete testis of the rat. The Anatomical Record 209: 185-195. Morita, J. 1966. Some observations on the fine structure of the human ductuli efferentes testis. Archivum Histologicum Japonicum 26: 341-365. Morris, S. A., Howarth Jr., B., Crim, J. W., Rodriguez de Cordoba, S., Esponda, P. and Bedford, J. M. 1987. Specificity of sperm-binding Wolffian duct proteins in the rooster and their persistence on spermatozoa in the female host glands. Journal of Experimental Zoology 242: 189-198.

 Reproductive Biology and Phylogeny of Birds Mulder, R. A. and Cockburn, A. 1993. Sperm competition and the reproductive anatomy of male Superb Fairy-Wrens. The Auk 110: 588-593. Munro, S. S. 1938. Functional changes in fowl sperm during their passage through the excurrent ducts of the male. Journal of Experimental Zoology 79: 71-92. Nakai, M., Hashimoto, Y., Kitagawa, H., Kon, Y. and Kudo, N. 1988. Microvasculature of the epididymis and ductus deferens of domestic fowls. Japanese Journal of Veterinary Science 50: 371-381. Nakai, M., Hashimoto, Y., Kitagawa, H., Kon, Y. and Kudo, N. 1989a. Histological study on seminal plasma absorption and spermiophagy in the epididymal region of domestic fowl. Poultry Science 68: 582-589. Nakai, M., Hashimoto, Y., Kitagawa, H., Kon, Y. and Sugimura, M. 1989b. Effects of ligation of the ductus deferens on the fowl epididymal region. Japanese Journal of Veterinary Science 51: 521-529. Nakai, M. and Nasu, T. 1991. Ultrastructural study on junctional complexes of the excurrent duct epithelia in the epididymal region in the fowl. Journal of Veterinary Medical Science 53: 677-681. Neaves, W. B. 1975. Leydig cells. Contraception 11: 571-606. Neaves, W. B. 1977. The blood-testis barrier. Pp. 125-162. In A. D. Johnson and Gomes, W. R. (eds.), The Testis. Volume IV. Academic Press, New York. Nicander, L. 1954. Glycogen secretion in the epididymis. Nature 174: 700-701. Nicander, L. 1957. On the regional histology and cytochemistry of the ductus epididymidis in rabbits. Acta morphologica neerlando-scandinavica 1: 99-118. Nicander, L. 1970. On the morphological evidence of secretion and absorption in the epididymis. Morphological Aspects of Andrology 1: 12-124. Nicander, L. and Glover, T. D. 1973. Regional histology and fine structure of the epididymal duct in the golden hamster (Mesocricetus auratus). Journal of Anatomy 114: 347-358. Nicholls, T. J. and Graham, G. P. 1972. Observation on the ultrastructure and differentiation of Leydig cells in the testis of the Japanese quail (Coturnix coturnix japonica). Biology of Reproduction 6: 179-192. Nickel, R., Schummer, A. and Seiferle, E. 1977. Urogenital system. Pp. 70-84. In Anatomy of the Domestic Birds. Verlag Paul Parey, Berlin and Hamburg. Nishida, T. 1964. Comparative and topographical anatomy of the fowl. XLII. Blood vascular system of the male reproductive organs. Japanese Journal of Veterinary Science 26: 211-221. Nonaka, S., Tanaka, Y., Okada, Y., Takeda, S., Harada, A., Kanai, Y., Kido, M. and Hirokawa, N. 1998. Randomization of left-right asymmetry due to loss of nodal cilia generating leftward flow of extraembryonic fluid in mice lacking KIF3B motor. Cell 95: 829-837. Odor, D. L. and Blandau, R. J. 1985. Observations on the solitary cilium of rabbit oviductal epithelium: its motility and ultrastructure. American Journal of Anatomy 174: 437-453. Olson, G. E. and Hinton, B. T. 1985. Regional differences in luminal fluid polypeptides of the rat testis and epididymis revealed by two-dimensional gel electrophoresis. Journal of Andrology 6: 20-34. Onagbesan, O. M., Safi, M., Decuypere, E. and Bruggeman, V. 2004. Development changes in inhibin a and inhibin/activin bA and bB mRNA levels in the gonads during post-hatch prepuberal development of male and female chickens. Moelcular Reproduction and Development 68: 319-326.

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Osman, D. I. 1978. On the ultrastructure of modified Sertoli cells in the terminal segment of seminiferous tubules in the boar. Journal of Anatomy 127: 603-613. Osman, D. I. 1980. The connection between the seminiferous tubules and the rete testis in the domestic fowl (Gallus domesticus)—morphological study. International Journal of Andrology 3: 177-187. Osman, D. I. and Plöen, L. 1978. The terminal segment of the seminiferous tubules and the blood-testis barrier before and after efferent ductule ligation in the rat. International Journal of Andrology 1: 235-249. Osman, D. I., Ekwall, H. and Plöen, L. 1980. Specialized cell contacts and the bloodtestis barrier in the seminiferous tubules of the domestic fowl (Gallus domesticus). International Journal of Andrology 3: 553-562. Palade, G. 1975. Intracellular aspects of the process of protein synthesis. Science 189: 347-358. Pease, D. C. 1956. Infolded basal plasma membranes found in epithelia noted for their water transport. Journal of Biophysical and Biochemical Cytology, Supplement 2: 203-208. Pellietier, R. M. 1990. A novel perspective: the occluding zonule encircles the apex of the Sertoli cell as observed in birds. American Journal of Anatomy 188: 87-108. Pfeiffer, D.C. and Vogl, A. W. 1993. Ectoplasmic (“Junctional”) specializations in Sertoli cells of the rooster and turtle: evolutionary implications. The Anatomical Record 235: 33-50. Plöen, L 1972. An electron microscope study of the immediate effects on spermateliosis of a short-term experimental cryptorchidism in the rabbit. Virchows Archives Abt B Zellpatologie 10: 293-309. Plöen, L. 1973a. An electron microscope study of the delayed effects on rabbit spermateliosis following experimental cryptorchidism for twenty-four hours. Virchows Archives Abt B Zellpatologie 14: 159-184. Plöen, L. 1973b. A light microscope study of the immediate and delayed effects on rabbit spermiogenesis following cryptorchidism for twenty-four hours. Virchows Archives Abt B Zellpatologie 14: 185-196. Pudney, J. and Fawcett, D. W. 1984. Seasonal changes in fine structure of the ductuli efferentes of the ground squirrel Citellus lateralis (Say). The Anatomical Record 188: 453-476. Ramos Jr., A. S. and Dym, M. 1977. Ultrastructure of the ductuli efferentes in monkeys. Biology of Reproduction 17: 339-349. Reid, B. L. and Cleland, K. W. 1957. The structure and function of the epididymis. I. The histology of the rat epididymis. Australian Journal of Zoology 5: 223-246. Riddle, O. 1918. Further observation on relative size and form of the right and left testes of pigeons in health and disease and as influenced by hybridity. The Anatomical Record 14: 283-334. Riddle, O. 1925. On the sexuality of the right ovary of birds. The Anatomical Record 30: 365-383. Risley, P. L. and Skrepetos, C. N. 1964a. Histochemistry of distribution of cholinesterases in the testis, epididymis and vas deferens of the rat. The Anatomical Record 148: 231-249. Risley, P. L. and Skrepetos, C. N. 1964b. Cholinesterase distribution in the rat epididymis and vas deferens after castration and sex hormone treatments. The Anatomical Record 150: 195-208. Robaire, B. and Hermo, L. 1988. Efferent ducts, epididymis, and vas deferens: Structure, functions, and their regulation. Pp. 999-1080. In E. Knobil and J. Neill (eds.), The Physiology of Reproduction. Raven Press, New York.



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Roosen-Runge, E. C. 1961. The rete testis in the albino rat: Its structure, development and morphological significance. Acta Anatomica 45: 1-30. Roosen-Runge, E. C. and Holstein, A. F. 1978. The human rete testis. Cell and Tissue Research 189: 409-433. Rothwell, B. 1973. The ultrastructure of Leydig cells in the testis of the domestic fowl. Journal of Anatomy 116: 245-253. Rothwell, B. 1975. Designation of the cellular component of the peritubular boundary tissue of the seminiferous tubule in the testis of the fowl (Gallus domesticus). British Poultry Science 16: 527-529. Rothwell, B. and Tingari, M. D. 1973. The ultrastructure of the boundary tissue of the seminiferous tubule in the testis of the domestic fowl (Gallus domesticus). Journal of Anatomy 114: 321-328. Rothwell, B. and Tingari, M. D. 1974. The ultrastructural differentiation of the boundary tissue of the seminiferous tubule in the testis of the domestic fowl. British Veterinary Journal 130: 587-592. Russel, L. D. 1996. Mammalian Leydig cell structure. Pp. 43-96. In A. H. Payne, M. P. Hardy and L. D. Russel (eds.), The Leydig Cell. Cache River Press, Vienna, Illinois. Salt, W. R. 1954. The structure of the cloacal protuberance. Auk 71: 64-73. Scheib, D. 1973. Les cellules secretices testiculaires du poussin de la caille japonaise: differenciation de leur ultrastructure et rapports avec leurs potentialities steroidogenes. Development, Growth and Differentiation 15: 315-328. Siller, W. G. and Hindle, R. M. 1969. The arterial blood supply to the kidney of the fowl. Journal of Anatomy 104: 117-135. Sinowatz, F., Wrobel, K.-H., Sinowatz, S. and Kugler, P. 1979. Ultrastructural evidence for phagocytosis of spermatozoa in the bovine rete testis and testicular straight tubules. Journal of Reproduction and Fertility 57: 1-4. Skinner, M. K. 1987. Cell-cell interactions in the testis. Annals of New York Academy of Sciences 513: 158-171. Skinner, M. K., Norton, J. N., Mullaney, B. P., Rosselli, M., Whaley, P. D. and Anthony, C. T. 1991. Cell-cell interactions and the regulation of testis function. Annals of New York Academy of Sciences 637: 354-363. Soley, J. T. 1992. A histological study of spermatogenesis in the ostrich (Struthio camelus). Ph.D. thesis, University of Pretoria, Pretoria, 187 pp. Sprando, R. L. and Russel, L. D. 1987. A comparative study of Sertoli cell cytoplasmic specializations in selected non-mammalian vertebrates. Tissue and Cell 19: 479493. Stallcup. O. T. and Roussel, J. D. 1965. Development of lactic acid dehydrogenase enzyme system in the testis and epididymis of young dairy bulls. Journal of Dairy Science 48: 1511-1516. Stanley, A. J. and Witschi, E 1940. Germ cell migration in relation to asymmetry in the sex glands of hawks. The Anatomical Record 76: 329-342. Stefanini, M. A., Orsi, A. M., Grégorio, E. A., Viotto, M. J. S. and Baraldi-Antoni, S. M. 1999. Morphologic study of the efferent ductules of the pigeon (Columba livia). Journal of Morphology 242: 247-255. Stoffel, M. H. and Friess, A. E. 1994. Morphological characteristics of boar efferent ductules and epididymal duct. Microscopy Research and Techniques 29: 411-431. Suzuki, F. and Nagano, T. 1978. Regional differentiation of cell junctions in the excurrent duct epithelium of the rat testis as revealed by freeze-fracture. The Anatomical Record 191: 503-520.

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Tingari, M. D. 1971. On the structure of the epididymal region and ductus deferens of the domestic fowl (Gallus domesticus). Journal of Anatomy 109: 423-435. Tingari, M. D. 1972. The fine structure of the epithelial lining of the excurrent duct system of the testis of the domestic fowl (Gallus domesticus). Quarterly Journal of Experimental Physiology 57: 271-295. Tingari, M. D. 1973. Histochemical localization of 3b- and 17b-hydroxysteroid dehydrogenases in the male reproductive tract of the domestic fowl (Gallus domesticus). Histochemical Journal 5: 57-65. Tingari, M. D. and Lake, P. E. 1972a. The intrinsic innervation of the reproductive tract of the male fowl (Gallus domesticus). A histochemical and fine structural study. Journal of Anatomy 112: 257-271. Tingari, M. D. and Lake, P. E. 1972b. Ultrastructural evidence for resorption of spermatozoa and testicular fluid in the excurrent ducts of the testis of the domestic fowl, Gallus domesticus. Journal of Reproduction and Fertility 31: 373381. Tingari, M. D. and Lake, P. E. 1972c. Histochemical localization of glycogen, mucopolysaccharides, lipids, some oxidative enzymes and cholinesterases in the reproductive tract of the male fowl (Gallus domesticus). Journal of Anatomy 112: 273-287. Traciuc, P. E. 1967. L’anatomie microscopique de l’épididyme chez Sterna hirundo L. Anatomischer Anzeiger 121: 382-386. Traciuc, E. 1969. La structurede l’epididyme de Coloeus monodula (aves, Corvidae). Anatomische Anzeiger 125: 49-67. Tuck, R. R., Setchell, B. P., Waites, G. M. H. and Young, J. A. 1970. The composition of fluid collected by micropuncture and catheterization from the seminiferous tubules and rete testis of rats. European Journal of Physiology 318: 225-243. Veeramachaneni, D. N. R. and Amann, R. P. 1991. Endocytosis of androgen-binding protein, clusterin, and transferrin in the efferent ducts and epididmis of the ram. Journal of Andrology 12: 288-294. Weaker, F. J. 1977. The fine structure of the interstitial tissue of the testis of the ninebanded armadillo. The Anatomical Record 187: 11-27. White, I. G., Wallace, J. C., Wales, R. G. and Scott, T. W. 1961. The occurrence and metabolism of glycerylphosphorycholine in semen and the genital tract. Proceedings of the IVth International Congress of Animal Reproduction and Artificial Insemination, The Hague, 266-269. Williams, D. D. 1958. A histological study of the effects of subnormal temperature of the testis of the fowl. The Anatomical Record 130: 225-242. Witschi, E. 1935. The origin of asymmetry in the reproductive system of birds. American Journal of Anatomy 56: 119-141. Wolfson, A. 1954. Sperm storage at lower than body temperature outside the body cavity in some passerine birds. Science 120: 68-71. Zamboni, L., Akuffo, V. and Bakst, M.R. 2004. Aquaporins are observed in the duct epithelia of the epididymal region of the Large White turkey. Poultry Science 83: 1917-1920. Zhu, L. J., Zong, S. D., Phillips, D. M., Moo-Young, A. J. and Bardin, C. W. 1997. Changes in the distribution of intermediate filaments in the rat Sertoli cells during the seminiferous epithelium cycle and postnatal development. The Anatomical Record 248: 391-405.

n n

CHAPTER

3

Anatomy and Evolution of Copulatory Structures Robert Montgomerie1 and James Briskie2

3.1

INTRODUCTION

Unlike most other animal Classes, Aves (birds) is a taxon in which males of some species possess an intromittent organ (IO), whereas males of other species do not (King 1981a). Indeed, birds are virtually unique among internal fertilizers that most species lack an IO. Thus, in birds at least, the IO is not necessary for internal fertilization. This raises the question, then, whether the avian IO has evolved as a primary sexual trait simply for the delivery of sperm, as is sometimes assumed for other taxa, or as a secondary sexual trait (Eberhard 1990; Briskie and Montgomerie 1997). Most of the scant literature on avian IOs has focused on the seemingly odd presence of IOs in a few orders, but it is in fact their absence in so many species that is the evolutionary puzzle. Despite this interesting question about the absence of IOs in birds, relatively little is known about the anatomy, physiology, and evolution of structures that facilitate copulation in this taxon. There was quite a bit of interest in the phallus of ratites in the 19th century with J. Müller’s (1836) anatomical study being the most comprehensive—King (1981a) provides a useful review of this early work. Later, Eckhard (1876), R. Müller (1908) and Liebe (1914) showed that the Mallard (Anas platyrhynchos) phallus is erected by a lymphatic, rather than blood-vascular mechanism as had previously been thought. Gerhardt (1933) and other early workers also noted that there were two types of true phalluses in birds—those with and without a blind cavity— and there was a smattering of other studies on the phalluses of wild birds in the early part of the 20th century. Beyond that early work, only the non1 2

Department of Biology, Queen’s University, Kingston, ON K7L 3N6, Canada School of Biological Sciences, University of Canterbury, Christchurch, New Zealand

$ Reproductive Biology and Phylogeny of Birds intromittent phalluses of the domestic chicken (Gallus gallus) and turkey (Meleagris gallopavo) received much anatomical or physiological study (King 1981a). However, during the past decade or so, there has been renewed interest in the avian IO, with new hypotheses to explain the evolutionary patterns of occurrence (Briskie and Montgomerie 1997; Wesolowski 1999; Briskie and Montgomerie 2001; Wesolowski 2001), comparative analyses of anatomical structure (Coker et al. 2002), detailed anatomical work using modern techniques (Oliveria and Mahecha 2000; Oliveria et al. 2003, 2004), and some excellent natural history on the most bizarre copulatory structures yet discovered in birds (Birkhead et al. 1993; Mulder and Cockburn 1993; Birkhead and Hoi 1994; Wilkinson and Birkhead 1995; Winterbottom et al. 1999, 2001; McCracken 2000; McCracken et al. 2001). In this chapter we present an overview on what is known about the anatomy, histology, and physiology of the avian phallus—both intromittent (phallus protrudens) and non-intromittent (phallus nonprotrudens) forms—as well as other copulatory structures involved directly in the transfer of sperm from male to female. King’s (1981a) comprehensive review is our point of departure for this material and should be referred to by the reader needing more detail on work done up to 1980. We also discuss the evolutionary history of IOs in birds in relation both to phylogeny and to the various selective pressures that may have favored the evolutionary loss of the IO and the appearance of phallus-like copulatory structures in some lineages. Our goal in this chapter is to provide a framework for understanding the evolution of all copulatory structures in birds in the context of both the evolutionary history of birds and the influence of both natural and sexual selection on the modification and loss of phalluses in some lineages.

3.2

EVOLUTIONARY HISTORY OF COPULATORY STRUCTURES

The various kinds of copulatory structures found in modern birds (Neornithes) can be mapped onto the most recent avian phylogenies (e.g., Figs. 3.1-3.3) to illustrate their probable evolutionary history. In birds, a true phallus (both intromittent and non-intromittent) is found only in the orders Struthioniformes (ostrich, rheas, emu, kiwis and cassowaries), Tinamiformes (tinamous), Anseriformes (waterfowl and screamers), and Galliformes (pheasants, grouse, megapodes and currasows). The true phallus is clearly homologous to the phallus of crocodiles, turtles and tortoises (King 1981a), their closest living relatives (Fig. 3.1). A phallus-like structure (or pseudophallus) used in copulation is known from only three other bird genera (see below); in only one of these is it intromittent, and in none is it synapomorphic with the crocodile phallus. Male birds in all remaining avian taxa apparently have no phallus, but instead copulate solely by cloacal apposition, usually for only a few seconds (see Volume 6B, Chapter 6). Problems in rooting the avian tree with respect to the Crocodylia has made placement of the sister taxa Galliformes and Anseriformes controversial with

Anatomy and Evolution of Copulatory Structures

%

Fig. 3.1 Phylogenetic relationship of birds to crocodiles, turtles, lizards, snakes, and some extinct dinosaurs. Modified after Padian, K. and Horner, J. R. 2002. Trends in Ecology and Evolution 17: 120-124, Fig. 3.

respect to the Palaeognathae and Neognathae (Cracraft et al. 2004). Thus to provide some context for the discussion of phallus evolution in birds, we have mapped the avian phallus and other copulatory organs onto two different phylogenies that have recent support (Figs. 3.2 and 3.3). These two phylogenetic hypotheses are based on both molecular and morphological data and are different in many respects from earlier phylogenies based on morphology alone. The comprehensive phylogeny (‘Tapestry’; see Chapter 1) of bird evolution proposed by Sibley and Ahlquist (1990), based on DNA-DNA hybridization, grouped the ratites (ostrich, rheas, and tinamous) into one clade (Palaeognathae), and the Galliformes and Anseriformes into a separate clade

& Reproductive Biology and Phylogeny of Birds

Fig. 3.2 Summary of the Sibley-Ahlquist ‘Tapestry’ of bird phylogeny, based on DNA hybridization, illustrating the evolution of the avian phallus. The Megapodiidae are marked as equivocal because IOs have been recorded in some species but not others, based on very sketchy evidence (see text). Modified after Cracraft, J., et al. 2004. Pp 468-489. In Cracraft, J. and Donoghue, M. J. (eds), Assembling the Tree of Life. Oxford University Press, New York, Fig. 27.2.

Anatomy and Evolution of Copulatory Structures

'

Fig. 3.3 Summary of a recent consensus phylogeny of birds, based on both molecular and morphological evidence, illustrating the evolution of the avian phallus. The Megapodiidae are marked as equivocal because IOs have been recorded in some species but not others, based on very sketchy evidence (see text). Modified after Cracraft, J., et al. 2004. Pp 468-489. In Cracraft, J. and Donoghue, M. J. (eds), Assembling the Tree of Life. Oxford University Press, New York, Fig. 27.10.

  Reproductive Biology and Phylogeny of Birds (Galloanserae) that is the sister clade to the Palaeognathae (Fig. 3.2). According to this phylogenetic hypothesis, the clade comprising the Palaeognathae and the Galloanserae is separate from the clade comprising all other birds (Neoaves). Though the Sibley-Ahlquist Tapestry was radically different from accepted phylogenies at the time, it received widespread support because it was based on DNA analyses, and made sense in many respects. Problems with both the use of DNA-DNA hybridization and the interpretation of data, however, encouraged other researchers to employ a large number of other kinds of molecular and phylogenetic analyses that continue to refine our understanding of the relationships among and within the clades that Sibley and Ahlquist (1990) proposed. Probably the best consensus phylogeny currently available, based on nuclear genes as well as other traits, agrees with the Sibley-Ahlquist Tapestry in many respects, but does not isolate the Galloanserae and Palaeognathae in a separate clade (Fig. 3.3). The evolution of avian phalluses maps onto both of these general molecular phylogenies in a similar fashion (Figs. 3.2 and 3.3), with the true phallus being monophyletic in both cases. The intromittent true phallus is clearly the ancestral type and is shared by all species in the Palaeognathae and several families of the Galloanserae; the non-intromittent true phallus, on the other hand, has probably arisen only once, in the ancestor to the clade comprising the Phasianidae, Odontophoridae, and Numididae, though possibly also in the Megapodiidae (see below). Several other copulatory structures that have so far been discovered in birds are all in the Neoaves, but only in the Psittaciformes (genus Coracopsis) has a copulatory structure evolved with an intromittent function. In all other species in which the genitalia of males appear to have been modified for copulation, none is known to have a structure that is inserted into the female’s cloaca.

3.3 3.3.1

FORM AND FUNCTION Ancestral Phalluses

The true phalluses of birds all have traits similar to the phalluses of their closest living relatives—the crocodiles, turtles, and tortoises (Fig. 3.4)—and appear to be monophyletic (Figs. 3.1-3.3). Given our current understanding of the evolutionary relationships among birds and dinosaurs (Fig. 3.1), it seems quite likely that theropod dinosaurs also possessed an intromittent phallus (Larson and Frey 1992) not unlike that of crocodiles, chelonians, and the intromittent true phalluses of birds. The phalluses of male chelonians (turtles and tortoises) comprise a thickening along the midline of the ventral wall of the proctodeum (Wood Jones 1915). In most species, the caudal portion of the phallus is separated from the wall of the proctodeum so that most of the phallus is free (Fig. 3.4A), and is inserted into the female during copulation. The phallus is made up of two fibroelastic bodies (fibrous tissue with large vascular spaces), which in

Anatomy and Evolution of Copulatory Structures

 

Fig. 3.4 Typical intromittent true phalluses of (A, B) turtles and (C) crocodiles showing the location of the ejaculatory groove (phallic sulcus). B. Cross section of A. Not drawn to scale, nor were scale bars shown on the original drawings. Modified after King, A.S. 1981a. Pp 107-147. In King, A. S. and McLelland, J. (eds), Form and Function in Birds. Academic Press, London, Fig. 3.2.

most species are fused into a single structure of erectile tissue (Fig. 3.4). The fibroelastic bodies are separated by a median ejaculatory groove, called the phallic sulcus, that extends from the opening of the urogenital sinus nearly to the tip of the phallus. As the phallus becomes erect, the fibroelastic bodies swell such that this groove becomes a closed channel along which the semen travels (Wood Jones 1915). The phalluses of crocodilians are clearly homologous to those of chelonians, but the free part is longer and projects more prominently from the proctodeal wall when it is not erect, the shape tends to be more cylindrical (Fig. 3.4C), and the glans may be more complex (Gerhardt 1933). In addition, the crocodilian phallus is distinctly curved, particularly when it is erect, compared to the relatively straight phallus of turtles and tortoises (Fig. 3.4). In chelonians and crocodilians, the mechanism of erection appears to be blood vascular, though King (1981a) hints that this needs to be better documented. In both taxa, females also have a phallus (i.e., clitoris) that is very similar to that of the male, but is much smaller and cannot be extruded from the cloaca.

3.3.2

Bird Phalluses

King (1981a) and earlier workers classified true avian phalluses into intromittent and non-intromittent types based on their gross anatomy, and further divided the intromittent phallus into two types based on the presence (Type B) or absence (Type A) of a blind cavity. Type B phalluses are similar in



Reproductive Biology and Phylogeny of Birds

shape to Type A but have a blind tubular cavity, like the invaginated finger of a glove, when flaccid, and tend to be twisted in a spiral from the base to the tip when erect. This categorization, based on the presence/absence of a blind cavity, appeared to make some evolutionary (phylogenetic) sense at one time, but more recent work on the anatomy and histology of tinamou phalluses (Oliveira and Mahecha 2000), as well as the best current phylogenies of birds (Fig. 3.3), suggest that the blind cavity may be an adaptive trait, possibly with some erectile function. Thus, the distinction between Type A and B phalluses does not appear to be particularly useful from a phylogenetic perspective. Like the phalluses of crocodilians and chelonians (Fig. 3.4), all intromittent true phalluses in birds are composed of fibrous, erectile tissue that arises from the ventral wall of the proctodeum and have a ventral sulcus (ejaculatory groove) along which the semen travels during ejaculation (e.g., Figs. 3.5-3.10). [In birds, the cloaca has three compartments separated from each other by folds: (1) the proctodeum, nearest the vent, (2) the urodeum, a narrow zone that the urogenital ducts empty into (except in tinamous and rheas; Oliveira et al. 2003), and (3) the coprodeum, into which the rectum empties (King 1981b).] When the phallus is erect, the lips of the ejaculatory groove also become engorged, sealing the groove and thus preventing the spillage of semen. Intromittent phalluses all have a fixed base of fibrous tissue and a free conical, or tubular portion (composed of two fused fibrolymphatic bodies) that is everted during erection and copulation. Such phalluses usually curve towards the male’s left, and thus during copulation will most likely deposit sperm into the female’s left oviduct, which in birds is the functional one (King 1981a). The fixed base of the phallus has lymphatic spaces that fill and become dilated during erection (Oliveira and Mahecha 2000); vascular bodies in the floor of the urodeum provide the lymph that engorges the phallus,

Fig. 3.5 Intromittent true phallus of the male Spotted tinamou in its erect state: A. Caudal end. B. Left side view. Modified from Oliveira, C. A. and Mahecha, G. A. B. 2000. Annals of Anatomy 182: 161-169, Figs. 18 and 19.

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possibly through the action of the cloacal sphincter muscle. The base of the phallus also has glandular tissue that secretes mucous that lubricates the phallus and facilitates both eversion and, presumably, copulation (Komárek and Marvan 1969). Below we review the taxonomic distribution of phalluses among extant birds; classification and species numbers are taken from Dickinson (2003).

3.3.2.1

Palaeognathae

Males of all species of Palaeognathae studied so far have an IO that is similar to the crocodilian phallus (Fig. 3.4C), characterized a conical base of fibrous tissue, and a bend to the bird’s left when erect due to the two fused fibrolymphatic bodies being laterally asymmetrical (e.g., Figs. 3.5-3.8). In all species studied so far, the female also has a phallus that is much smaller than that of the male and does not extrude from the cloaca (e.g., Fig. 3.6B). Tinamidae (tinamous, 47 species). Tinamous were classified by King (1981a) as having a Type A phallus (i.e., lacking a blind cavity), based on work by Müller (1836) and Gerhardt (1933) on Crypturellus cinnamomeus. However, a recent anatomical and histological study of the Spotted tinamou (Nothura maculosa) clearly documents the presence of a blind cavity (Fig. 3.5A; Oliveira and Mahecha 2000) like that in the Anseriformes, rheas, emus, and cassowaries. It is quite possible that other species of tinamou possess such a cavity but there is too little information available (even in Gerhardt 1933) to be certain. The phallus of the Spotted tinamou is the best studied in this family, based on a sample of 26 males captured at different times of the year (Oliveira and

Fig. 3.6 North Island kiwi (Apteryx australis mantelli): A. Intromittent true phallus of the male. B. Vestigial phallus (clitoris) of the female. Modified after Caithness, T. A. 1971. International Zoo Yearbook 11: 206-208, Figs. 2 and 3.

 " Reproductive Biology and Phylogeny of Birds Mahecha 2000). To describe the morphology of the phallus, Oliveira and Mahecha (2000) conducted modern histological and anatomical studies of birds with the phallus both flaccid and erect (the latter induced by cloacal massage). In this species, the phallus is composed of a fixed, fibrous, conical base attached to the floor of the proctodeum and a tubular portion with a blind-ended tube at the tip (Fig. 3.5A). When the phallus is not erect, it lies entirely within a phallic pouch in the floor of the proctodeum. The erect phallus is distinctly coiled in a spiral (Fig. 3.5B) and extends about 3 cm out of the male’s vent during copulation, directed towards the male’s left. The fixed base of the phallus of both Spotted and Red-winged tinamous (Rhynchotus rufescens) also has intraepithelial plasma cells that increase at least 8-fold in number during the breeding season (Oliveira et al. 2003). These cells contain granular material indicative of immunoglobulin accumulation that might be important for an immune response to protect the phallus tissue from infection resulting from sexually transmitted diseases. Alternatively, secretions from these cells may be added to the seminal fluid and thus protect sperm within the female’s reproductive tract (Oliveira et al. 2003). Apterygidae (kiwis, 3 species). Except that it is asymmetrical and curves to the male’s left, the phallus of male kiwis (Fig. 3.6A) is very similar to that of crocodilians and chelonians (Fig. 3.4). The phallus of the female (i.e., clitoris) is very small and cannot be everted from the cloaca (Fig. 3.6B), making this a relatively easy trait for sexing these externally monomorphic birds (Caithness 1971). Casuariidae (cassowaries, 3 species). Phalluses have been observed in the Southern (Casuarius casuarius) and Dwarf Cassowaries (C. bennetti), and in the latter were described as being indistinguishable from the phalluses of rheas and emus (Gerhardt 1933). Dromaiidae (emus, 3 species). Emus are reported to have the same phallus structure as rheas (Müller 1836, Gerhardt 1933). Struthionidae (ostrich, 1 species). The large, bright red phallus of the male Ostrich (Struthio camelus) is up to 20 cm long when flaccid and extrudes 40 cm or more out of the male’s cloaca when erect (Fig. 3.7A; Gerhardt 1933). Even in its flaccid state, it has to be partly protruded from the cloaca to allow defecation as it takes up so much space in the cloacal lumen that it blocks the opening of the ureter. At rest, it lies within a wide pocket in the ventral wall of the proctodeum. As in other paleognaths, the walls of the ejaculatory groove (Fig. 3.7B) in the Ostrich are composed of erectile tissue that fills with lymph during erection, and presumably seals the lip of the groove to form a tube down which the semen passes. During erection a pair of retractor muscles (m. lecator phalli) extrude the flaccid phallus from its pocket (Fig. 3.7A), followed by the vascular bodies filling the phallus with lymph. King (1981a), however, notes that the literature on this lymphatic mechanism (and the tissues involved) is old, equivocal, and biased. It is unlikely that the mechanisms of erection in

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Fig. 3.7 Intromittent true phallus of the male Ostrich: A. Erect phallus extruded from vent, with dissection showing musculature. B. Cross section of A (at about the dotted line) showing asymmetrical sizes of the left and right fibrolymphatic bodies. Not drawn to scale (especially since the erect phallus should be about 40 cm long, or about 10 times the width of the vent), nor were scale bars shown on the original drawings. Modified after King, A. S. 1981a. Pp 107-147. In King, A. S. and McLelland, J. (eds), Form and Function in Birds, vol. 2. Academic Press, London, Figs. 3.3b and 3.4d.

ostriches is different from that in the other paleognaths but this deserves some study using modern techniques Rheidae (rheas, 2 species). Müller’s (1836) description of the phallus of the Greater rhea (Rhea americana) is still the best and most detailed (Fig. 3.8). The other species of rhea has not been studied but its phallus should have essentially the same structure. In the Greater rhea, the resting phallus has an orifice at the tip that leads to a blind tube or cavity, as in both the Spotted tinamou and the Anseriformes. When the phallus is erect, about half of this blind cavity is evaginated. The proximal end of this cavity is continuous with the phallic sulcus, and when the phallus is at rest (i.e., flaccid) part of the phallic sulcus is actually pulled into the cavity. Unlike the other paleognaths studied so far, the fibrous bodies at the base of the rhea’s phallus are spirally intertwined, and the free tubular portion of the phallus continues in a slight left-turning spiral throughout its length (Fig. 3.8). In addition a band of elastic fibers (lig. elasticum phalli) runs along the entire length of the everted phallus and probably retracts the phallus during detumescence (Müller 1836; Gerhardt 1933).

 $ Reproductive Biology and Phylogeny of Birds

Fig. 3.8 Intromittent true phallus of the male Rhea. Not drawn to scale, nor was there a scale bar shown on the original drawing. Modified after Briskie, J. V. and Montgomerie, R. 1997. Journal of Avian Biology 28: 73-86, Fig. 1a, who redrew this from King, A. S. 1981a. Pp 107-147. In King, A. S. and McLelland, J. (eds), Form and Function in Birds, vol. 2. Academic Press, London, Fig. 3.3d.

3.3.2.2

Neognathae

Galloanserae (452 species) The Galliformes and Anseriformes comprise the Galloanserae (Sibley and Ahlquist 1990), recognizing that these two avian orders are sister taxa (Fig. 3.3). Like the paleognaths, both of these orders possess a true phallus but only in the Anseriformes, and probably in the Cracidae and Megapodidae, is it intromittent. Thus the common ancestor of these two orders most likely had an IO but the size of this phallus became reduced and no longer involved in intromission in the galliform lineage leading to the Phasianide, Numididae, and Odontophoridae, and possibly in the Megapodiidae (Fig. 3.3). Anseriformes (162 species) Anatidae (ducks, geese, swans; 158 species). The family Anatidae is distinguished by having (i) the species with the longest (relative to body size) IO of any bird (McCracken 2000), (ii) the species with the best-studied IO, and (iii) by far the largest number of species for which there are quantitative data on IO size and morphology (Coker et al. 2002). Not only is the Anatidae the most speciose family of birds with IOs, but it displays a wide range of mating systems and male reproductive tactics and thus provides a useful model for studying the evolution of IOs in relation to sexual selection. Coker et al. (2002) have made an excellent start at this but there is still much to be done. The Anatidae are also relatively easy to keep and study in captivity, even while breeding (Johnsgard 1978), and so would lend themselves well to both critical

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experiments about IO function in relation to structure and detailed anatomical/physiological study during the critical period when males copulate. The phallus of the male Mallard is probably better studied than that of any other bird, save possibly the non-intromittent phallus of the domestic chicken. At rest, the Mallard’s phallus lies coiled within a thin peritoneal sac in the ventral wall of the proctodeum; when erect, the base of the phallus fills the male’s vent and the phallus extrudes 4 cm out of the cloaca in wild birds (but 8 cm in domesticated stock; Rautenfeld et al. 1974). The right fibrolymphatic body is much larger that the left and the phallus twists 3-4 turns in a left spiral from the base to the tip (Fig. 3.9). These two bodies are fused and thus continuous with each other but are separated at the surface by a deep ejaculatory groove (phallic sulcus) along which semen travels during ejaculation. The surface of the phallus is relatively smooth at the base but within a half turn of the spiral the surface of the fibrolymphatic bodies becomes cornified with rough transverse ridges about 2 mm apart. The tip of the IO has a small opening into a blind cavity, typical of Type B avian phalluses (Fig. 3.9). The Mallard phallus becomes erect by filling with lymph from two small (1 ¥ 4 cm) vascular bodies in the cloacal wall near the base of the phallus. Peristaltic contractions of the cloacal sphincter increase the flow of lymph to the fibrolymphatic bodies (Guzsal 1974). The swelling of these bodies seals the lips of the phallic sulcus, converting it into a closed tube such that semen can leave the phallus only at the tip. The phallus detumesces when the cloacal sphincter relaxes and lymph flows out of the phallus into general circulation.

Fig. 3.9 Intromittent true phallus of the male Mallard as viewed from the left side when fully erect. No scale bar shown on the original drawing. Note that the right fibrolymphatic body is much smaller than the left, and that the ejaculatory groove (phallic sulcus) is between them. Modified after Briskie, J. V. and Montgomerie, R. 1997. Journal of Avian Biology 28: 73-86, Fig. 1a, who redrew this from King, A. S. 1981a. Pp 107-147. In King, A. S. and McLelland, J. (eds), Form and Function in Birds, vol. 2. Academic Press, London, Fig. 3.6.

 & Reproductive Biology and Phylogeny of Birds This process is assisted by contractions of two pairs of muscles as well as by elastic fibers in the phallus that were stretched during erection. Detumescence begins with the immediate invagination of the base of the phallus, then over about 30 s the entire phallus is retracted into the cloaca by the action of the retractor muscles and the elastic fibers. Finally, over a 2-4 min period, the whole flaccid phallus is folded back inside the peritoneal sac, aided by the antiperistaltic action of the cloacal sphincter as well as by mucous that is secreted from the glandular base of the phallus. Coker et al. (2002) studied the IOs of 54 species of Anatidae for which they had detailed, scaled drawings of IOs (one per species) made from formalinpreserved museum specimens of males taken during the breeding season. As they did not work with fresh or live specimens, it is unknown whether the species that they studied have phalluses that are similar in structure to that of the Mallard. From each drawing, they measured the flaccid length of the IO, estimated its circumference, and quantified the numbers and heights of knobs and ridges (Fig. 3.10, Plate 3.1) on its surface. In the species studied, flaccid IO length varied from 1.25 cm in the Red-breasted goose (Branta ruficollis) to an incredible 28.5 cm in the Australian blue-billed duck (Oxyura australis). IO length was not related to body length (r = –0.11, P = 0.36) in this sample of species, suggesting that variation in size might be adaptive. Indeed, the largest geese and swans had among the smallest IOs (e.g., 2.4 cm in the Canada goose, Branta canadensis, and 2.3 cm in the Tundra swan, Cygnus columbianus), whereas the relatively small stiff-tailed ducks (subfamily Oxyurinae) had among the largest (e.g., 23.6 cm in the Ruddy duck, O. jamaicensis). Even between closely-related species of similar size, the IO can be quite different in

Fig. 3.10 Intromittent true phallus of the North American ruddy duck, showing knobs and ridges (not to scale). The base of the phallus is to the left and the view is from the right side of the phallus. Modified after Coker, C. R. et al. 2002. Auk 119: 403-413, Fig. 1.

Anatomy and Evolution of Copulatory Structures

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size (e.g., 9.2 cm in the Common eider, Somateria mollissima, versus 15.0 cm in the King eider, S. spectabilis; 9.2 cm in the White-faced whistling duck, Dendrocygnus viduata, versus 18.8 cm in the Black-bellied whistling duck, D. autumnalis). There was also considerable variation in both the number (density) and size of knobs and ridges on the surface of the IOs, with the density of ridges strongly negatively correlated with IO circumference (r = – 0.86, P <0.05). Thus larger IOs tended to have more knobs than ridges (Fig. 3.11). The stiff-tailed ducks have long been known to have large IOs especially in relation to their body size. Indeed, the IO of the Australian blue-billed duck has been described as being so large that, after copulation, the male rolls on his back and prods his long, detumescing IO back into his cloaca with his bill (Marchant and Higgins 1990). Males of both the Australian blue-billed duck and the North American ruddy duck apparently often preen their IO after copulation (McCracken 2000). In this same subfamily, McCracken et al. (2001) recently discovered a male Argentine lake duck (O. vittata) with an IO that was 42.5 cm long in its everted, flaccid state, fully as long as the duck itself (Plate

Fig. 3.11 Results of Principal Component Analysis on IO sizes and surface structures (see Fig. 3.10) in the family Anatidae, grouped according to the presumed frequency of forced extrapair copulations (FEPCs). Each data point represents one species; outlines encompass each mating system type, with the range of monogamous birds shaded to improve clarity. Modified after Coker, C. R. et al. 2002. Auk 119: 403-413, Fig. 2.

CMYK

CMYK

CMYK

! Reproductive Biology and Phylogeny of Birds

Plate 3.1 A,B Intromittent true phallus of the male Argentine lake duck: A. Extending from male, pulled out to its full extent but not erect. B. Showing knobs Plate 3.1 Contd. ...

CMYK

Anatomy and Evolution of Copulatory Structures

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3.1A)! McCracken (2000) had previously described the IO of this species as averaging 22.3 cm (range 19.0-24.5 cm, n = 4 males), thus suggesting that this one male had a particularly long IO. Despite its length, the width of the IO averaged only 6.5 mm (range 4.5-8 mm) at its base. In this species, the inverted, detumescent IO lies entirely within a blind pouch in the ventral wall of the cloaca, as in other anatids. The base of the IO is covered with a dense array of large, sharp, black-tipped white spines, most dense on the ventral side of the IO, and pointing outward and backward toward the cloaca when everted (Plate 3.1B). Thus copulation may proceed by the male everting the first 1-2 cm of his IO and inserting this into the female where the spines would clasp the female’s cloaca while the male everts the rest of his IO inside the female (McCracken 2000). Spines also occur along a series of spiral grooves extending along the full length of the everted IO (Plate 3.1B). Anseranatidae (magpie goose; 1 species). To the best of our knowledge, the phallus of the Magpie goose (Anseranas semipalmata) has never been studied, but there is no reason to expect that it differs much in structure from the typical anatid phallus. Anhimidae (screamers; 3 species). The phallus of screamers has never been studied, but looks superficially like that of the Mallard (R. M., unpublished data). Galliformes (290 species) Megapodiidae (scrub fowl and brush turkeys; 22 species). The Australian brush-turkey (Alectura lathami), the Malleefowl (Leipoa ocellata), and species in the genus Aepypodius are all reported to have an intromittent phallus, whereas at least some other megapodes in the genus Megapodius apparently do not (Brom and Dekker 1992; Jones et al. 1995), though no details are available on the source of these observations. It is unknown what sort of phallus occurs in other genera of this family. If Fig. 3.3 correctly depicts the evolutionary relationships in the Galloanserae, and if the Cracidae do have an anatid-like IO (see below), then the absence of an IO in some megapodes would be an additional instance of the independent evolutionary loss of the intromittent function of the avian phallus. Understanding the patterns of phallus evolution in the megapodes, particularly the evolutionary transformation from an intromittent to a non-intromittent phallus, presents an interesting challenge that might provide some useful insights into the reasons for IO loss in birds. Plate 3.1 Contd. ...

and spines in everted but flaccid state. C. Copulation in the Red-breasted merganser (Mergus serrator): typical of waterfowl that copulate on the water, the female’s cloaca is submerged, so the male phallus may help to prevent water from entering the cloaca and damaging sperm during copulation. Original photographs by Kevin McCracken (A, B) and Iztok Škornik (C).

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Reproductive Biology and Phylogeny of Birds

Cracidae (guans and currasows; 50 species). Gerhardt (1933), as well as Gadow and Selenka (1891), report that cracids have a phallus resembling that of the Mallard but details are sketchy and some modern work is really needed on this taxon. Gadow and Selenka (1891) report on three independent accounts of a cracid phallus from the early literature. In one of these accounts, the phallus is described as a spiral structure about 35 mm long with an ejaculatory groove and an opening (presumably a blind cavity) at the tip. Gerhardt (1933) studied, but did not dissect, a museum specimen of the Black currasow (Crax alector), and describes a flaccid phallus that resembled that of the Mallard and was 2.4 cm long with a blind cavity that could be everted to increase the total length to 3.9 cm. Given our current understanding of the relationship of the Cracidae to the other Galloanserae (Fig. 3.3), a detailed study of phallus anatomy within this taxon would be particularly useful. It would not be too surprising if the cracid phallus resembled that of the anatids since they share a common ancestor but further study might help us to understand how the intromittent phallus was lost in the clade comprising Phasianidae, Odontophoridae and Numididae because this clade also shares a common ancestor with the cracids (Fig. 3.3). Numididae (Guineafowl; 6 species). The guineafowl are expected to have a non-intromittent phallus like that of the chicken but its structure has never been described. Odontophoridae (New World quails; 32 species). The New World quails are also expected to have a non-intromittent phallus like that of the chicken but its structure has never been described. Phasianidae (pheasants and grouse; 180 species). All of the Phasianidae are expected to have a non-intromittent phallus like that of the chicken and turkey but there have not, to the best of our knowledge, been any studies of any other species in this family. The phalluses of the domestic chicken and turkey have, however, been very well studied. Both species have a non-intromittent phallus on the ventral lip of the vent that, even when not erect, can readily be seen by manually depressing this lip. This phallus appears to be homologous in structure to the intromittent true phalluses of the Neognathae and Palaeognathae. In the domestic chicken, the erect phallus is roughly heart-shaped with a median groove (Fig. 3.12A) into which the paired deferent ducts discharge semen directly. During erection of the phallus, both the vent and the floor of the proctodeum evert for a few seconds exposing the phallus externally. The chicken phallus has vascular bodies embedded in the cloacal wall in the same position as those of the Mallard, attesting to their common evolutionary origin, though their actual structures are substantially different (see King 1981a for details). The lymphatic channels of the vascular bodies connect with the lymphatic channels of both the phallic bodies and the lymphatic folds. Thus erection of the chicken phallus is accomplished by engorgement with lymph, as in the Mallard, and results in the eversion of the phallic area of the vent

Anatomy and Evolution of Copulatory Structures

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Fig. 3.12 Non-intromittent true phalluses of male A. Domestic chicken. B. Domestic turkey. Not drawn to scale, nor were scale bars shown on the original drawings. A modified after King, A. S. 1981a. Pp 107-147. In King, A. S. and McLelland, J. (eds), Form and Function in Birds, vol. 2. Academic Press, London, Fig. 3.8a; B modified after Briskie, J. V. and Montgomerie, R. 1997. Journal of Avian Biology 28: 73-86, Fig. 1e, who drew this from a photograph in King, A.S. 1981a. Pp. 107-147. In King, A.S. and McLelland, J. (eds), Form and Function in Birds, vol. 2. Academic Press, London, Fig. 3.10b.

and the adjacent cloacal floor. Both the phallic bodies and the lymphatic folds swell to meet along the midline of the phallus, thus creating a median groove (Fig. 3.12A) down which the semen flows during ejaculation. Immediately after ejaculation, lymph drains from the phallic bodies and lymphatic folds via the lymphatic vessels of the pudendal artery (Nishiyama 1955). The phallus of the domestic turkey is structurally and functionally similar to that of the chicken but differs in having two prominent prismoid humps separated by a deep furrow (Fig. 3.12B). The lymphatic folds extend obliquely across the floor of the proctodeum and have 3-4 prominent oblique ridges on their surface. Neoaves (about 9500 species) There is no strong evidence for a true phallus in any of the Neoaves, and direct evidence for a truly intromittent copulatory structure is only convincing for the Coracopsis parrots. During the breeding season, male Greater (C. vasa) and Lesser vasa parrots (C. nigra) of Madagascar and the Comoro Islands have a very large, heavily vascularized cloacal protrusion (Fig. 3.13A). This large fleshy bag is usually everted during copulation when it becomes dark red in color and extends 50 mm or more out of the cloaca. During one copulation the male mounted the female’s back and his cloacal protrusion appeared to completely enter the female’s cloaca, which stretched to accommodate it (Wilkinson and Birkhead 1995). After intromission, the male slipped off the female’s back but their cloacas remained interlocked for 104 min. When the male and female eventually disengaged, both sexes ejected a small volume of liquid and a small (5-15 mm) phallus-like organ was

!" Reproductive Biology and Phylogeny of Birds

Fig. 3.13 Pseudo-phalluses of male A. Vasa parrot. B. Buffalo weaver. C. Superb fairy wren. Arrows indicate the opening to the cloaca. Modified after Briskie, J. V. and Montgomerie, R. 1997. Journal of Avian Biology 28: 73-86., Fig. 1c, d, f.

observed at the tip of the cloacal protrusion. Some detailed anatomical work is needed to assess the structure of this apparent phallus. The remaining extant bird species in the Neoaves do not appear to have a phallus that is homologous to that of the Palaeognathae and the Galloanserae. Thus the true phallus appears to have been lost during the early evolution of this entire clade (Fig. 3.3). Although the literature from the 19th century does suggest some vestigial phallic structures might be present in some species of Neoaves (see King 1981a for details), it is far from certain whether these are homologous to the true phallus. For example, in the Emberizidae, Wolfson (1954a) reported that a pair of papillae formed by the proctodeal wall could be protruded from the vent when the cloacal protuberance was squeezed. These papillae meet and formed a median groove that carried semen during ejaculation. Wolfson (1954a) suggested the structure was analogous to the phallus of the rooster and functioned in the same manner. However, it is not clear whether these papillae are intromittent or whether they simply assist in passing the ejaculate to the female. An intromittent function has been proposed by a variety of authors but at present the exact positioning of the papillae during copulation is unknown. Papillae have now been observed in a variety of other passerine species (Briskie 1993; Birkhead et al. 1991; Birkhead and Hoi 1994; Nakamura and Matsuzaki 1995; Lombardo 2001; Chiba and Nakamura 2003). Although papilla size varied slightly across a small number of species measured by Briskie (1993), this variation did not appear to be related to differences in social mating system. The length of the papillae in the Alpine accentor (Prunella collaris) increases in the breeding season (Chiba and Nakamura 2003), suggesting that the increased size may be important for its functioning during copulation. However, no seasonal changes in the size of the papillae were found in the Tree swallow (Tachycineta bicolor; Lombardo 2001). As the size of the papillae in all species examined so far has been relatively small (about 1-3 mm long), it is unlikely that they penetrate very far into the female’s reproductive tract even if they are intromittent. More research is needed to

Anatomy and Evolution of Copulatory Structures

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confirm the presence and anatomical/histological details of papilla-like structures in other Neoaves (especially non-passerines), and to determine the exact role they play in sperm transfer.

3.3.3

Bird Cloacal Protuberances and other Structures

During the breeding season, the cloacal region of male passerine birds enlarges to such a degree that a protuberance forms as an outpocketing of the ventral abdominal wall (Fatio 1864; Salt 1954; Wolfson 1952, 1954a; Middleton 1972). In most species, this cloacal protuberance is formed primarily by the vastly enlarged seminal glomera (the distal ends of the ductus deferens). The seminal glomera function in sperm storage, and Wolfson (1954b) found that sperm are kept cooler in the cloacal protuberance as a result of its position outside the body cavity. However, it has also been suggested that the cloacal protuberance may function as a copulatory organ by elevating the cloaca from the body wall, thereby facilitating contact with the female genital area (Wolfson 1954a). The cloacal protuberance varies in size across species, and reaches its largest dimensions in those species with highly promiscuous mating systems in which males are subject to intense sperm competition (Nakamura 1990; Birkhead et al. 1991; Briskie 1993; Mulder and Cockburn 1993; Schulze-Hagen et al. 1995; Tuttle et al. 1996; Castro et al. 1996; Dixon and Birkhead 1997). Although such large cloacal protuberances are clearly the result of large seminal glomera containing large numbers of sperm, the enlarged size of the cloacal protuberance might also function to increase the efficiency of copulation in these species (Birkhead et al. 1993). Those species that copulated more frequently sported larger cloacal protuberances (Birkhead et al. 1993), but there was no relation between the duration of copulation and cloacal protuberance size. It is interesting to note that, in the Bearded tit (Panurus biarmicus), the cloacal protuberance is formed primarily from gelatinous connective tissue overlain by muscle layers, and not by the seminal glomera (which remain within the body cavity; Birkhead and Hoi 1994). As a result, the extension of the cloacal protuberance outside the body cavity cannot play a role in the temperature regulation of stored sperm in this species. Thus this cloacal protuberance may function primarily to improve ejaculate transfer by longer and betterpositioned cloacal contact (Sax and Hoi 1998). The enlarged cloacal protuberance of the New Zealand stitchbird (Notiomystis cincta) likewise contains only the ejaculatory ducts and not the seminal glomera (Castro et al. 1996). Low et al. (2005) found that the cloacal protuberance in this species not only increased in size over the breeding season but also changed orientation such that the opening to the vent rotated anteriorly by about 60 degrees. This “cloacal erection” was suggested to facilitate ejaculate transfer as it would allow males to more directly align the opening of their cloaca with that of the female during their face-to-face copulation (Low et al. 2005).

!$ Reproductive Biology and Phylogeny of Birds In a few species, the cloacal region has become modified further in ways that suggests some copulatory function. The skin beside the cloaca of both Red-billed (Bubalornis niger) and White-billed buffalo weavers (B. albirostris) of Africa have become highly modified to form an external phalloid organ (Fig. 3.13B) that is employed during copulation, although it is now known not to be intromittent (Winterbottom et al. 1999, 2001). This organ lies immediately anterior to the cloaca, is composed of connective tissue, has no sperm ducts, and is non-erectile. In the Red-billed buffalo weaver, the length of the male’s phalloid organ averages 15.7 mm (±0.29 mm SE, n = 109) whereas the female’s is much smaller (6.1±0.19 mm, n = 68). Interestingly, resident males had a significantly longer phalloid organ than non-residents, and residents with a harem of females had a significantly longer organ than those without a harem (Winterbottom et al. 2001). During copulation the male’s phalloid organ is rubbed vigorously against the female and appears to stimulate the male to orgasm and ejaculation (Winterbottom et al. 1999). In male Superb fairy wrens (Malurus cyaneus), the cloacal protuberance has a cartilaginous projection from its anterior surface (Fig. 3.13C; Mulder and Cockburn 1993). The location and phallus-like shape of this projection suggest that it may be a copulatory structure but its function is unknown. Other species of fairy wrens appear to have a similar projection (Tuttle and Pruett-Jones 2004) but details are lacking. Detailed studies of the dynamics of sperm transfer are needed in species with both large cloacal protuberances and pseudophalluses before their true nature can be determined. With only a handful of species studied in any detail, it is likely that a variety of other modifications to the external genitalia in the Neoaves may be discovered by careful observers.

3.4

ADAPTIVE SIGNIFICANCE OF IOs

To date, seven different hypotheses have been proposed to explain the pattern of presence/absence in the intromittent phalluses of birds. The true avian phallus was probably lost only once during the evolution of birds (Fig. 3.3) and is thus absent in all of the Neoaves, comprising >95% of extant avian species. Reasons for this loss remain obscure, in part because of the paucity of data on both the anatomy and physiology of phalluses in extant species, but also because of the technical (and ethical) difficulties in conducting critical experiments and the lack of power in a comparative analysis with only two states. The evolutionary history of avian IOs, on the other hand, is a little more complex in that these copulatory structures have originated at least once in birds (in the vasa parrots) and may have been lost three times if both the phylogeny and the data on the presence/absence of IOs shown in Fig. 3.3 is correct: once in the common ancestor of the Phasianidae-NumididaeOdontophoridae clade, once (at least) in the Megapodiidae, and once in the lineage leading to the Neoaves, the large sister clade of the Galloanserae (Fig. 3.3). This pattern of loss/gain is correct only if all of the Cracidae have

Anatomy and Evolution of Copulatory Structures

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an IO and the IO has been lost once in the Megapodiidae, but more work is needed to confirm this. In addition to this pattern of IO loss/gain, copulatory structures have arisen independently in the buffalo weavers, and possibly the fairy wrens but neither of these pseudophalluses appears to be intromittent. In this section, we summarize the various hypotheses proposed so far to explain the evolutionary loss of the IO in most birds, and evaluate the evidence for and against each of them. Because the IO appears to have been lost only three times in the evolutionary history of birds, we cannot employ modern comparative methods to evaluate these hypotheses. Instead, we draw on a variety of anatomical, physiological, and behavioral data at the species and family level in birds, and we use some recent data on variation in IO size within species and families to address these hypotheses. We have classified the hypotheses as being the result of natural and sexual selection and we present them individually but there is clearly no reason that more than one of them could not explain the loss of IOs. Thus, for example, different hypotheses may explain the maintenance of IOs in different taxa. In addition, the disappearance of the IO in avian evolutionary history logically requires that IOs are costly, and that cost must be due to either development or maintenance. All of these hypotheses have previously been evaluated in detail (Briskie and Montgomerie 1997, 2001). We therefore provide only a brief summary here plus new information that has come to light since those previous reviews.

3.4.1

Natural Selection Hypotheses

H1 Water Damage Lake (1981) suggested that an IO might prevent water from entering the female’s cloaca during copulation where it would either damage sperm (via osmotic shock), dilute the ejaculate, or wash it out of the cloaca. Thus IOs would be favored in species that copulate on the water (e.g., Plate 3.1C), but lost, due to some cost (e.g., see H4 and H5), in lineages that copulate out of the water. This hypothesis gains some support from the fact that the majority of the Anatidae, which have IOs, copulate on the water, whereas almost all species in the many other families of aquatic birds that copulate on land do not have IOs (Briskie and Montgomerie 1997). On the other hand, many birds that copulate on the water (e.g., some puffins, auklets, murrelets, and phalaropes) also lack an IO and do not seem to be at a particular disadvantage with respect to water damage, though at least a few species lift their cloacas clear of the water when copulating (Volume 6B, Chapter 6). On balance, the water damage hypothesis might provide an explanation for the maintenance of an IO in lineages that typically copulate on the water, but it cannot explain variation in IO size in those taxa nor can it explain why the IO has been lost in the majority of bird species. H2 Genital Contact King (1981a) suggested that an IO might provide an advantage for sperm transfer in species where cloacal contact is made difficult by the bird’s

!& Reproductive Biology and Phylogeny of Birds anatomy or its environment. Thus large-bodied, long-legged, and flightless birds (who may be less able to use their wings for balancing) might have difficulty maintaining cloacal apposition during sperm transfer. Briskie and Montgomerie (1997) suggested that this explanation was weak because even birds with an IO employ several methods to maintain genital contact and there was no clear pattern with respect to body size, long-leggedness, and the presence of an IO. Nevertheless, it is clear that the IO of vasa parrots is essential for maintaining their long period of genital contact as the male dismounts the female. It also seems to us that an IO would provide some selective advantage to males when cloacal contact is difficult to maintain, especially during forced copulations and interference from rival males. Coker et al. (2002), for example, found that ducks and geese with high levels of forced extrapair copulations were more likely to have longer IOs, and IOs with more and larger knobs, than monogamous species (Fig. 3.14A,B,C) where forced extrapair copulations are thought to be rare. In the subfamily Oxyurinae, in particular, males often display in groups to females and copulation can be quite tumultuous (McCracken 2000), so a relatively large, spiny IO might help a male to maintain genital contact with a female. H3 Copulation Duration Wesolowski (1999) proposed that the loss of an IO in most birds would result from selection for shorter copulations, either to reduce the period when copulating pairs would be vulnerable to predation or to minimize the period when the male would have to balance unsteadily on the female’s back (as in H2 above). Contrary to expectations from this hypothesis, males in species with an IO mounted females for shorter durations during copulation than those without an IO, controlling for body size (Briskie and Montgomerie 2001). Even within the non-passerines, species without an IO do not mount for shorter periods than those with an IO (Fig. 3.15A). Thus there is, so far, no empirical support for this interesting idea. Wesolowski (2001) correctly pointed out that the amount of time that a male spends mounting a female may not be an accurate index of the duration of copulation, but accurate data on the actual duration of cloacal contact in birds are rare (Volume 6B, Chapter 6) so this idea is difficult to test quantitatively without more data. H4 Flight Costs The loss of an IO might be expected in flying animals to reduce flight costs and increase maneuverability. Indeed, many birds with an IO are flightless or relatively poor fliers. With few exceptions, though, the mass of a bird’s IO is unlikely to be more than a tiny proportion of its body mass and thus its influence on flight would be negligible. Höhn (1960), for example, showed that the mass of the Mallard’s IO at the height of the breeding season was only 0.3% of the male’s body mass. Thus this hypothesis may only explain the extremely large IOs in ostriches, which are flightless.

Anatomy and Evolution of Copulatory Structures

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Fig. 3.14 Size and structure of the male intromittent organ (IO) in species from the family Anatidae in relation to the expected frequency of forced extrapair copulations FEPCs: A. IO length, controlling for variation in body size. B. Percent of the IO surface that is covered by knobs or ridges (see Fig. 3.10). C. Size of knobs or ridge. D. Showing residual IO size in relation to residual testes mass (both controlling for body size), where the latter is an index of the intensity of sperm competition. Redrawn from data in Coker, C. R. et al. Auk 119: 403-413, Figs. 3 and 4.

H5 Sexually Transmitted Diseases Briskie and Montgomerie (1997) suggested that sexually transmitted diseases (STDs) might pose a significant cost to the maintenance of an IO in birds. They argued that STDs might be especially problematic in birds because they are the only homeotherms that have a common cloaca for both defecation and reproductive functions. Thus, in birds, the male inserts his IO into a warm site through which fecal material passes, potentially an excellent environment for the proliferation of pathogens (e.g., Stewart and Rambo 2000). Wesolowski (1999) argued that STDs were unlikely to be prevalent in most birds, based on Lombardo’s (1998) suggestion that STDs should be rare in birds because, in seasonal breeders, the dispersal rates of highly virulent STDs should be low. Unfortunately, there are few hard data to address this idea but we do know

" Reproductive Biology and Phylogeny of Birds

Fig. 3.15 Tests of adaptive hypotheses to explain the pattern of presence (black bars and symbols) and absence (open bars and symbols) of intromittent organs (IOs) in birds: A. In non-passerines there is no difference in mean copulation duration (controlling for body mass) between species with (n = 16) and without (n = 167) an IO (ANCOVA on log-transformed variables, F1,180 = 0.03, P = 0.86; bar graphs show least squares means±95%CL). B. Male and female contributions to incubation in relation to the presence/absence of male IO (data from 10 families with and 86 without an IO). C. Relation between the volume of one egg and female body mass in species with (Anseriformes) and without (Galliformes) an IO in the Galloanserae. A from data in Briskie, J. V. and Montgomerie, R. 2001. Journal of Avian Biology 32: 184-187, Fig. 1; B and C from data in Briskie, J. V. and Montgomerie, R. 1997. Journal of Avian Biology 28: 73-86, Figs. 3 and 4.

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that virulent STDs have been isolated in the domestic chicken (Sheldon 1993), and that bacteria are sexually-transmitted in the Red-winged blackbird (Agelaius phoeniceus; Westneat and Rambo 2000). Moreover, both Spotted and Red-winged tinamous have plasma cells in the epithelium of the fixed base of the IO, suggesting an immunoprotective function (Oliveira et al. 2003). The dramatic increase in the number of these cells during the breeding season indicates enhanced immune function during this period, potentially to protect against STDs. While there are other potential explanations for the function of these cells, this finding does raise the intriguing possibility that birds with IOs are particularly susceptible to STDs. Given that IOs are clearly not needed for internal fertilization in birds, Briskie and Montgomerie (1997) argued that selective pressures due to the costs of development and maintenance, especially in the face of STDs, would favor the disappearance of the IO unless there was a particular benefit to retaining it. Two such benefits could accrue as a result of sexual selection via male-male competition or female choice, as follows.

3.4.2

Sexual Selection Hypotheses

H6 Sperm Competition (Male-Male Competition) Briskie and Montgomerie (1997) proposed that IOs may be favored particularly when there is competition among males to fertilize a female’s ova. They argued that IOs, and longer IOs, would allow a male to deposit his sperm higher up the female’s reproductive tract, closer to the site of sperm storage at the utero-vaginal junction. In addition, the spikes, knobs and ridges on some IOs might facilitate the displacement or removal of sperm of rival males from the female’s urogenital tract. Briskie and Montgomerie (1997) tested this hypothesis indirectly by comparing the incidence of IOs in species with male, female, and biparental incubation, arguing that species with male-only care would be more-strongly selected to ensure their paternity and thus maintain an IO. This prediction was supported by an analysis at the family level (Fig. 3.15B). Interestingly, the IO of domesticated Mallards (mean length 8 cm) is about twice as large as in the wild bird (mean length 4 cm). Rautenfeld et al. (1974) suggested that this difference was due to the increased sexual activity of the domesticated form but we see no reason that IO size should be influenced by sexual activity alone. Rather it seems to us that this difference in IO size may be due to increased sperm competition in domestic flocks, where groups of males have constant access to females in the absence of male territoriality, and possibly because of reduced costs of both development and maintenance of an IO in captivity. This apparently evolutionary change of IO size in domestic mallards deserves some in-depth research. Coker et al. (2002) also support this sperm competition hypothesis as an explanation for the interspecific variation in size of anatid IOs. They analyzed data on the IO size of 54 species of ducks and geese in relation to both mating systems and relative testes size (controlling for body size), an index of the

"

Reproductive Biology and Phylogeny of Birds

intensity of sperm competition (Briskie and Montgomerie, Chapter 9). Mating systems of 28 of these species in relation to the expected rate of forced extrapair copulations (FEPC) were classified as 1 = monogamous (presumably no FEPCs), 2 = rare FEPCs, 3 = frequent FEPCs, and 4 = polygynous or promiscuous (presumably FEPCs common), based on extensive natural history observations of the species studied. Both the presumed incidence of FEPCs (Fig. 3.14A) and the relative testes mass of males were significantly positively correlated with IO size (Fig. 3.14D). As noted above, males in species that were more likely to engage in male-male competition also had more and larger knobs and spikes on their IOs (Fig. 3.14B, C). Inceased number and sizes of knobs, spikes, and ridges on an IO would (i) enhance a male’s ability to stay inside the female’s cloaca while being harassed by rival males, (ii) allow a male to fully erect his IO inside the female, and (iii) potentially enhance a male’s ability to displace or extract the sperm of rival males who had previously recently copulated with the female, much in the manner of the odonate IO (Waage 1979). While males of all the species studied by Coker et al. (2002) have an IO, the patterns they uncovered show that both the size and structure of the avian IO are quite variable interspecifically and thus appear to respond to selection resulting from sperm competition. In the absence of sperm competition, it is therefore at least plausible that the phallus would be reduced in size over evolutionary time, due to costs of IO development and maintenance, until it is no longer intromittent. Given the potential advantage for even a small IO in the Anatidae to prevent water damage to sperm, we might not expect to see the disappearance of the IO in species that copulate on the water. Thus, it would be interesting to compare the IO sizes of species that copulate on and off the water, while controlling for the intensity of sperm competition. Similarly, IO size and structure in relation to mating patterns need to be examined in both the cracids and tinamous, two groups in which the large number of species would enable some rigorous tests of this hypothesis. H7 Female Choice Briskie and Montgomerie (1997) also suggested that the evolutionary loss of an IO in birds may have been driven by female choice. They reasoned that, if females could somehow prefer males that did not force copulations, the IO might serve no selective advantage. This hypothesis rests on four assumptions: (i) that IOs are not needed for internal fertilization in birds, (ii) that the main function of male IOs in birds is to force inseminations and/or to ensure that semen is deposited well into the female’s reproductive tract, (iii) that females have a mechanism to reject or reduce the fitness of males that they prefer not to mate with, and (iv) that IOs are too costly to develop and maintain if they do not provide any advantage to males that possess them. Assumption (i) is clearly true, and assumptions (ii) and (iv) are supported by some limited evidence as presented above. In accordance with assumption (iii), female birds potentially have the ability to abort or discard at relatively little cost any ova that are fertilized by non-preferred males. Since the ova of

Anatomy and Evolution of Copulatory Structures

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all birds are fertilized individually at intervals of usually 24-48 h (Howarth 1974), and there appears to be last male sperm precedence (Birkhead 1998), females have the potential to identify the paternity of each egg laid and lay it outside their nest, thus selecting against disfavored males. Briskie and Montgomerie (1997) reasoned that the costs of selective abortion or abandonment of ova/eggs would be related to their size relative to the female’s body size, and thus such a tactic would be most feasible for females that laid relatively small eggs. Consistent with this prediction, they found that females in species where the male has an IO lay relatively large eggs compared to species without male IOs (Fig. 3.15C). Indeed this pattern is particularly striking in the Galloanserae where male Anatidae have an IO, frequently engage in forced copulations, and females generally lay very large eggs, whereas in the Phasianidae, Odontophoridae and Numididae females lay relatively small eggs, males have a non-intromittent phallus, and forced copulations are less frequently observed. In other words, if eggs are large and costly to produce because they are large relative to body size, then females may be reluctant to abort or abandon such eggs, even if they are sired forcibly by males that the female does not prefer as a sire. In these species with relatively large eggs, males retain an advantage by forcing copulations and maintaining an IO, while in the smaller-egged species, the IO is proposed to have disappeared because females are able to control fertilizations through refusing to raise an unwanted male’s offspring. Like the sperm competition hypothesis, this female choice hypothesis predicts that IO size will be positively correlated with the frequency of forced extrapair copulations, but makes the unique prediction that IO size will also be positively related to the costs of egg production. Certainly among the Anatidae, the Oxyurinae have relatively large eggs compared to other species (Johnsgard 1978), and they also have the largest IOs, but a more comprehensive analysis of this prediction is clearly needed.

3.5

FUTURE DIRECTIONS

Despite research on avian copulatory organs that spans almost two centuries, we still have only the most rudimentary knowledge of IO structure and function in all but a few species. Even the taxonomic distribution of IOs and phalluses among taxa has not been comprehensively documented, including even some uncertainty about whether or not males in the Cracidae and Megapodidae possess an IO. There is clearly a need for more detailed anatomical work. Though such studies have not been popular for some time, modern anatomical techniques (e.g., Oliveira et al. 2000, 2004; Oliveira and Mahecha 2003), and the various hypotheses about the adaptive significance of IOs in birds presented above, make this a potentially attractive field of study. Some of the adaptive hypotheses summarized here could also be tested with experimental studies of captive birds. Many species in the Anseriformes and Galliformes can readily be bred in captivity so that hypotheses about

"" Reproductive Biology and Phylogeny of Birds water damage, sperm competition, female choice, and STDs might be tested with some clever experiments. There is certainly much to be learned about the incidence of STDs in birds and their effect on the anatomy, physiology, and costs of maintaining phalluses. With controlled experiments, it should also be possible to quantify the influence of IO size and structure on the outcome of sperm competition, especially within species in which there is natural variation in IO size (e.g., domestic versus wild Mallards). Selection experiments with domesticated Mallards would add some useful insights into the heritability and evolvability of IO size in birds. Because the taxonomic distribution of phalluses and IOs in birds is relatively simple, with both being largely ancestral traits (Fig. 3.3), there is virtually no scope for the sorts of comparative studies that have given us some insights into the evolution of other reproductive traits in birds (e.g., Briskie et al. 1997; see Chapter 9). Nonetheless, considerable progress can still be made in our understanding of both IO evolution in birds and the reasons for the loss of IOs in most of this Class by simply conducting more comprehensive anatomical research. Half a century ago , Fisher (1955) lamented the fact that ‘The “modern” trend in biological sciences seems all too often to imply that “anatomy as such” may be overlooked in the evolution of the “better and more accepted” avenues of approach to biological problems.’ Plus ça change... The time is ripe for some detailed anatomical work on avian copulatory structures, informed by modern phylogenies and some of the hypotheses about their adaptive significance that we have outlined above. We hope this chapter will inspire an enterprising graduate student to take up this challenge.

3.6

ACKNOWLEDGMENTS

We are grateful to Meghan Goodchild, Kristen Scott, and Christina Cliffe for help in compiling data and searching out references; and to Tim Birkhead and Barrie Jamieson for numerous excellent comments on the manuscript. Our work on avian reproductive tactics is supported by grants from the Natural Sciences and Engineering Research Council of Canada (to R. M.) and the University of Canterbury (to J. V. B.).

3.7 LITERATURE CITED Benz, G. D. 1983. Myology and the histology of the phalloid organ of the buffalo weaver (Bubalornis albirostris). Auk 100: 501-504. Birkhead, T. R. 1998. Sperm competition in birds. Reviews of Reproduction 3: 123129. Birkhead, T. R., Briskie, J. V. and Møller, A. P. 1993. Male sperm reserves and copulation frequency in birds. Behavioral Ecology and Sociobiology 32: 85-93. Birkhead, T. R., Hatchwell, B. J. and Davies, N. B. 1991. Sperm competition and the reproductive organs of the male and female Dunnock Prunella modularis. Ibis 133: 306-311. Birkhead, T. R. and Hoi, H. 1994. Reproductive organs and mating strategies of the bearded tit Panurus biarmicus. Ibis 136: 356-360.

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Birkhead, T. R., Stanback, M. T. and Simmons, R. E. 1993. The phalloid organ of buffalo weavers Bubalornis. Ibis 135: 326-331. Briskie, J. V. 1993. Anatomical adaptations to sperm competition in Smith’s Longspurs and other polygynandrous passerines. Auk 110: 875-888. Briskie, J. V. 1998. Avian genitalia. Auk 115: 826-828. Briskie, J. V. and Montgomerie, R. 1997. Sexual selection and the intromittent organ of birds. Journal of Avian Biology 28: 73-86. Briskie, J. V. and Montgomerie, R. 2001. Efficient copulation and the evolutionary loss of the avian intromittent organ. Journal of Avian Biology 32: 184-187. Briskie, J. V., Montgomerie, R. and Birkhead, T. R. 1997. The evolution of sperm size in birds. Evolution 51: 937-945. Brom, T. G. and Dekker, R. W. R. J. 1992. Current studies on megapode phylogeny. Zoologische Verhandelingen (Leiden) 278: 7-17. Caithness, T. A. 1971. Sexing kiwis. International Zoo Yearbook 11: 206-208. Castro, I., Minot, E. O., Fordham, R. A. and Birkhead, T. R. 1996. Polygynandry, faceto-face copulation and sperm competition in the Hihi Notiomystis cincta (Aves: Meliphagidae). Ibis 138: 765-771. Chiba, A. and Nakamura, M. 2003. Anatomical and histophysiological characterization of the male cloacal protuberance of the polygynandrous Alpine Accentor Prunella collaris. Ibis 145: E83-E93. Coker, C. R., McKinney, F., Hays, H., Briggs, S. V. and Cheng, K. M. 2002. Intromittent organ morphology and testis size in relation to mating system in waterfowl. Auk 119: 403-413. Cracraft, J., Barker, F. K., Braun, M., Harshman, J., Dyke, G. J., Feinstein, J., Stanley, S., Cibois, A., Schikler, P., Beresford, P., García-Moreno, J., Sorenson, M. D., Yuri, T. and Mindell, D. P. 2004. Phylogenetic relationships among modern birds (Neornithes): toward an avian tree of life. Pp 468-489. In Cracraft, J. and Donoghue, M. J. (eds), Assembling the Tree of Life. Oxford University Press: New York. 592 pp. Dickinson, E. C. (ed.) 2003. The Howard and Moore Complete Checklist of the Birds of the World, 3rd Edition. Princeton University Press, Princeton, New Jersey. 1056 pp. Dixon, A. and Birkhead, T. R. 1997. Reproductive anatomy of the reed bunting: a species which exhibits a high degree of sperm competition through extra-pair copulations. Condor 99: 966-969. Eberhard, W. G. 1990. Animal genitalia and female choice. American Scientist 78: 134141. Eckhard, C. 1876. Ueber die Erection der Vögel. Beitrage zur Anatomie und Physiologie 7: 116-125. Fatio, M. V. 1894. Note sur une particularite de l’appareil reproducteur male chez l’Accentor alpinus. Revue et Magazine de Zoologie pure et appliquée 27: 65-67. Fisher, H.I. 1955. Avian anatomy, 1925-1950, and some suggested problems. Pp 57105. In Wolfson, A. (ed.), Recent Studies in Avian Biology. University of Illinois Press: Urbana. 479 pp. Gadow, H. and Selenka, E. 1891. Vögel. Anatomischer Thiel. Vol. 6 Winter, Leipzig. Gerhardt, U. 1933. Kloake and Begattungsorgane. In Bolk, I., Goppert, E., Kallius, F. and Lubosh, W. (eds), Handbuch der vergleichenden Anatomie der Wirbeltiere, Vol. 6. Urban and Schwarzenberg: Berlin. Guzsal, E. 1974. Erection apparatus of the copulatory organ of ganders and drakes. Acta Veterinaria Academiae Scientiarum Hungaricae 24: 361-373.

"$ Reproductive Biology and Phylogeny of Birds Howarth, B. 1974. Sperm storage as a function of the female reproductive tract. Pp. 237-279. In Johnson, A.D. and Foley, C.E. (eds), The Oviduct and its Functions. Academic Press, New York. 369 pp. Höhn, E. O. 1960. Seasonal changes in the mallard’s penis and their hormonal control. Proceedings of the Zoological Society of London 134: 547-555. Johnsgard, P. A. 1978. Ducks, Geese and Swans of the World. University of Nebraska Press, Lincoln. 404 pp. Jones, D. N., Dekker, R. W. R. J. and Roselaar, C. S. 1995. Bird Families of the World: The Megapodes. Oxford University Press, Oxford. 282 pp. King, A. S. 1981a. Phallus. Pp 107-147. In King, A.S. and McLelland, J. (eds), Form and Function in Birds, vol. 2. Academic Press, London. 496 pp. King, A. S. 1981b. Cloaca. Pp 63-105. In King, A.S. and McLelland, J. (eds), Form and Function in Birds, vol. 2. Academic Press, New York. 496 pp. Komárek, V. and Marvan, F. 1969. Beitrag zur mikroskopischen Anatomie des Kopulationsorganes der Entesvögel. Anatomischer Anzeiger 124: 467-476. Lake, P.E. 1981. Male genital organs. Pp 1-61. In King, A.S. and McLellend, J. (eds), Form and Function in Birds. vol. 2. Academic Pres, London. 496 pp. Larson, P. L. and Frey, E. 1992. Sexual dimorphism in the abundant Upper Cretaceous theropod Tyrannosaurus rex. Journal of Vertebrate Paleontology 12: 38A. Liebe, W. 1914. Die mannliches betattungsorgane der hausente. Jenaische Zeitschrift für Naturwissenschaft 51: 627-696. Lombardo, M. P. 1998. On the evolution of sexually transmitted diseases in birds. Journal of Avian Biology 29: 314-321. Lombardo, M. P. 2001. Individual and seasonal variation in external genitalia of male tree swallows. Auk 118: 789-795. Low, M., Castro, I. and Berggren, A. 2005. Cloacal erection promotes vent apposition during forced copulation in the New Zealand stitchbird (hihi): implications for copulatory efficiency in other species. Behavioral Ecology and Sociobiology 58: 247-255. Marchant, S. M. and Higgins, P. J. (eds) 1990. Handbook of Australian, New Zealand and Antarctic Birds. Volume 1, Ratites to Ducks, Part B Australian Pelican to Ducks. Oxford University Press, Melbourne. 1406 pp. McCracken, K. 2000. The 20-cm spiny penis of the Argentine lake duck (Oxyura vittata). Auk 117: 820-825. McCracken, K., Wilson, R. E., McCracken, P. J. and Johnson, K. P. 2001. Are ducks impressed by drakes’ display? Nature 413: 128. Middleton, A. L. A. 1972. The structure and possible function of the avian seminal sac. Condor 74: 185-190. Mulder, R. A. and Cockburn, A. 1993. Sperm competition and the reproductive anatomy of male superb fairy-wrens. Auk 110: 588-593. Müller, J. 1836. Uber zwei verschiedene typen in dem bau der erectilen mannlichen geschlechtsorgane bei den straussartigen vogeln. Gelesen in den kgl. Akad. Wiss. Physikal. Abhandl. 137-177. Müller, R. 1908. Über den Tannenberg’schen Korper. Archiv für die gesamte Physiologie 122: 455-483. Nakamura, M. 1990. Cloacal protuberance and copulatory behavior of the Alpine Accentor (Prunella collaris). Auk 107: 284-295. Nakamura, M. and Matsuzaki, Y. 1995. Sex determination based on cloacal protuberances in the Japanese accentor Prunella ribida. Journal of the Yamashina Institute of Ornithology 27: 78-88.

Anatomy and Evolution of Copulatory Structures

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Nishiyama, H. 1955. Studies on the accessory reproductive organs in the cock. Journal of the Faculty of Agriculture of Kyushu University 10: 277-305. Oliveira, C. A., Geraldo, I., Poblete, P. C. P., Macedo, G. F. and Machecha, G. A. B. 2003. Intraepithelial plasma cells in the avian copulatory organ of two tinamou species: quantitative variation during the breeding season. Anatomy and Embryology 207: 409-416. Oliveira, C. A. and Mahecha, G. A. B. 2000. Morphology of the copulatory apparatus of the spotted tinamou Nothura maculosa (Aves: Tinamiformes). Annals of Anatomy 182: 161-169. Oliveira, C. A., Silva, R. M., Santos, M. M. and Mahecha, G. A. B. 2004. Location of the ureteral openings in the cloacas of tinamous, some ratite birds, and crocodilians: a primitive character. Journal of Morphology 260: 234-246. Padian, K. and Horner, J. R. 2002. Typology versus transformation in the origin of birds. Trends in Ecology and Evolution 17: 120-124. Rautenfeld, D. B., Preuss, F. and Fricke, W. 1974. Neue daten zur Erektion und Reposition des Erpelphallus. Praktische Tierarzt 10: 553-556. Salt, W. R. 1954. The structure of the cloacal protuberance of the Vesper sparrow (Pooecetes gramineus) and certain other passerine birds. Auk 71: 64-73. Sax, A. and Hoi, A. 1998. Individual and temporal variation in cloacal protuberance size of male Bearded Tits (Panurus biarmicus). Auk 115: 964-969. Schulze-Hagen, K., Leisler, B. and Birkhead, T. R. 1995. Prolonged copulation, sperm reserves and sperm competition in the Aquatic Warbler Acrocephalus paludicola. Ibis 137: 85-91. Sheldon, B. C. 1993. Sexually transmitted disease in birds: occurrence and evolutionary significance. Philosophical Transactions of the Royal Society B 339: 491-497. Sibley, C. G. and Ahlquist, J. E. 1990. Phylogeny and Classification of Birds: A Study in Molecular Evolution. Yale University Press, New Haven. Stewart, R. and Rambo, T. B. 2000. Cloacal microbes in house sparrows. Condor 102: 679-684. Tuttle, E. M. and Pruett-Jones, S. 2004. Estimates of extreme sperm production: morphological and experimental evidence from reproductively promiscuous fairy-wrens (Malurus). Animal Behaviour 68: 541-550. Tuttle, E. M., Pruett-Jones, S. and Webster, M. 1996. Cloacal protuberances and extreme sperm production in Australian fairy-wrens. Proceedings of the Royal Society B 263: 1359-1364. Waage, J. K. 1979. Dual function of the damselfly penis: sperm removal and transfer. Science 203: 916-918. Wesolowski, T. 1999. Reduction of phallus in birds—an avian way to safe sex? Journal of Avian Biology 30: 483-485. Wesolowski, T. 2001. Reduction of phallus in birds—a reply to Briskie and Montgomerie. Journal of Avian Biology 32: 188. Westneat, D. F. and Rambo, T. B. 2000. Copulation exposes female Red-winged Blackbirds to bacteria in male semen. Journal of Avian Biology 31: 1-7. Wilkinson, R. and Birkhead, T. R. 1995. Copulation behaviour in the vasa parrots Coracopsis vasa and C. nigra. Ibis 137: 117-119. Winterbottom, M., Burke, T. and Birkhead, T. R. 1999. A stimulatory phalloid organ in a weaver bird. Nature 399: 28.

"& Reproductive Biology and Phylogeny of Birds Winterbottom, M., Burke, T. and Birkhead, T. R. 2001. The phalloid organ, orgasm and sperm competition in a polygynandrous bird: the red billed buffalo weaver (Bubalornis niger). Behavioral Ecology and Sociobiology 50: 474-482. Wolfson, A. 1952. The cloacal protuberance—a means for determining breeding condition in live male passerines. Bird Banding 23: 159-165. Wolfson, A. 1954a. Notes on the cloacal protuberance, seminal vesicles, and a possible copulatory organ in male passerine birds. Bulletin of the Chicago Academy of Sciences 10: 1-23. Wolfson, A. 1954b. Sperm storage at lower than body temperature outside the body cavity in some passerine birds. Science 120: 68-71. Wood Jones, F. 1915. The chelonian type of genitalia. Journal of Anatomy and Physiology 49: 383-406.

CHAPTER

4

Developmental Anatomy of the Female Reproductive Tract Monika Jacob1 and Murray R. Bakst2

4.1

INTRODUCTION

In vertebrates, the reproductive system arises as bilateral anlagen, and consequently, paired genital organs are commonly found in the adult. In birds, the female reproductive system is unique. Although paired anlagen appear, only the left genital primordia further develop to functional organs except in birds of prey (order Ciconiiformes (=Falconiformes); families: Cathartidae (American vultures), Accipitridae (kites, eagles, hawks, and allies), Falconidae (caracaras and falcons). In cases of persisting right genital organs, double oviducts are less frequently observed than double ovaries. However, according to Kinsky (1971) they have also been found in other orders than Falconiformes. The reason for the unilateral development of female genital organs might be to reduce weight for flying (see discussion by Gilbert, 1979). The question than arises why the falconiforms allow themselves the luxury of two genital tracts. To reduce weight their hard-shelled eggs are perhaps relatively small and are laid down at an early stage of development. The weight of the egg in relation to the maternal body weight in falconiforms varies from 2.75% in the Whitetailed eagle, Haliaeetus albicilla (Accipitridae) to 8.5 % in Falco (Falconidae) and thus is at a lower level compared with non-falconiforms (Starck 1965). A relatively long incubation time and a small number of eggs laid are also found in falconiforms in comparison with other birds. Yet, these data do not yield an entirely satisfactory explanation for either a single or a double female genital tract. When ovulation occurs, the mature follicular oocyte is released from the ovary and is received by the oviduct. This tract surrounds the ovum with the albumen, the shell membranes, and the shell to form the characteristic avian 1 2

Abteilung für Anatomie und Embryologie, Ruhr-Universität Bochum, Bochum, Germany Biotechnology and Germplasm Laboratory, ARS/USDA, Beltsville, Maryland 20705 USA

# Reproductive Biology and Phylogeny of Birds egg. In addition to forming and transporting the egg, the oviduct is the site of sperm storage, sperm transport, fertilization and early embryonic development. (See Chapter 11 for more details about the role of the oviduct in reproduction.). In this chapter we focus on the development and differentiation of the left oviduct from the Müllerian (paramesonephric) duct and the concurrent regression of the right Müllerian duct. We further present the macroanatomy, histology and ultrastructure of the oviduct in hens. Data are mainly based on studies of the domestic fowl, Gallus domesticus, since these birds have been systematically studied. Only fundamental peculiarities of wild birds (as far as known) are mentioned in this chapter. A link is made to the Wolffian duct and the mesonephros, which form the reproductive tract in the male, but does not further develop in females. Only remnants of these anlagen may be found in hens.

4.2

EARLY MÜLLERIAN DUCT DEVELOPMENT

The Müllerian ducts, the anlagen of the female genital tracts in vertebrates, arise in both sexes by inductive activity of the Wolffian (mesonephric) ducts. The first sign of Müllerian duct development is a thickening of the coelomic epithelium adjacent to each Wolffian duct in stage 19 HH chick embryos [HH designates Hamburger and Hamiliton (1954) normal table of staging chicken embryos]. This thickened epithelium forms the bilateral Müllerian ridge (Fig. 4.1A-C) extending from cranial to caudal on the lateral side of the mesonephros. Interestingly, nephrostome-like depressions are found within the cranial part of both ridges (Fig. 4.1C, D). They are localized caudal and lateral to a group of external (coelomic) glomeruli (Fig. 4.1A, C), which are well developed even in the pronephros of higher vertebrates, including humans, and are comparable to the glomeruli of the other kidney generations (Jacob et al. 1977, 1979, 1986). Peritoneal funnels of cranial mesonephric tubules are also found adjacent to the external glomeruli (Jacob 1986; Hiruma and Nakamura 2003). The nephrostome-like depressions at the lateral side of each cranial Fig. 4.1 Gallus gallus, Fowl. A. Transverse section of the rostral part of the Müllerian ridge (arrow) from a stage 23 HH (4 days) embryo. Scale bar = 50 µm. WD, Wolffian duct; EG, external glomerulus B. Schematic drawings of serial sections from the rostral part of the Müllerian ridge (arrows) adjacent to the Wolffian duct (WD). Note the foveolae within the epithelium of the Müllerian ridge. Scale bar = 100 µm. C. Scanning electron micrographs of the mesonephros (Mn) from a stage 24 HH (4, 5 days) embryo. Irregularly arranged external glomeruli (arrowheads) are found on the medial side of the mesonephros near the mesenterium (Mt). Note the nephrostomal-like foveolae within the rostral part of the Müllerian ridge. Scale bar = 100 µm. Go, Gonads; *, Müllerian ridge D. Detail from the Müllerian ridge with nephrostomal-like openings. Scale bar = 10 µm. Original.

Developmental Anatomy of the Female Reproductive Tract

Fig. 4.1

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#

Reproductive Biology and Phylogeny of Birds

Fig. 4.2 Gallus gallus, Fowl. A. Scanning electron micrograph from a section through the rostral part of the mesonephros (stage 26 HH, 5 days). The arrow Fig. 4.2 Contd. ...

Developmental Anatomy of the Female Reproductive Tract

#!

mesonephros represent only shallow epithelial invaginations from which cells detach to form the anlage of the paired Müllerian duct (Fig. 4.1B). According to Abdel-Malek (1950) they may considered as pronephrostomes that have already shifted laterad while the pronephric tubules have already degenerated. Similarly, in primitive vertebrates, like Ichthyophis (Gymnophiona: Amphibia), pro-and mesonephric nephrostomial tubules are found with an outer ciliated funnel at the lateral side (Semon 1890, 1892). The fate of nephrostomal tubules was intensely studied by Wrobel and Süß (2000) in larvae of the Gymnophiona (Ichthyophis kohtaoensis) as well as in bovine embryos. Their comparative studies furnished evidence that the nephrostomial tubules are involved in the formation of the Müllerian duct infundibulum. It can be speculated that the lateral depression of the coelomic epithelium in chick embryos might be a relicts of such an outer funnel, but the complete evidence for this is lacking. Abdel-Malek (1950) has suggested that a highly pronounced nephrostome at the posterior part of somite 17 eventually forms the abdominal ostium of the oviduct in the chicken. However, the caudal nephrostome-like depressions fuse and form the abdominal ostium of the oviduct (Jacob et al. 1999, 2000). Such a “funnel field” is shown in Fig. 4.2A in a six-day chick embryo (stage 28 HH). One day later (stage 30 HH), a deep oval ostium has formed (Fig. 4.2B), which is subdivided by a crest of rounded cells presumably growing inwards to form the ostium. Therefore, the old opinion which up to now is found in textbooks of embryology (e.g. Hamilton 1965), that the cranial part of the Müllerian duct develops by fusion of the lips of a longitudinal groove arising within the cephalic end of the tubal ridge does not hold true for higher vertebrates, including birds. Our results argue for an outgrowth of the Müllerian duct anlage from the funnel region since at early developing stages, the abdominal aperture as well as the Müllerian duct proper represent highly proliferating tissues as revealed by the BrdU-anti BrdU reaction (Fig. 4.2C, D). Fig. 4.2 Contd. ...

indicates the funnel region with several nephrostomal-like foveolae. On the cut plane, the Wolffian duct (WD) and the anlage of the Müllerian duct (MD) are visible. B. The same region in a stage 30 HH (7 days) embryo. One oval ostium with a crest of cells has formed. C. and D. BrdU-anti-BrdU reaction through the funnel region and a more caudal part of the MD, note the high proliferation within the MD anlage (asterisks) as compared with the WD. E. Electron micrograph from the not yet fully canalized part of the Müllerian duct (MA) in a stage 26 HH embryo. The arrow indicates a nephrostome-like foveola. F. The tip of the Müllerian duct (MD) with a group of densely packed mesenchymal cells from a stage 25 HH (4.5-5 days) embryo. G. At stage 28 HH (5,5-6 days) the caudal part of the MD is found in close vicinity to the WD, but separated from the coelomic epithelium (CE) by a wide acellular space. The wall of the WD, which is in contact with the Müllerian duct, is double-layered and much thicker on the opposite side, both duct share a common basal lamina. Scale bars in A, B. E and G = 10 µm, in C and D = 50 µm, and F = 2,5 µm. Original.

#" Reproductive Biology and Phylogeny of Birds

Fig. 4.3 Gallus gallus, Fowl. A-C stage 28 HH A. Scanning electron micrograph from a section through the rostral part of the Müllerian duct (MD). A secondary duct Fig. 4.3 Contd. ...

Developmental Anatomy of the Female Reproductive Tract

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Thus, the Müllerian duct develops from a placodal-like funnel region, which invaginates to form a pit. Here cells detach from the coelomic epithelium and grow caudad (Fig. 4.2E). At first a solid cord of cells is established (Fig. 4.2F). The caudal migrating end still preserves a mesenchymal, rod-like configuration without a basal lamina. The cranial part transforms to a canalized, epithelial duct (Fig. 4.2G), which gradually opens into the deepened ostium. A graphic reconstruction based on serial sections from a stage 28-chick embryo has illustrated that caudal to the abdominal ostium additional apertures appear at intervals of about 100 µm (Jacob et al. 1999). They are connected with the main Müllerian duct by accessory ducts (Fig. 4.3A). Balfour and Sedgwick (1878) already described multiple openings of the cranial end of the Müllerian duct, and considered them as parts of a rudimentary head kidney (pronephros) in the chicken. According to Hamilton (1965), two or more coelomic apertures of the oviduct usually occur near the ostium on the fifth day since the groove-like anterior part of the Müllerian duct does not close uniformly. However, such a mechanism could not be observed in our studies. The Müllerian duct elongates very rapidly. At stage 28 HH it has a length of about 2 mm. The caudal part is found in close vicinity to the Wolffian duct and both ducts are enclosed by a common basal lamina (Fig. 4.3B, 4.2G). At the same time, a broad space filled with a dense fibrillar matrix has been established between the Müllerian duct and coelomic epithelium (Fig. 4.3C). Because of their close relation, it has been speculated that cells from the Wolffian duct contribute to the Müllerian anlage (see for discussion Didier, 1973). However, our own experimental and morphological data have clearly shown that Wolffian duct cells do not invade the Müllerian duct epithelium. Isotopic grafts of the tip of the Wolffian duct between chick and quail embryos provided almost normally developed embryos. In such cases a normal chickderived Müllerian duct develops adjacent to the quail Wolffian duct, but no quail (Wolffian duct) cells, possessing their characteristic distribution of heterochromatin, are found within the Müllerian duct (Fig. 4.3D). Fig. 4.3 Contd. ...

(2) is found beside the primary duct (1). B. The immunostaining of laminin exhibits the common basal lamina between Wolffian duct (WD) and MD. C. The electron micrograph reveals the dense fibrillar matrix between MD and coelomic epithelium. D. Transverse section through a chimera with grafted quail WD at the place of the chick duct. Feulgen staining demonstrates the quail nuclei within the WD epithelium. The MD only contains only chick cells. Courtesy of Dr. H. J. Jacob. E and F. Scanning electron micrograph from a transverse section (for technique see Jacob et al. 1999) through the rostral part of the MD separated from the WD by circular layer of mesenchyme. The detail shows a dense fibrillar matrix between the mesenchymal cells. Scale bars in A, B, E = 10 µm, in C = 1 µm and D = 50 µm, and F = 5 µm. Original.

#$ Reproductive Biology and Phylogeny of Birds

Fig. 4.4 Gallus gallus, Fowl. A. Sagittal section through the mesonephos and Müllerian duct (MD) of a stage 32 HH (7,5 days) embryo. In contrast to the Wolffian Fig. 4.4 Contd. ...

Developmental Anatomy of the Female Reproductive Tract

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Nevertheless, the developmental relationship between Wolffian and Müllerian duct has teratological importance in all vertebrates. Gruenwald (1941), first, delivered experimental evidence that destruction of the Wolffian duct results in kidney and Müllerian duct aplasia. Likewise in Emx-2 and GATA-3 deficient mice with absent Wolffian ducts, Müllerian ducts fail to develop (see Saino 2003). During further development, a mesenchymal sheath surrounds the epithelial duct (Fig. 4.3E). According to Didier (1973), and our own observations, the mesenchyme of the Müllerian duct is formed by epitheliomesenchymal transformation of the pseudostratified epithelium of the Müllerian ridge. The basal lamina ruptures, cells form pseudopodia and detach from the epithelium to surround the epithelium of Müllerian duct. Between the concentric layers of mesenchymal cells a dense extracellular matrix is established (Fig. 4.3F). Up to the eighth day (stage 33/34 HH), the development of the right and left, female and male duct is identical. The Müllerian duct still runs parallel to the Wolffian duct (Fig. 4.4A). But a dense mesenchymal tunic separates it from the dilated Wolffian duct. The latter has a wide lumen, and is surrounded by loose mesenchyme. The epithelium of the Müllerian duct has thickened and is now pseudostratified (Fig. 4.4B). It is highly proliferative as indicated by many mitotic figures. On the eighth day, the involution of the rostral part of the mesonephros has started. Thus, the anlage of the ostium abdominale is now found behind the lung anlage (Fig. 4.4C) and is attached to the mesonephros by a double-layered fold (ligament) of serosa epithelium. The middle part of the Müllerian duct projects over the lateral surface of the mesonephros, surrounded by the serosa epithelium of the former Müllerian ridge, which has now a flat appearance like the other parts of the splanchnopleure (Fig. 4.4D). The data here presented have mainly been obtained from the order Galliformes (species Gallus gallus and Coturnix coturnix japonica) but similar developmental processes were also found in the order Anseriformes in the Bean goose, Anser anser, and the domestic duck. A homology of avian Müllerian duct development with that of amphibians was postulated by Didier (1973). We believe that the early development of the paramesonephric ducts is similar in all vertebrates including humans. Fig. 4.4 Contd. ...

duct (WD), the MD is surrounded by dense mesenchyme. Scale bar = 100 µm. B. Detail of the MD shows a pseudostratified epithelium with many mitotic figures. Scale bar = 10 µm. C Oblique transverse section of a stage 34 HH (8 days) embryo. Arrows, Müllerian duct; G, gonad; L, liver; Lu, lung; Ms, mesonephros; St, stomach. The funnel region of the MD lies dorsal to the lung anlage. Scale bar = 0.5 mm. D. Detail of the left MD. Scale bar = 10 µm. Original.

#& Reproductive Biology and Phylogeny of Birds

4.3 REGRESSION OF THE RIGHT MÜLLERIAN DUCT AND DIFFERENTIATION OF THE LEFT ONE The early development of the two Müllerian (paramesonephric) ducts is identical in both sexes. At day seven they reach the cloaca with the compact distal end, but there is no opening into the cloaca prior to hatching. An occluding plate between oviduct and cloaca even persists up to the first breeding season (from Gilbert, 1979). In the male embryo, both Müllerian ducts, and in the female embryo, the right Müllerian duct undergos regression starting at the eighth day of development. According to Lutz-Ostertag (1954) the regression of the male duct occurs in two phases: first, the epithelium undergoes cell death; and second, the ducts are transformed into dense mesenchyme. The process of regression is obviously highly conserved in vertebrates as has been described in morphological studies in reptilians (Austin 1989, 1995), birds (Forsberg and Olivecrona 1963) and mammals (Dyche 1979; Trelstad et al. 1982; Wartenberg 1985). In male embryos apoptosis appears to be the main cellular mechanism in Müllerian duct regression. The central molecule in the apoptotic machinery is the anti-Müllerian hormone (AMH) also named Müllerian-inhibiting substance (MIS) that is a member of the transforming growth factor-beta (TGF-beta) superfamily (see Behringer 1995, Visser 2003, Rey et al. 2003) (see also Chapter 12). Like other TGF-beta family members, AMH appears to signal through two transmembrane receptors, both serine/threonine kinases. The AMH type II receptor, which has been clearly defined, binds to the ligand, and, it is proposed, triggers the formation of a complex with an assumed type 1 receptor ALK2 (activin-like kinase). The signaling pathway to activate the proapoptotic gene probably uses cytoplasmatic Smads (mothers against decapentaplegic related gene product) proteins (Clarke et al. 2001). The AMH type II receptor is exclusively expressed in the mesenchymal sheath surrounding the epithelial lining of the Müllerian duct, whereas apoptotic cells are localized within the epithelium (Baarends et al. 1994; Roberts et al. 1999). Thus, AMH functions indirectly via a paracrine mechanism. One of the first morphological features in Müllerian duct regression is the loss of the basal lamina (Trelstad et al. 1982) and the condensation of the mesenchymal sheath (Wartenberg 1985). These processes start in the male chick embryo around the eighth day of development. The formation of an “epithelial cuff” by the condensed mesenchyme is shown in Fig. 4.5C in a 9day-old embryo. As compared with the female ducts (Fig. 4.5A, B) the lumen has narrowed, and the diameter of the duct is decreased. Apoptotic bodies appear within the epithelium. Figure 4.5D from an 11.5-day-old male embryo reveals that the epithelium has disappeared. Only dense mesenchyme mark the former location of the Müllerian duct. The molecular and cellular events during Müllerian duct regression were demonstrated by Allard et al. (2000) including apoptosis and epithelial-

Developmental Anatomy of the Female Reproductive Tract

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Fig. 4.5 Gallus gallus, Fowl. A. Transverse section through the right Müllerian duct (MD) of a stage 36 HH (10 days) female embryo at the level of the gonad without sign of regeression. B. Section of the left MD at stage 33/34 HH (8 days) at the same level revealing many mitotic figure in the epithelium. C Transverse section of the right MD of a stage 35 HH (9 days) male embryo. The diameter is smaller than that of a comparable duct in the female. Note also the narrow lumen and the apoptotic bodies. D. At stage 37/38 HH (11, 5 days) the MD (here the left side) is replaced by dense mesenchyme. Scale bars = 25 µm. Original.

mesenchymal transformation. b-catenin, in association with lymphoid enhancer factor 1, is suggested to be involved in this process. Furthermore, Roberts et al. (2002) propose potential roles of the matrix metalloprotease MMP2 in mediating duct regression. Not all cells of the regressing Müllerian duct undergo apoptosis. Viable cells transform into mesenchyme and are able to migrate into other locations such as the mesonephric tubules (Hutson et al. 1984; Austin 1995). The mechanisms leading to the regression of the right Müllerian duct in female chick embryos are probably quite similar to those in male duct regression with AMH as a trigger. While in mammals, AMH is synthesized only in Sertoli cells immediately after testicular differentiation and in ovarian granulosa cells after birth, the situation is quite different in birds. Prior to 14 days of development, AMH is produced in the ovary. The left ovary synthesizes AMH at a level similar to that seen in the testis (Hutson et al. 1981). Its maximum level is at 14 days of development, latter than the testes,

$ Reproductive Biology and Phylogeny of Birds

Fig. 4.6 Gallus gallus, Fowl. Schematic drawing to show the differentiation of the left and the regression of the right MD. A. Stage 34 HH (8 days). Both MD are equally formed. B. Stage 37 HH (11 days). The regression of the right MD has Fig. 4.6 Contd. ...

Developmental Anatomy of the Female Reproductive Tract

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but at a time when the right ovary (ovotestis) is drastically reduced in size (Teng 1987). It is this peak in AMH synthesis by the left ovary that may account for regression of the right ovary. Similarly, chick AMH mRNA expression in the ovary was found to peak at the seventeenth day but in a lower amount than in the testis (Carré-Eusèbe et al. 1996). In mammals, the onset of AMH expression depends on SOX9 -a member of the SRY gene family. However, in the chick embryo AMH is expressed in males one day before SOX9 transcripts appear and SOX9 is never expressed in females (Oreal et al. 1998). Furthermore, the chicken differs from mammals in that AMH is expressed in dispersed medullary cells of the indifferent gonad (Oreal et al. 2002). With the early differentiation of the left ovary AMH concentrates in the outer medullary zone whereas a dispersed expression is preserved in the right ovary. Since AMH expression in indifferent and female gonads is not correlated with factors as in testis or follicles, Oreal et al. (2002) and Lasala et al. (2004) postulated a bird-specific control mechanism of AMH transcription. The apoptotic pathway in the female right Müllerian duct is described as caspase-3 mediated (Teng 2000) and shows the typical DNA laddering due to internucleosomal fragmentation. Recently, Ha et al. (2004) have shown that MMP2 (matrixmetalloprotease) mRNA of the right female Müllerian duct was significantly higher than on the left side at days 15 to 18 of incubation coinciding with the time of regression, and that diethylstilbestrol could decrease MMP2 expression. The female left Müllerian duct and even its right duct (Teng 1987) are much more resistant to AMH than the male ducts, and it is therefore obvious that a special protection exist. Many studies have shown that estrogenic hormones prevent the induction of left duct regression (Wolff, Et 1939; Hutson et al. 1982, 1985). While estrogen is synthesized in both male and female gonads in chick embryos (Woods and Erton 1978), it is much higher in the ovary than in the testis especially on the twelfth to the fifteenth day of incubation (Guichard et al. 1977). Furthermore, nuclear estrogen receptors were found to be higher in the female left Müllerian duct (MacLaughlin et al. 1983). The development of the Müllerian duct has been intensively studied by Lutz-Ostertag (1954). Figure 4.6 is based upon her data. In the 8-day-old Fig. 4.6 Contd. ...

started cranially. The right ovary is already smaller than the left. The mesonephros is well developed, and both Wolffian ducts (WD) are functional. C. Stage 40 HH (14 days) with regression of the cranial part of the right MD. D. Stage 45 HH (20 days). On the left side, dilatation of the caudal MD. On the right side, rudimentary duct without ostium abdominale. Reduced size of mesonephros, but the WDs are still present. E. Adult: The different parts of the oviduct have developed on the left side. On the right side, small appendix of rudimentary duct. The metanephros with upper, middle, and lower lobes is formed. The mesonepros is involuted. Only rudiments of the WD persist. Drawing after Lutz-Ostertag 1954 made by K. Barteczko.

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Reproductive Biology and Phylogeny of Birds

embryo (Fig. 4.6A), both ducts have equal length. The mesonephroi are well developed with both gonads symmetrically positioned at their medial sides. The first difference in length is seen in the 10-day-old embryo and becomes clearly visible in the 11-day-old embryo (Fig. 4.6B). While the left gonad has grown, the right gonad began to regress. After 14 days of incubation (Fig. 4.6C) the size of the right ovary is significantly reduced and the right Müllerian duct has regressed nearly completely with only its caudal part evident. But during the period of regression the abdominal ostium is suggested to remain open (Lutz-Ostertag 1954). The mesonephros is still functioning although some nephrons are involuted (Volle and Beaumont 1964), and both Wolffian ducts are well developed. Prior to hatching (Fig. 4.6D), the right oviduct is still rudimentary with a length of about 4.5 mm while the left one has grown steadily and has differentiated into several regions. The future shell gland (uterus) can be distinguished as a dilatation of the tract near its distal end. The mesonephroi are reduced in size with a big left and a small right ovary attached on the ventral and medial side. The Wolffian ducts are likewise preserved. Histological examination of the oviductal mucosa shortly after hatching (Fig. 4.7A-C) shows well-developed primary folds with secondary folding starting. Cell strands of the dense lamina propria contact the basal lamina. They appear to help in glandular formation. The nonciliated epithelium is irregular and consists of columnar or pseudostratified cells. Berg et al. (2001) demonstrated that exposure of female quail embryos to ethynyloestradiol induces precocious differentiation of the epithelium and tubular glands in immature birds In the adult chicken (Fig. 4.6E), the mesonephros is replaced by the metanephros. Rudiments of the Wolffian ducts might persist. The left ovary lies adjacent to the upper part of the kidney (metanephros) and ventral to the aorta. It has an irregular shape resulting from different growth of follicles. The left oviduct, which is straight and small before sexual maturity, increases enormously in length and in diameter in the breeding season. The oviduct differentiates into five morphologically distinct regions: infundibulum, magnum, isthmus, the prominent shell gland, and vagina. The caudal part, the vagina is separated from the cloaca proper by the occluding plate, which is ruptured after mating or at the onset of egg production. On the right side a rudimentary Müllerian duct persist as an appendix of the cloaca. The normal growth of the right and left female Müllerian duct is summarized in Table 4.1. The data are adapted from Teng (1987) and LutzOstertag (1954). The latter are measured in White Leghorn embryos. The differences in absolute values may be due to different methods or different strains. However the ratios are similar. An overview of the development of the oviduct in duck embryos was likewise presented by Lutz-Ostertag (1954). Up to day 9, the female Müllerian ducts are of equal length. The regression of the right duct starts in day-10

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Fig. 4.7 A. Longitudinal section of the left oviduct from a one-day-old chicken. The future magnum is dilated. The mucosa reveals primary folds with a dense lamina propria. B. Detail from a well-developed primary fold with secondary ones that appear leaf-like. C. The luminal epithelium is columnar and non-ciliated. Cells of the dense stroma attach the basal lamina. D. Transverse section through the distal region of the right oviduct and the rudimentary Wolffian duct (*) (WD) between ureter (U) and oviduct (O). The undifferentiated oviduct reveals a columnar epithelium with a thick basal lamina surrounded by a dense mesenchymal layer. E. Detail of the WD to show the well developed columnar epithelium and the small mesenchymal layer. Scale bars: A, B, D = 100 mm, C and E = 25 mm. Original.

$" Reproductive Biology and Phylogeny of Birds Table 4.1

The normal growth of Müllerian duct (MD) in female chick embryos

Days of incubation 6 8 10 11 14 16 21 The data

Length of left MD (mm)

Length of the right MD (mm)

3.8 3.8 5.3 5.3 (7.5) (7.5) 7.5 7.5 (10.6) (9.6) 9.0 7.8 (11.5) (8.7) 104 6.4 (17.9) (4.8) 19.2 4.7 (21.5) (4.6) 22.2 4.4 (28.0) (7.5) are taken from Teng (1987) and, in parenthesis, Lutz Ostertag (1954)

embryos in the same manner as in the chick embryo. Apoptotic bodies are visible reaching their maximum during days 14 and 15. The regression is terminated at the nineteenth day of incubation with a rudiment similar in appearance to that observed in the chick.

4.4

THE RIGHT MÜLLERIAN DUCT AFTER HATCHING

While the left oviduct is a length of 22 or 28 mm respectively (see Table 4.1) at the time of hatching, the right oviduct is rudimentary with a length of 4 to 9.3 mm according to authors such as Benoit 1950 (cited by Hlozankova and Zalenka 1978), Lutz-Ostertag 1954; Romanoff 1960; Teng 1987. Figure 4.7D shows a persistent right oviduct shortly after hatching. It is poorly differentiated with a columnar epithelium, a thick basal lamina, and a dense mesenchymal sheath. Observing 63 female chicks (hybrid combination Ross 1), Hlozankova and Zelenka (1978) found a rudimentary right oviduct in all animals. Their length and width increase in relation to the growing hen. At 15 months, their average length was of 93.1 mm and the diameter had enlarged three-fold (see Table 4.2). However, there was also significant variation in shape and size. Two narrowings separate an ampulla, which could be compared with the shell gland, from the caudal retroperitoneal part and from a thin cranial cap-like Table 4.2

Size of the right Müllerian duct (MD) after Hlozankova and Zelenka (1978)

Age in days Average length (mm) Average width (mm)

22

36

57

450

17.3 4.6

23.7 7.1

24.8 6.9

93.1 13.8

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part that extends into the meso (ligament). In some hens this upper part was separated from the ampulla by connective tissue. The rudiments are always filled with a clear liquid and in some cases they have dilated to a cystic structure. Interestingly, when the occluding plate is ruptured there is a clear fluid released from the vagina (unpublished observations, MB). Sell (1959) found well-developed right oviducts in 80% of the White Plymouth Rock hens examined. They possessed all regions of a normal oviduct, but in all cases the infundibulum was shorter than normal, and eggs were only found in the left oviduct. The incidence of a fully functional persistent right oviduct is described in White Leghorn by Bickford (1965), and in the ring necked pheasant by Purohit et al. (1977).

4.5 RUDIMENTS OF MESONEPHROS AND WOLFFIAN DUCT IN FEMALES AFTER HATCHING The mesonephros is functional until 3 days before hatching (Wendler 1965). The regression of mesonephric tubules continues up to the 3rd or 4th week after hatching (Budras 1972), but rudiments persist even in old adults. This has been described in detail by Kummerlöwe (1931) for many wild birds such as Turdus and Accipiter. Budras (1972) intensely investigated the remnants of mesonephros in Gallus. He always found cranial tubules (epoophoron) located between adrenal gland and ovary and some of them are supposed to grow into these adjacent organs. Inside the capsule of the adrenal gland and at the border between ovarian medulla and cortex they transform into interrenal (adrenal cortical-like) and interstitial nodules, respectively. The epoophoron seems to be an additional source of sex-steroid producing cells similar to the corpus luteum of mammals (Budras 1972). The pathological significance of epoophoron-derived tumors with virilisation of hens has been described by Boring and Pearl (1918). Because of the close relationship between the mesonephros and gonads, several authors (Witschi 1935, 1956, cited by Carlon et al. 1983; Carlon et al. 1983) claimed that, as in other vertebrates, the part of the chick embryo’s mesonephros which does not form nephrons contributes to the medullar cells of the gonads. Rodemer et al. (1986), using chick-quail chimeras, showed that from the third to the sixth segment, the ventromedial part of the differentiating mesonephros participates in the formation of stromal cells to the gonads. The mesonephric (Wolffian) ducts persist in female mammals as Gartner’s ducts and can be the source of hydatids of Morgagni (Motta 1966; Jacob and Barteczko 2005) or cysts. Glandular acini opening in Gartner’s duct were also found in the camel (Shehata 1978), and are discussed as accessory glands of the female genital tract. In female birds rudimentary Wolffian ducts persist as permanent structures running parallel with the ureter within the dorsal ligament of the oviduct. King (1975) stated that the adult hen possesses on the right side a potentially viable and complete system of the male duct including the rete, the epoophoron and the mesonephric duct, while the fate of the left

$$ Reproductive Biology and Phylogeny of Birds duct remains uncertain. In contrast to Domm (1927), Brode (1928) and Kar (1947) (all cited by King 1975) concluded that the left mesonephros and duct involuted nearly completely before hatching. We found in the one day-old female chicken (Fig. 4.7D, E) on both sides a patent Wolffian duct with a narrow lumen surrounded by a cuboidal or columnar epithelium and some layers of mesenchymal cells. At the beginning of the breeding season the Wolffian ducts respond to androgenic hormones produced by the female and might be prominent in many avian species (for references see Gilbert 1979). Yet, no information is available about the functional significance of such gland-like structures.

4.6 STRUCTURE OF THE HEN’S OVIDUCT DURING EGG PRODUCTION 4.6.1

Overview

The oviduct in mature birds in egg production is a relatively large tubular organ that has three primary functions all within the realm of reproduction. These include the following: egg formation; sperm selection, storage and transport; and, fertilization and early embryonic development. Briefly, at ovulation the fimbriated region of the infundibulum guides the ovum to the ostium abdominale, the opening of the oviduct. Then within minutes, sperm, if present, make contact with the inner perivitelline layer (IPVL) initiating the process of fertilization. Whether fertilized or not, the ovum accrues secretory material released from the infundibular mucosa forming the thin continuous layer and the more extensive outer perivitelline layer (OPVL). Within 1520 min, the ovum, now referred to as an egg-mass, enters the magnum, which is the largest of the five oviductal regions. The egg albumen proteins are synthesized in the magnal tubular glands and secreted (see Burley and Vadehra, 1989 for a comprehensive review on albumen composition and chemistry). After about 3 hr, the albumenous egg mass enters the isthmus for 75-90 min where the inner and outer shell membranes are formed. The egg mass then reaches the uterus (shell gland) and during the next 18-20 hr the calcareous shell is deposited and there is a movement of “plumping” fluid (uterine transudate) into the egg mass. At oviposition the hard-shelled egg mass is expelled through the vagina, which serves as a conduit between the uterus and cloaca. The vagina also plays a vital role in sperm selection, transport and storage. If fertilized, and depending on the species, the egg at oviposition contains a blastoderm with 20,000 to 60,000 cells. The time from ovulation to oviposition is referred to as the “ovulatory cycle” and in domestic birds it is about 23-25 hr. These events are summarized in Fig. 4.8 and presented in greater detail in Chapter 11 of this volume. To more fully understand and appreciate these functions, a comprehensive understanding of the tissue and cell structures and their interrelationships is necessary. The descriptions provided will focus on the anatomy, histology and to a lesser extent the ultrastructure of each oviductal segment. Unless otherwise stated, the description will be that of the commericial turkey. For

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Developmental Anatomy of the Female Reproductive Tract

Fig. 4.8 Drawings illustrating formation of the hard-shelled egg in Gallus gallus and the segments of the left oviduct corresponding to the different stages of the ovulatory cycle. A. Length of the segment in mm and schedule of the ovulatory cycle in minutes. B. Alteration of the egg mass during passage along the oviduct. C. Segments of the oviduct with transverse sections (D) and details of the mucosa (E). a, left ovary; b, infundibulum; c, funnel-like fimbriated region of the infundibulum showing an ovum (t) entering the ostium abdominale; d, tubular neck region of infundibulum; e, magnum; f, isthmus with pars translucens (g) and the tubular shell gland (h); I, uterus (shell gland); k, vagina; u, continuous and outer perivitelline layer; v, albumen; x, shell membrane; z, shell. (From: Komarek V, Malinovsky L, Lemez L: 1982. Anatomia avium domesticarum et embryologia galli. Part 2. Priroda, Bratislava,Tab. LVI, with permission of the publishers).

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$& Reproductive Biology and Phylogeny of Birds more detail discussion on the overall anatomy of the oviduct refer to Aitken (1971), Hodges, (1974), and King (1975). Anatomical nomenclature will follow that suggested in Handbook of Avian Anatomy: Nomina Anatomica Avium (Baumel, 1993).

4.6.2

Infundibulum

The infundibulum is the most anterior segment of the oviduct. It is divided into a funnel like region, the fimbria (also referred to as the ampulla), which grasps the ovulated ovum and guides it into the ostium abdominale, and the more elongated, tubular neck region (also referred to as the chalaziferous region). The neck gradually merges with the proximal segment of the magnum. If fertilization is to take place, sperm must be present in the funnel or upper neck region. The surface mucosa is thrown into a series of primary and smaller secondary folds clearly more voluminous and longitudinally orientated at the neck region than at the fimbria, which are shorter and more randomly orientated (Bakst and Howarth 1975). The mucosal folds in the neck region begin to form widely scattered subepithelial tubular glands just caudal to the fimbria (Fig. 4.9A, B). Moving caudally into the neck region the tubular glands proliferate into dense clusters and eventually mix with clusters of the larger tubular glands characteristic of the proximal magnum (Fig. 4.10A, B). Secretions derived from the infundibular tubular glands contribute to the formation of the continuous layer and OPVL. Together, these tertiary investments (formed by oviducal secretions) envelop and add strength to the IPVL as well as serving as a block to pathological polyspermy (see Chapter 11). The pseudostratified columnar epithelium lining the mucosa surface at the fimbriated region is composed of a dense array of ciliated cells that more caudally give rise to alternating ciliated and secretory cells in the neck (Fig. 4.10B). At the base of some neck folds, nonciliated cuboidal predominate and in cross section appear as ‘glandular grooves’. When these folds are apposed to an egg mass the nonciliated cells forming the glandular grooves are exteriorized and clearly visible (Fig. 4.9A). True tubular glands are observed throughout the neck region and resemble the tubular glands of the the magnum and isthmus (Fig. 4.10A, B). Structurally the tubular glands of the neck region are compound multicellular glands. The spherical secretory granules densely packed in the apical half of the secretory Fig. 4.9 A. When apposed to an ovum, the mucosa of the midregion of the infundibulum becomes distended exposing its nonciliated basal surface. An opening to a tubular gland is observed (arrow) in this scanning electron micrograph. After Bakst, M.R. 1978. Poult. Sci. 57: 1065-1069. Fig. 2. B. A squash preparation of unfixed infundibular mucosa from the same region as Fig. 4.9A is viewed by differential interference contrast (DIC) microscopy. With the plane of the optical section just subjacent to the surface epithelium portions of several tubular glands (arrows) are observed among the capillaries in the loose connective tissue. After Bakst, M.R.1994. Biology of Reproduction 50: 987-992, Fig. 5.

Developmental Anatomy of the Female Reproductive Tract

Fig. 4.9

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Fig. 4.10 A. A squash preparation of unfixed infundibular mucosa from caudal neck region is viewed by DIC microscopy. Secretory granules are abundant only in the supra-nuclear cytoplasm of the secretory cells permitting an unobstructed view Fig. 4.10 Contd. ...

Developmental Anatomy of the Female Reproductive Tract

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cells when secreted presumably are the source of the tertiary investment around the ovum. It has long been speculated that the infundibulum is a secondary sperm storage site (see Bakst et al., 1994 for review) but this role has been recently questioned (see Chapter 11). Regardless of the role of the infundibulum in sperm storage, it is the site of fertilization. To be successful, sperm must find their way to the 3 mm diameter germinal disc at the surface of the ovum, interact with the appropriate sperm receptors on the IPVL surface before the OPLV is formed.

4.6.3

Magnum

In egg production the magnum is responsible for the synthesis and secretion of the albumen proteins (see Burley and Vadehra 1989 for a discussion on albumen chemistry). Visually, the longitudinally orientated folds are high, voluminous and ivory in color prior to the passage of an egg mass. Immediately after passage, the folds appear somewhat diminished in size and have a pale reddish-brownish coloration. This volume change is due to the depletion of secretory material from the tubular glands. However, full volume and the ivory coloration of the folds are replenished prior to the next ovulation. The pseudostratified columnar epithelium lining the mucosa surface contains a near equal distribution of non-ciliated secretory and ciliated cells. Using transmission electron microscopy (TEM), its clear that the non-ciliated secretory cells are mucin secreting cells, reminescent of goblet cells (Blom 1973; Sandoz et al. 1976). Tubular glands are prominent in sections of the magnum and clearly present at all levels of the folds. Like the neck of the infundibulum, the tubular glands are compound multicellular glands lacking a transition between the tubular gland and surface epithelium (see Blom 1973). Toward the distal 2-3 cm of the magnum, the mucosal folds appear less voluminous and the goblet cells in the surface epithelium appear to be the dominant cell type. This is sometimes referred to as the mucous part of the magnum. The transition between the magum and isthmus is abrupt and easily recognized by eye as a 1-3 mm wide, nearly translucent band, the pars translucens.

4.6.4

Isthmus

Slightly shorter and smaller in diameter than the magnum, the isthmus synthesizes and secretes an array of proteins and glycoproteins that form the Fig. 4.10 Contd. ...

of the nuclei (arrow). B. From the same region as Fig. 4.10A, a section containing the ciliated surface epithelium (C) and several tubular glands are observed. Secretory granules are abundant only in the supra-nuclear cytoplasm of the secretory cells permitting an unobstructed view of the nuclei (arrows). Original.

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Reproductive Biology and Phylogeny of Birds

shell membranes. Given this mix, Burley and Vadehra (1989) point out that it was incorrect to refer to the shell membranes are being composed of keratins. The longitudinally orientated mucosal folds are somewhat smaller than the folds of the magnum and contain tubular glands endowed with a dense array of secretory granules. Like the magnum, the surface epithelium is composed of both non-ciliated secretory cells and ciliated cells arranged as a pseudostratified columnar epithelium. Interestingly, the openings of the tubular glands are slightly larger than those observed in the magnum and by SEM appear as dimples (Bakst and Howarth 1975) or shallow depressions (Blom 1973) distributed on the mucosal surface. There was disgreement in the literature regarding the caudal aspect of the isthmus and whether it should be considered part of the isthmus or uterus (Fig. 4.11A). The mucosal folds of the tubular shell gland resembles that of the isthmus. However, the tubular glands may (Richardson 1935) or may not (Aitkin 1971) differ from the uterine pouch. Solomon (2002) reviewed the arguments and suggested that the term “tubular shell gland” as it was commonly used. This term concurs with that found in Nomina Anatomica Avium (Baumel 1979) “Pars cranialis uteri” suggesting that the tubular segment caudal to the isthmus is the cranial aspect of the uterus.

4.6.5

Uterus

Also referred to as the shell gland, the uterus has two distinct regions, the more cranial tubular shell gland (see previous paragraph), and the pouch-like portion of the uterus. Less distinct is the recessus uterinus, described as a funnel shaped, greyish white color at the caudal end of the uterus (Baumel 1979). Its mucosa appears to be more a transitional zone between the uterovaginal junction (UVJ), the latter being the most cranial aspect of the vagina defined by the presence of sperm storage tubules (SST). These structures are best appreciated when the recessus uterinus, UVJ, and cranial half of the vagina are collectively dissected free of connective tissue and laid straight (Fig. 4.11A). Each region becomes clearly visible after they are liberated from the enveloping connective tissue. The mucosa folds are not clearly longitudinal but form leaf-like structures that seem to be arranged more circumferentially. This circumferential orientation is lost as the connective tissue band surrounding the pouch is dissected away. The pseudostratified columnar epithelium contains nearly an equal distribution of non-ciliated secretory cells and ciliated cells. The tubular glands of the uterine pouch are less voluminous than in the isthmus or magnum.

4.6.6

Vagina

The most caudal segment of the oviduct, the vagina, is coiled and folded and held in that complex configuration by a dense covering of connective tissue (Fig. 4.11B). This same connective tissue also binds the coprodeum, the most cranial compartment of the cloaca, in juxtaposition with the vagina. It should

Developmental Anatomy of the Female Reproductive Tract

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Fig. 4.11 A. After isolating and stripping off the connective tissue around the uterus and vagina, the recessus uterinus (oval) and extended vagina are observed. Also observed is the Pars cranialis uteri, the tubular shell gland (box). The uterovaginal junction is just caudal to the recessus uterinus, and is defined by the presence of sperm-storage tubules (SST). After Bakst, M.R. 1998. J. Exp. Zool. 282: 618-626, Fig. 4. B. The vagina (encircled) has been cut free of the connective tissue that envelops it with the coprodeum (upper arrow). However, the dense connective tissue still keeps the vagina as a tightly coiled structure. The lower arrow highlights the ostium cloacale oviductus sinistri (Baumel 1993), the opening of the left oviduct. After Bakst, M.R. 1993. Chapter 2, Manipulation of the Avian Genome, Fig. 4.

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Fig. 4.12 A. A squash preparation of unfixed UVJ mucosa viewed by DIC microscopy showing the midregion of a sperm storage tubule containing a few sperm. Basal nuclei (arrows) and more apical lipid droplets characterize this simple columnar epithelium. After Bakst, M.R. 1993. Chapter 2, Manipulation of the Avian Genome, Fig. 10. B. The complexity of the mucosal folds in the distal vaginal is observed in this section. Note where the folds are tightly apposed the lumen is not discernible (arrow). Original.

Developmental Anatomy of the Female Reproductive Tract

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be noted that with the removal of the connective tissue binding the distal uterus, vagina and coprodeum and subsequent straightening of the vagina, at no time was there observed a thickening of the Tunica muscularis (MB, personal observations). Thus, the existence of a vaginal sphincter around the cranial aspect of the vagina analogous to the mammalian cervix (Baumel 1979) should be questioned. About 24 nearly parallel primary folds are longitudinally orientated and are further subdivided into secondary and tertiary folds. The complexity of the mucosal folds is best appreciated in histological sections (Fig. 4.12B). Tightly apposed folds with seemingly interdigitating ciliated surfaces were often observed with one or more sperm between the cilia. By SEM, the mucosa surface appears to be densely ciliated although nonciliated secretory cells are also observed in its pseudostratified columnar epithelium. At the juncture of the distal vagina with the urodeum, the central zone of the cloaca in which the urogenital ducts empty, the ciliated epithelium of the vagina ends rather abruptly and is replaced with the secretory (mucin) cells of the urodeum. Unlike the other oviductal segments, the vaginal segment does not possess tubular glands. However, the UVJ at the cranial end of the vagina is defined by the presence of SST (Fig. 4.12A) These differ from the tubular glands seen elsewhere in the oviduct as the SST are composed of simple columnar nonsecretory cells characterized by a basal nuclei and an accumulation of intracellular lipid. A more detailed account of SST structure and function is found in Chapter 11. The development of the SST was recently examined by Holm and Ridderstrale (2002) in quail and found to coincide with the development of the oviduct at maturation. Sperm-storage tubule numbers, spatial distribution, and morphology (Birkhead and Moller 1998) as well as the spatial and temporal patterns of sperm filling and emptying (Briskie 1996; Bakst and Vinyard 2002) appear to vary between and within species.

4.7 LITERATURE CITED Abdel-Malek, E. T. 1950. Early development of the urogenital system in the chick. Journal of Morphology 86: 599-626. Aitken, R. N. C. 1971. The Oviduct. Pp.1237-1289. In D. G. Bell and B. M. Freeman (eds), Physiology and Biochemistry of the Domestic Fowl. Volume 3. (Academic Press, London. Allard, S., Adi, P., Gouédard, L., di Clemente, N., Josso, N., Orgebin-Crist, M-C., Picard, J-Y. and Xavier, F. 2000. Molecular mechanisms of hormone-mediated Müllerian duct regression: involvement of b-catenin. Development 127: 3349-3360. Austin, H. B. 1989. Müllerian duct regression in the American alligator (Alligator mississippiensis). Its morphology and testicular induction. Journal of Morphology 251: 329-338. Austin, H. B. 1995. DiI analysis of cell migration during Müllerian duct regression. Developmental Biology 169: 29-36. Baarends, W. M., van Helmond, M. J., Post, M., van der Schoot, P. J., Hoogerbrugge, J. W., de Winter, J. P., Uilenbroik, J. T., Karels, B., Wilming, L. G., Meijers, J. H., Themmen, A. P. and Grootegoed, J. A. 1994. A novel member of the

%$ Reproductive Biology and Phylogeny of Birds transmembrane serine/threonine kinase receptor family is specifically expressed in the gonads and in the mesenchymal cells adjacent to the Müllerian duct. Development 120: 189-197. Bakst, M. R. and Howarth, B. 1975. SEM preparation and observations of the hen’s oviduct. Anatomical Record 181: 211-226. Bakst, M. R., Wishart, G. J., and Brillard, J. P. 1994. Oviducal sperm selection, transport, and storage in poultry. Poultry Science Reviews 5: 117-143. Bakst, M. R. and Vinyard B. T. 2002. Oviducal sperm storage in turkeys: spatial distribution of sperm within the uterovaginal junction sperm-storage tubules. Journal of Experimental Zoology 292: 205-209. Balfour, F. M. and Sedgwick, A. 1878. On the existence of a rudimentary head-kidney in the embryo chick. Proceedings of the Royal Society 27: 443-446. Baumel, J. J., King, A. M, Breazile, J. E., Evans, H. E., and Vanden Berge, J. C. (eds.) 1993. Handbook of Avian Anatomy: Nomina Anatomica Avium, Second Edition, Nuttall Ornithological Club, Cambridge MA. 779 pp. Behringer, R. R. 1995. The Müllerian inhibitor and mammalian sexual development. Philosophical Transactions of the Royal Society of London. Series B. Biological Sciences 350: 285-289. Berg, C., Holm, L., Brandt, I. and Brunström, B. 2001. Anatomical and histological changes in the oviducts of Japanese quail, Coturnix japonica, after embryonic exposure to ethynyloestradiol. Reproduction 121: 155-165. Bickford, A. A. 1965. A fully formed and functional right oviduct in a single comb White Leghorn pullet. Avian Disease 22: 464-470. Birkhead, T. R. and Moller, A. P. (eds). 1998. Sperm Competition and Sexual Selection. Academic Press, New York. 282 pp. Blom, L. 1973. Ridge pattern and surface ultrastructure of the oviducal mucosa of the hen (Gallus domesticus). Det Kongelige Danske Videnskabernes Selkab Biologiske Skrifter 20: 1-40. Boring, A. M. and Pearl, R. 1918. Sex studies. XI. Hermaphrodite birds. Journal of Experimental Zoology 25: 1-48. Budras, K-D. 1972. Das Epoophoron der Henne und die Transformation seiner Epithelzellen in Interrenal- und Interstitialzellen. Advances in Anatomy, Embryology and Cell Biology 46(3). Springer, Berlin. 74 pp. Burley, R. W. and Vadehra, D. V. 1989. The Avian Egg: Chemistry and Biology. Wiley Interscience-John Wiley and Sons, New York. 472 pp. Carlon, N., Pizant, J. and Stahl, A. 1983. Mesonephric origin of the gonadal primitive medulla in chick embryos. Anatomy and Embryology 166: 399-414. Carré-Eusèbe, D., di Clemente, N., Rey, R., Pieau, C., Vigier, B., Josso, N. and Picard, J-Y. 1996. Cloning and expression of the chick anti-Müllerian hormone gene. The Journal of Biological Chemistry 271: 4798-4804. Clarke, T. R., Hoshiya, Y., Yi, S.E., Liu, X., Lyons, K. M. and Donahoe, P. K. 2001. Müllerian inhibiting substance signaling uses a bone morphogenetic protein (BMP)-like pathway mediated by ALK2 and induces Smad6 expression. Molecular Endocrinology 15: 946-959. Didier, E. 1973. Recherches sur la morphogénèse du canal de Müller chez les Oiseaux I. Etude descriptive. Wilhelm Roux’ Archiv 172: 271-286. Dyche, W. J. 1979. A comparative study of the differentiation and involution of the Müllerian duct and Wolffian duct in the male and female fetal mouse. Journal of Morphology 162: 175-209.

Developmental Anatomy of the Female Reproductive Tract

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Forsberg, J. G. and Olivecrona, H. 1963. Degeneration processes during the development of the Muellerian ducts in alligator and female chicken embryos. Zeitschrift für Anatomie und Entwicklungsgeschichte 124: 83-96. Gilbert, A. B. 1979. Female genital organs. Pp. 237-360. In A.S. King and J. McLelland (eds), Form and Functions in Birds. Volume 1. Academic Press, London, New York, Toronto, Sydney, San Francisco. Gruenwald, P. 1941. The relation of the growing Müllerian duct to the Wolffian duct and its importance for the genesis of malformations. The Anatomical Record 81: 1-19. Guichard, A., Cedard, L., Mignot, T. M., Scheib, D. and Haffen, K. 1977. Radioimmunoassay of steroids produced by cultered chick embryonic gonads: Differences according to age, sex and side. General and Comparative Endocrinology 32: 255-265. Ha, Y., Tsukada, A., Saito, N. and Shimada, K. 2004. Changes in mRNA expression of MMP-2 in the Müllerian duct of chicken embryo. General and Comparative Endocrinology 139: 131-136. Hamburger, V. and Hamilton, H. L. 1951. A series of normal stages in the development of the chick embryo. Journal of Morphology 88:49-92. Hamilton, H. L. 1965. Lillie’s Development of the Chick. An Introduction to Embryology. 3rd edition. Holt, Rinehart and Winston, New York. 624 pp. Hiruma, T. and Nakamura, H. 2003. Origin and development of the pronephros in the chick embryo. Journal of Anatomy 203: 539-552. Hlozankova, E. and Zelenka, J. 1978. Right Müllerian duct in the domestic fowl during postnatal ontogenesis. Anatomischer Anzeiger 144: 208-213. Hodges, R. D. 1974. The Histology of the Fowl. Academic Press, London, 648 pp. Holm, L. and Ridderstrale, Y. 2002. Development of sperm storage tubules in the quail during sexual maturation. Journal of Experimental Zoology 292: 200-205. Hutson, J., Ikawa, H. and Donahoe, P. K. 1981. The ontogeny of Müllerian inhibiting substance in the gonads of the chicken. Journal of Pediatric Surgery 16: 822-827. Hutson, J., Ikawa, H. and Donahoe, P. K. 1982. Estrogen inhibition of Müllerian inhibiting substance in the chick embryo. Journal of Pediatric Surgery 17: 953-959. Hutson, J. M., Fallat, M. E. and Donahoe, P. K. 1984. The fate of the grafted quail Müllerian duct in the chick coelom. Journal of Pediatric Surgery 19: 345-352. Hutson, J. M., Donahoe, P. K. and MacLaughlin, D. T. 1985. Steroid modulation of Müllerian duct regression in the chick embryo. General and Comparative Endocrinology 57: 88-102. Jacob, H. J., Jacob, M. and Christ, B. 1977. Die Ultrastruktur der externen Glomerula. Ein Beitrag zur Nierenentwicklung bei Hühnerembryonen. Verhandlungen der Anatomischen Gesellschaft 71: 909-912. Jacob, H. J., Jacob, M. and Christ, B. 1979. Feinstrukturelle Untersuchungen zur Entwicklung der Vorniere von Hühnerembryonen. Verhandlungen der Anatomischen Gesellschaft 73: 547-554. Jacob, H. J., Jacob, M. and Christ, B. 1986. The early development of the intermediate mesoderm in the chick. Pp 61-68. In R. Bellairs, D. A. Ede and J. W. Lash (eds), Somites in Developing Embryos. NATO ASI Series, Series A: Life Sciences, Volume 181. Plenum Press, New York. Jacob, M. and Barteczko, K. 2005. Contribution to the origin and development of the appendices of the testis and epididymis in humans. Anatomy and Embryology 209: 287-302.

%& Reproductive Biology and Phylogeny of Birds Jacob, M., Konrad, K. and Jacob, H. J. 1999. Early development of the Müllerian duct in avian embryos with reference to the human. An ultrastructural and immunohistochemical study. Cells Tissues Organs 164: 63 -81. Jacob, M., Jacob, H. J., Barteczko, K. and Süß, B. 2000. Early development of urogenital system in vertebrates especially origin of Wolffian and Müllerian duct. Journal of Egyptian German Society of Zoology 32: 58-73. King, A. S. 1975. Aves urogenital system. Pp 1919-1964. In R. Getty (ed.), Sisson and Grossman’s The Anatomy of the Domestic Animals. Volume 2, 5th edition. Saunders, Philadelphia. Kinsky, F. C. 1971. The consistent presence of paired ovaries in the kiwi (Apteryx) with some discussion of this condition in other birds. Journal of Ornithology, Leipzig 112: 334-357. Kümmerlöwe, H. 1931. Vergleichende Untersuchungen über das Gonadensystem weiblicher Vögel. 3. Ausgewählte Beispiele aus verschiedenen Vogelordnungen. Zeitschrift für mikroskopisch-anatomische Forschung 24: 455-631. Lasala, C., Carré-Eusèbe, D., Picard J-Y. and Rey, R. 2004. Subcellular and molecular mechanisms regulating anti-Müllerian hormone gene expression in mammalian and nonmammalian species. DNA Cell Biology 23: 572-585. Lutz-Ostertag, Y. 1954. Contribution a l’étude du développement et de la régression des canaux de Muller chez l’embryon d’oiseau. Bulletin Biologique de la France et de la Belgique 88: 333-412. MacLaughlin, D. T., Hutson, J. M. and Donahoe, P. K. 1983. Specific estradiol binding in embryonic Müllerian ducts: a potential modulator of regression in the male and female chick. Endocrinology 113: 141-145. Motta, P. 1966. Contributo alla struttura ed ultrastruttura dell’epitelio dell’idatide del Morgagni. Archivio di ostetricia e ginecologia. (Napoli) 71:473-496. Oréal, E., Pieau, C., Mattei, M. G., Josso, N., Picard, J-Y., Carré-Eusèbe, D. and Magre, S. 1998. Early expression of AMH in chicken embryonic gonads precedes testicular SOX9 expression. Developmental Dynamics 212: 522-532. Oréal, E., Mazaud, S., Picard, J-Y., Magre, S. and Carré-Eusèbe, D. 2002. Different patterns of anti-Müllerian hormone expression, as related to DMRT1, SF-1, WT1, GATA-4, Wnt-4, and Lhx9 expression, in the chick differentiating gonads. Developmental Dynamics 225: 221-232. Purohit, V.D., Basrur, P. K. and Reinhardt, B.S. 1977. Persistent right oviduct in ringnecked pheasant. British Poultry Science 18: 177-178. Rey, R., Lukas-Croisier, C., Lasala, C. and Bedecarras, P. 2003. AMH/MIS: what we know already about the gene, the protein and its regulation. Molecular and Cellular Endocrinology 211: 21-31. Richardson, K.C. 1935. The secretory phenomena in the oviduct of the fowl, including the process of shell formation examined by microincineration techniques. Philosophical Transactions of the Royal Society of London 225B: 149195. Roberts, L.M., Hirokawa, Y., Nachtigal, M. W. and Ingraham, H. A. 1999. Paracrinemediated apoptosis in reproductive tract development. Developmental Biology 208: 110-122. Roberts, L.M., Visser, J.A. and Ingraham, H.A. 2002. Involvement of a matrix metalloproteinase in MIS-induced cell death during urogenital development. Development 129: 1487-1496. Rodemer, E.S., Ihmer, A. and Wartenberg, H. 1986. Gonadal development of the chick embryo following microsurgically caused agenesis of the mesonephros and

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using interspecific quail-chick chimaeras. Journal of Embryology and Experimental Morphology 98: 269-285. Romanoff, A. A. 1960. The Avian Embryo. Macmillan, New York. Sandoz, D., Boisvieux-Ulrich, E., and Delaunay, M. C. 1976. Ciliogenese dans les cellules a mucus de l’oviducte de caille. I. Etude ultrastructurale chez la caille en ponte. Journal of Cell Biology 71: 449-459. Saino, K. 2003. Development of mesonephric kidney. Pp 75-86. In P. D. Vize, A. S. Woolf and J. B. L. Bard (eds). The Kidney. From Normal Development to Congenital Disease. Academic Press, New York. Sell, J. 1959. Incidence of persistent right oviducts in the chicken. Poultry Science 38: 33-35. Semon, R. 1890. Über die morphologische Bedeutung der Urniere in ihrem Verhältnis zur Vorniere und Nebenniere und über ihre Verbindung mit dem Genitalsystem. Anatomischer Anzeiger 5: 455-482. Semon, R. 1892. Studien über den Bauplan des Urogenitalsystems der Wirbeltiere. Jenaer Zeitschrift für Medizin und Naturwissenschaften 26 NF 19: 89-203. Shehata, R. 1978. Comparative study of Gartner’s duct in the camel. Acta Anatomica 100: 532-537. Solomon, S. E. 2002. The oviduct in chaos. World’s Poultry Science Association 58: 4148. Starck, D. 1999. Embryologie. Ein Lehrbuch auf allgemein biologischer Grundlage. 2nd edition. Georg Thieme Verlag, Stuttgart, Germany. 693 pp. Teng, C. S. 1987. Quantification of Müllerian inhibiting substance in developing chick gonads by a competitive enzyme-linked immunosorbent assay. Developmental Biology 123: 255-263. Teng, C. S. 2000. Differential expression of c-Jun proteins during Müllerian duct growth and apoptosis: caspase-related tissue death blocked by diethylstilbestrol. Cell Tissue Research 302: 377-385. Trelstad. R. L., Hayashi, A., Hayashi, K. and Donahoe, P. K. 1982. The epithelialmesenchymal interface of the male rat Müllerian duct: loss of basement membrane integrity and ductal regression. Developmental Biology 92: 27-40. Visser, J. A. 2003. AMH signaling: from receptor to target gene. Molecular and Cellular Endocrinology 211: 65-73. Volle, G. and Beaumont, A. (1964) Phénomènes histologiques de la régression du mésonephros de l’embryon de Poulet. Comptes rendus des séances de la Société de biologie et de ses filiales 158: 5-7. Wartenberg, H. 1985. Morphological studies on the role of the periductal stroma in the regression of the human male Müllerian duct. Anatomy and Embryology 171: 311-323. Wendler, D. 1965. Der histochemische Aktivitätswandel des proximalen Urnierennephrons während der Entwicklung des Hühnchens. Zeitschrift für Anatomie und Entwicklungsgeschichte 124: 478-503. Wolff, Et. 1939. L’action du diéthylstilboestrol sur les organes génitaux de l’embryon de Poulet. Comptes rendus hebdomadaires des séances de l’Académie des sciences 209: 1532-1533. Woods, J. E. and Erton, L. H. 1978. The synthesis of estrogens in the gonads of the chick embryo. General and Comparative Endocrinology 24: 413-423. Wrobel, K. H. and Süß, F. 2000. The significance of rudimentary nephrostomial tubules for the origin of the vertebrate gonad. Anatomy and Embryology 201: 273-290.

n n

CHAPTER

5

Endocrinology of Reproduction G.E. Bentley1, K. Tsutsui2 and J.C. Wingfield3

5.1

INTRODUCTION

Birds reproduce in virtually all habitats on earth from the poles to the equator and alpine meadows to the driest deserts. Because all avian species are oviparous, they are restricted to land for breeding purposes. Thus, the only major ecosystem in which birds are unable to nest is the ocean. Nonetheless, marine habitats provide rich trophic resources for birds nesting on islands and coastal areas. Restriction of the reproductive process to the oviparous mode might suggest that birds have an equally restricted hormone control system. Compared with other vertebrate taxa this may be true but the complexity of avian breeding cycles and the broad spectrum of habitats in which they nest indicates great flexibility in these endocrine control systems. We approach the hormonal control systems of the class Aves in four major parts: 1) types of environmental signals that influence reproduction and how they are perceived; 2) the hypothalamus as an integrator of environmental information from the external and internal environments, biological clock, etc., that through neuroendocrine and neural secretions affects all aspects of reproduction; 3) the hypothalamo-pituitary unit that transduces environmental information processed by higher centers into endocrine secretions; 4) the functional gonads (ovary and testis) themselves. We do not provide much detail on the gonadal histology as this topic is covered in chapters 2, 4 and 7 in this book. This is an exciting era in neuroendocrinology as new peptides discovered in recent years provide even more flexibility in control mechanisms. This is timely in an age when global climate change, human disturbance and pollution require organisms to be even more flexible in how they cope with 1

Department of Integrative Biology, University of California, Berkeley, California 94720, USA. 2 Faculty of Integrated Arts and Sciences, Hiroshima University, Higashi-Hiroshima 7398521, Japan. 3 Department of Biology, University of Washington, Box 351800, Seattle, Washington, 98195, USA.

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such events. We hope that this review will focus new attempts to explore the breeding strategies and cycles of vertebrates in general.

5.2

ENVIRONMENTAL SIGNALS AFFECTING REPRODUCTION

Environmental signals that are indicative of transitions of life history stages (such as breeding) required in the near or immediate future can be used to trigger appropriate responses of the individual (Wingfield et al. 1999; Wingfield 2004). In other words, environmental signals can be used to: (a) Orchestrate development (ontogeny) of organisms and respond to environmental changes that require adjustment of developmental trajectories. Maternal effects are also important at this time. (b) Regulate development, maintenance and regression of morphological, physiological and behavioral traits characteristic of life history stages (e.g. migration, breeding, molt) throughout the adult life cycle. (c) Regulate when adjustments of homeostasis are to be made in advance of predictable changes in the environment. (d) Signal when alternate behavioral and physiological patterns should be triggered in response to unpredictable and disruptive events. (e) Adjust social interactions to maximize an individual’s ability to gain access to shelter, food resources, territories, mates etc. There is a complex cascade of events involved in the perception, integration and transduction of environmental signals into chemical signals (e.g. hormones, neurotransmitters) that then initiate appropriate morphological, physiological and behavioral adjustments (Ball 1993). There are five major processes involved: 1. Specialized receptors for specific external environmental signals that then transduce them into neural events. 2. Neural pathways from the specialized receptors project to target areas in the brain that integrate the environmental information in relation to internal physiological state (biological clock, social status; nutritional state, disease etc.). 3. These target areas of the brain then signal specific neurons in the hypothalamus to release neurosecretions that initiate a hormonal cascade that in turn triggers appropriate adjustments of morphology, physiology and behavior. 4. Neurosecretions and enzymes within the CNS can have separate regulatory effects on neurosecretions from the hypothalamus into the blood circulation. Note also that internal signals (from the biological clock, social status; nutritional state, disease etc.) may also act at the level of neurosecretions within the CNS. 5. Peripheral, blood borne, hormones (especially steroids) may feedback to act on the brain and alter responsiveness to environmental signals at either the specialized receptor, target brain area, or neurosecretory

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neuron levels. Again, these effects may be further regulated by the biological clock, social status; nutritional state, disease etc. It should also be borne in mind that possible neural pathways for environmental signals may be very different among the types of signal (e.g. food, temperature, social etc., Fig. 5.1). Furthermore, these neural pathways may not always be stimulatory. Some may inhibit neurosecretion, others may release an inhibition rather than stimulate directly. Clearly an almost limitless

Fig. 5.1 Proposed pathways of transduction for proximate information that may regulate reproductive function. In homeothermic vertebrates such as birds, direct effects of environmental cues on peripheral endocrine cells are not common (an example is the effect of gut contents on the hormone secreting cells of the gastric mucosa, Norris 1997). Most environmental information from the physical environment, social interactions and internal cues are transduced through the hypothalamo-pituitary unit. Note that some environmental cues may be signalized directly through central (CNS) or autonomic (ANS) nervous systems (e.g. behavioral responses to social interactions, ejaculation when mating, oviposition etc.). Others may be signalized through neuroendocrine secretions directly (i.e. from the pars nervosa). Most appear to be transduced through the hypothalamopituitary unit and have cascading effects on physiology, morphology and behavior associated with reproductive development, actual nesting and finally termination of breeding. An example of this would be the effects of changing photoperiod on GnRH and gonadotropin release that directly regulate gonadal function. Original.

&" Reproductive Biology and Phylogeny of Birds spectrum of specialized receptors, neural pathways and neuromodulator mechanisms exist, but we still know very little about these critical first steps in response to environmental signals. Next, the steps in the cascade of events that takes place after an individual is exposed to environmental signals, and the hormonal cascades that follow will be defined further. The types of environmental signals regulating transitions of life history states may vary markedly. For example, signals emanating from other individuals may be perceived by sensory receptors and transduced by areas of the CNS that are very different from physical environmental signals. Exactly how an organism perceives relevant environmental signals, and the influences those signals have, can be classified in two major ways. Firstly the effects may be directly on cells that respond with a physiological or morphological response (e.g. gut cells responding to changes in food composition, e.g. Norris 1997) or are signalized through the CNS (Fig. 5.1). Environmental cues perceived by the CNS allow the individual to respond neurally (e.g. via the autonomic nervous system) or by the neuroendocrine/endocrine system (i.e. secrete hormones). In the latter case neural signals are transduced into neuroendocrine secretions that then influence the rest of the endocrine system by blood borne hormones (Wingfield et al. 1999). External cues that are signalized may be perceived by various sensory modalities—visual, tactile, auditory, chemical (taste/smell) electrical, deep brain photoreceptors and baroreceptor. Many other sensory receptors remain to be identified such as those for detection of temperature responses, humidity etc. In addition to external cues, internal signals also have important influences on the neuroendocrine and endocrine systems. Examples are blood and stored levels of amino acids, free fatty acids, sugars, vitamins etc. Other examples include osmoregulatory functions (salt/water balance), the immune system and disease, endogenous rhythms (“internal clock”) and aging. One can imagine many potential internal signals. Even here, however, the mechanisms by which such signals are perceived may not always be intuitive.

5.2.1 The Reproductive Life History Stage and Its Organization There are three major phases to any life history stage (Jacobs and Wingfield 2000; Wingfield 2004; Fig. 5.2). The development phase consists of maturation of the reproductive system itself, as well as the activation of reproductive behaviors including establishment of a breeding territory, attraction of a mate etc. The mature capability phase (Fig. 5.2) is when actual nesting can begin. At this time the gonads are mature and females are ready to begin the final maturation of ovarian follicles leading to yolk deposition and ovulation. There are two major sub-phases, sexual and parental, with complex patterns of substages within them that can be repeated when re-nesting after failure of one breeding attempt, or when raising multiple broods within a season. Interactions between these sub-phases occur when the sexual phase terminates (the clutch is complete) and parental behavior begins. Because the

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Fig. 5.2 Phases and sub-stages of the reproduction life history stage in birds. The development phase consists not only of development of the reproductive system itself, but also establishment of a breeding territory, attraction of a mate etc. The mature capability phase is when actual nesting can begin. There are two major sub-phases, sexual and parental, with complex patterns of sub-stages within them that can be repeated when re-nesting after failure of one breeding attempt, or when raising multiple broods within a season. Finally the reproduction life history stage is terminated and the reproductive system regresses (often to a completely infantile state), reproductive behaviors wane and the next life history stage (e.g. molt) develops. Modified from Wingfield, J. C., Jacobs, J. D., Tramontin, A. D., Perfito, N., Meddle, S., Maney, D. L. and Soma, K. 1999. Pp. 85-128. In K. Wallen and J. Schneider (eds), Reproduction in Context. M.I.T. Press, Cambridge, Massachusetts, Fig. 4.2.

reproductive system remains mature throughout this process, switches between sub-phases and the sub-stages involved (e.g. courtship and copulation, nest building, incubation, feeding young etc.) can be rapid. Finally the reproduction life history stage is terminated (Fig. 5.2) and the reproductive system regresses (often to a completely infantile state), reproductive behaviors wane and the next life history stage (e.g. molt) develops.

&$ Reproductive Biology and Phylogeny of Birds

5.2.2 Classification of Environmental Signals Affecting Reproduction The spectrum of environmental signals known to influence reproduction in vertebrates is truly daunting. However, it is possible to classify these factors into three major types (Fig. 5.3, Wingfield 1980, 1983; Wingfield et al. 1999, note that the signals from unpredictable events are not included here). Initial predictive information (e.g. the annual change in day length) triggers the development phase of the reproductive life history stage, maintains mature capability and then regulates termination at the end of the breeding season (Fig. 5.3). Local predictive information (e.g. temperature, food supply, rainfall) may inhibit or accelerate effects of initial predictive information. These cues are also important to time onset of actual nesting at mature capability, and adjust timing of the termination phase (Fig. 5.3). All social interactions are clustered under synchronizing and integrating information that can influence the development phase and support functions such as territorial behavior, synchronize mates as they begin nesting and integrate transitions between sub-phases such as sexual to parental behavior (Wingfield 2006, Fig. 5.3).

Fig. 5.3 Different types of environmental signals from the physical environment and from social interactions can influence all three phases of the reproduction life history stage. Initial predictive information initiates gonadal development, maintains reproductive capacity and then terminates the breeding season. Local predictive and synchronizing and integrating information can influence all three phases too. Thus knowing phase of the reproduction life history stage will be an important determinant of how, for example, a given behavioral interactions will affect neuroendocrine and endocrine mechanisms. Modified from Wingfield, J. C., Jacobs, J. D., Tramontin, A. D., Perfito, N., Meddle, S., Maney, D. L. and Soma, K. 1999. Pp. 85-128. In K. Wallen and J. Schneider (eds). Reproduction in Context. M.I.T. Press, Cambridge, Massachusetts, Fig. 4.3.

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There is also strong evidence that behavioral interactions regulate the termination phase. Note how types of environmental signals act at different phases of the reproductive life history stage. This may indicate very different mechanisms by which a specific type of environmental signal may affect different phases (Wingfield et al. 1999; Wingfield 2004).

5.3 THE HYPOTHALAMUS AS A TRANSDUCER OF ENVIRONMENTAL CUES INTO HORMONAL SIGNALS The avian hypothalamus and the hypothalamo-pituitary unit are functionally composed of both the nervous and endocrine systems. They have been described in depth frequently (e.g. Wingstrand 1954; Oksche and Farner 1974; Mikami 1986; Scanes et al. 1984) and show clear homologies with other vertebrate taxa. A photomicrograph of this region in the White-crowned Sparrow (Zonotrichia leucophrys gambelii) is shown in Fig. 5.4.

Hypothalamus

The hypothalamus extends from the lamina terminalis, rostral to the optic chiasm, to the supramamillary decussation. The base and lateral walls of the third ventricle provide useful markers (Fig. 5.4). This region has been divided into preopticohypothalamic, tuberal, and mamillary regions (e.g. Kuenzel and van Tienhoven 1982), and most of the hypothalamic nuclei, neuroendocrine

Fig. 5.4 Sagittal section of the hypothalamo-pituitary unit in the white-crowned sparrow, Zonotrichia leucophrys gambelii. Modified from Farner, D. S., Wilson, F. E. and Oksche, A. 1967. Pp. 529–545. In L. Martini and W. F. Ganong (eds), Neuroendocrinology, vol. 2. Academic Press, New York, Fig. 2.

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5.3.1

&& Reproductive Biology and Phylogeny of Birds fibers, tracts and associated glial and ependymal cells are contained within it. Neuroendocrine cells within the hypothalamus project to the pars nervosa and the median eminence where they form terminal fields next to blood capillaries (Fig. 5.5, from Farner et al. 1967; Wingfield and Farner 1993). Avian hypothalamic nuclei are rather diffuse, perhaps numbering 15-20, although only a few have been well studied (Fig. 5.5, from Farner et al. 1967). Note also that regions of the avian telencephalon and diencephalon have undergone a major revision of nomenclature (Reiner et al. 2004). It is through these brain regions that many environmental signals perceived by sensory receptors are received as neural transmissions. These signals are converted into neuroendocrine secretions that then influence morphology, physiology and behavior (Scanes et al. 1984; Ball 1993; Ball and Balthazart 2002). Most environmental information from the physical environment, social interactions and internal cues are transduced through the hypothalamopituitary unit. Note that some environmental cues may be signalized directly through central (CNS) or autonomic (ANS) nervous systems (e.g. behavioral responses to social interactions, ejaculation when mating, oviposition etc.). Others may be signalized through neuroendocrine secretions directly (i.e. from the pars nervosa). Most appear to be transduced through the hypothalamopituitary unit and have cascading effects on physiology, morphology and behavior associated with reproductive development, actual nesting and finally termination of breeding. An example of this would be the effects of

Fig. 5.5 Schematic diagram of the major components of the hypothalamic-pituitary unit in birds (c.f. Fig. 5.1) and diagram of some avian hypothalamic nuclei. Modified after Farner, D. S., Wilson, F. E. and Oksche, A. (1967). Pp. 529-582. In L. Martini and W. F. Ganong (eds.), Neuroendocrinology, vol. 2. Academic Press, New York, Fig. 1.

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changing photoperiod on gonadotropin releasing hormone (GnRH) and other peptides (Fig. 5.6, see below) and gonadotropin release that directly regulate gonadal function (Fig. 5.6).

5.3.2

Median Eminence

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The median eminence is where neurohormones from hypothalamic neuroendocrine cells are released into the hypophysial blood portal system to stimulate or inhibit release of anterior pituitary hormones. This is a major switch point as environmental information relayed through hypothalamic neurons triggers a hormonal cascade that affects many aspects of an organism’s life cycle. In birds the median eminence is formed by the ventral floor of the hypothalamus, emerging posteriorly from the optic chiasm and forming the infundibular stalk (Figs. 5.4 and 5.5). It receives axons from hypothalamic nuclei and also contains glial cells. There are specialized ependymal cells that terminate on portal capillaries in juxtaposition with terminals of axons (Figs. 5.4, 5.5 and 5.7). Additional neurons of hypothalamic origin may also be found but they are not neuroendocrine (Kobayashi et al.

Fig. 5.6 A schematic representation of how signalized environmental information transduced by the brain results in secretion of hypothalamic peptides such as avian gonadotropin-releasing hormone-I (GnRH-I) and Lamprey GnRH-III (lGnRHIII) that stimulate gonadotropin release. Note that avian gonadotropin-inhibitory hormone (GnIH) inhibits at least luteinizing hormone secretion. These cascading effects regulate gonadal development and function such as secretion of sex steroids, avian inhibins etc. Original.

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' Reproductive Biology and Phylogeny of Birds 1970). Typically, hypothalamic neuron terminals filled with neurosecretory granules are separated from primary capillaries by two basement membranes (Fig. 5.7). These capillaries, that drain to the hypophysial portal system, have many fenestrations of the endothelial cells as found in capillaries of other endocrine tissues. Note that the perivascular spaces between the basement membrane and the capillary endothelium also contain cells and sometimes erythrocyctes (Fig. 5.7, Mikami et al. 1970). There are also pericytes that orient spirally around endothelial cells (Fig. 5.7). They may have a function as a mechanical support of the portal vessels as well as a contractile function regulating blood flow (Mikami et al. 1970). Additionally, the endothelial cells may have protrusions into the capillary lumen that have been suggested to have a role in regulation of blood flow. In some species, the median eminence and the primary portal capillaries are encased by the pars tuberalis of the adenohypophysis (Kobayshi et al. 1970; Vitums et al. 1964; Mikami et al. 1970).

5.3.3

Hypothalamic Nuclei and Tracts

Secretory neurons in the hypothalamus synthesize a number of hormones that are released into the peripheral blood circulation (e.g. those of the pars nervosa) and have effects largely on contraction of smooth muscle and water balance in the kidney. Neurosecretions from the median eminence enter a specialized blood portal system that vascularizes the pars distalis (anterior pituitary or adenohypophysis). These secretions either stimulate or inhibit the synthesis and secretion of hormones from endocrine cells within the pars distalis. They are commonly referred to as “releasing” or “inhibiting” hormones or factors (Scanes et al. 1984). The hypothalamic neurons, often clustered in groups (nuclei), project their axons to the pars nervosa and/or the median eminence (Fig. 5.5) forming tracts and terminal fields where axons may contact basement membranes of capillaries (Fig. 5.7). The median eminence narrows caudally into the tubular infundibular stalk terminating in the pars nervosa (Figs. 5.4 and 5.5). Neurosecretory material is transported through this stalk via the supraoptico-paraventriculo-hypophysial tract to the pars nervosa (see Wingstrand 1951, 1954; Oksche and Farner 1974 for detailed review). Many tracts have been indentified in the avian hypothalamus (Wingstrand 1951). Important examples are (1) the tractus supraoptico-hypophyseus, which originates in the supraoptic region and terminates in the pars nervosa; (2) the tractus tuberohypophyseus projecting from the infundibular nucleus to the median eminence; (3) the tractus hypophyseus posterior, mostly receiving fibers from the posterior hypothalamus; and (4) the tractus hypophyseus anterior, consisting of fibers from nuclei in the lateral hypothalamus, and from the paraventricular and preoptic areas that terminate in the median eminence. It is likely that there are many differences among species. The systematic use of immunocytochemical methods and in situ hybridizations for mRNA coding for peptides may resolve these issues.

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Fig. 5.7 A. Electron micrograph of anterior median eminence (AME) and primary capillary (PCp) of a photostimulated white-crowned sparrow, Zonotrichia leucophrys gambelii. B. Enlargement of primary capillary (PCp) that is widely extended on the surface of the median eminence and separated from it by two layers of basement membrane (Bm). Fenestrations (arrows) of the endothelial cells (En) are evident. A Pericyte (Pc) with a large nucleus is in the lower left corner. Perivascular spaces (PVS) are found between the basement membrane and the capillary endothelium. From Mikami et al. (1970). Zeitschrift für Zellforschung 106: 155-174. Springerverlag, Berlin, Fig. 2.

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5.3.4

Reproductive Biology and Phylogeny of Birds

Hypothalamo-hypophysial Portal System

There are two networks of primary portal capillaries arising separately from the ventral surfaces of the anterior and posterior divisions of the median eminence (Fig. 5.5). Anterior and posterior portal veins may be enclosed as a bundle by the pars tuberalis of the adenohypophysis, flowing to secondary capillary plexuses and sinuses in the rostral and caudal lobes respectively (Fig. 5.5, e.g. Vitums et al. 1964, 1966; Duvernoy et al. 1969, 1970; Singh and Dominic 1975). It is assumed that this portal system delivers neurohormones from neurons in specific areas of the hypothalamus to discrete areas in the pars distalis of the adenohypophysis (Vitums et al. 1966), although as yet there is no actual physiological evidence that supports this hypothesis (Wingfield and Farner 1993). The portal vessels may be the sole vascular supply to the pars distalis, and there is no unequivocal evidence of direct hypothalamic or other innervation, although recently such projections have been claimed to exist. The late Donald S. Farner described this hypothalamo-hypophysial system very well as follows: “defined hypothalamic nuclei contain cells of different functions despite their near-uniform appearance when examined by classical cytological and cytochemical techniques. The same may hold for fiber systems within anatomically defined tracts. A crude analogy can be drawn with industrial cities, railways and highways, and destinations. Factories (perikarya) in several cities (hypothalamic nuclei) produce a variety of products (neurohormones) some of which are produced in more than one city. It then follows that railways and highways (axons and tracts) are involved in delivery of a variety of products (neurohormones etc.) to different destinations—neurohemal organs, synapses, cerebrospinal fluid etc. The analogy is of course incomplete because nothing is included about communication and transport of materials to the factories” (from Wingfield and Farner 1993).

5.3.5

Pars Nervosa

The caudal portion of the infundibulum of the hypothalamus, just posterior to the median eminence, forms the pars nervosa (Figs. 5.4 and 5.5, Wingstrand 1951; Oksche and Farner 1974). The lumen may be open or closed and the neuroendocrine pars nervosa contains branched terminals and fibers of the supra-optico-paraventriculo-hypophysial tract, glial cells, pituicytes, and ependymal cells. The pars nervosa is a neurohemal organ, like the median eminence, from which hypothalamic neurohormones can be transferred into circulating systemic blood (Wingfield and Farner 1993). In birds there are two pars nervosa hormones—arginine vasotocin (AVT) and mesotocin (MT). Magnocellular bodies in the pre-optic, supra-optic and paraventricular regions of the hypothalamus synthesize these peptides and immunocytochemical techniques show each cell body expresses one hormone—either mesotocin-like or vasotocin-like. Note that all these cell types are found in the

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hypothalamic nuclei containing magnocellular neurons (Oksche and Farner 1964; Wingfield and Farner 1993).

5.3.6

Hormones of the Pars Nervosa

Mesotocin (Ile-8-oxytocin, MT) and arginine vasotocin (AVT) were first demonstrated in extracts of posterior pituitaries in Domestic fowl (Gallus gallus), Turkey (Meleagris galloparvo) and Goose (Anser anser). Both MT and AVT are amidated nonapeptides with a disulphide bridge and amino acid substitutions at positions 3 and 8 (Acher et al. 1993 for review). They are found in all flying birds studied to date as well as ratites, e.g. Ostrich (Struthio camelus) (Lazure et al. 1987; Michel et al. 1990a). The genes also express neurophysins that have specific properties of binding neurohypophysial hormones, carry them down the axon to terminal fields, and play a role in their release by exocytosis. Neurophysins are of two major types and appear to occur on a one-on-one molar ratio with the hormones they carry. Neurophysin I binds mesotocin. Neurophysin II binds AVT (George 1980; Michel et al. 1990ab; Hamann et al. 1992; Acher et al. 1993). Neurophysins are synthesized in perikarya, transported axonally, stored and released with the hormones themselves. They have a high degree of homology of amino acid sequence with mammalian neurophysins. The precursor molecules themselves are biologically inactive. Enzymatic cleavage produces the active peptides in the Golgi apparatus and in secretory granules prior to release. Neurophysin bound to hormone requires about 2-3 hours to migrate from the magnocellular neurons to terminal fields in the pars nervosa. In general, AVT and MT have the following functions (see George 1980 for review): 1. Water balance, through flow in semipermeable membranes. 2. Regulation of blood pressure. 3. Contraction of smooth muscle in gonadal ducts. 4. Effects on sexual behavior and aggression (Goodson and AdkinsRegan 1999). Note that actions 2 and 3 are on contraction or relaxation of smooth muscle. AVT has well known actions on contraction of smooth muscle, especially in the oviduct at oviposition. In support of this view, plasma levels of AVT increase at oviposition in chickens perhaps mediated by prostaglandins in oviductal tissue (see George 1980; Acher et al. 1993 for reviews). In addition to the action of AVT, galaninergic innervation of the uterine oviduct may be involved in oviposition in quail (Li et al. 1996; Tsutsui et al. 1997, 1998; Sakamoto et al. 2000). There is also evidence that injections of AVT in pigeons increase plasma levels of free fatty acids, glucose and growth hormone (George 1980). The biological effects of MT in birds are less well known. Both MT and AVT can act as neurotransmitters since such neurons project to many sites in the CNS including the median eminence (Acher et al. 1993). AVT has also been shown to increase male sexual behavior in the pigeon (Kihlström and Danninge 1972) and male zebra finch (Taenopygia guttata) (Goodson and Adkins-Regan 1999) and on aggression (Goodson 1998a,b; Goodson and Adkins-Regan 1999), as has been well demonstrated in other vertebrates such as amphibia (e.g. Moore 1987). More recently, there is

'" Reproductive Biology and Phylogeny of Birds some evidence that AVT can act to attenuate behavior- and crowding-induced aggression via a cardiovascular mechanism, rather than a receptor-mediated central mechanism, in European starlings (Sturnus vulgaris) (Nephew et al. 2005a). The mechanism of action of MT in birds is not well known.

5.4 ADENOHYPOPHYSIS (= ANTERIOR LOBE OF THE PITUITARY GLAND) Unlike the pars nervosa (neurohypophysis), the adenohypophysis is not part of the brain but derived from pharyngeal tissue and becomes associated with the median eminence during embryonic development (Wingstrand 1951; Gorbman et al. 1983; Norris 1997). The avian adenohypophysis consists of the pars tuberalis and pars distalis. There is no pars intermedia (Wingstrand 1951; Gorbman et al. 1983; Norris 1997).

5.4.1

Pars Distalis and Pars Tuberalis

Cells of the pars distalis are typically arranged in cords (Fig. 5.8). Using histological and cytochemical techniques a system of putative cytophysiological functions for cells of the avian pars distalis has been developed (for extensive reviews see Tixier-Vidal and Follett 1973; Mikami 1986; Mikami et al. 1975). The early investigations on the functions of these cells involved surgical removal of gonads, thyroid glands, and adrenals. Later, the morphology of secretory granules and other ultrastructural features of secretory cells were studied using electron-microscope techniques. Now immunocytochemical and in situ hybridization techniques provide more definitive evidence. Here we will focus on the gonadotropin and prolactin secreting cells as the major types directly involved in control of reproduction. Histological investigations with light and electron microscopy combined with reproductive state and following castration, lead to the conclusion that there are two types of gonadotropes in the rostral and caudal lobes of the pars distalis. We now know that these cell types secrete the gonadotropins folliclestimulating hormone (FSH) and luteinizing hormone (LH) that regulate gonadal function (e.g. Goldsmith and Follett 1980; Norris 1997; Kirby et al. 2005). These conclusions seem to be at least partially sustained because in chickens, in situ hybridization studies show that FSH and LH appear to be produced by different cells. In situ hybridization of mRNAs of the b subunit of LH match immunocytochemical localization in pituitaries at different stages of reproduction in white-crowned sparrows (Fig. 5.9, e.g. Kubokawa et al. 1994). Recent immunocytochemical evidence indicates that prolactin secreting cells (lactotropes) are found primarily in the rostral lobe of the pars distalis (Mikami 1986). Note that the term lactotrope is misleading in birds because most species do not lactate (except for crop milk in columbiiformes), and prolactin has many other actions (Wingfield and Farner 1993; Norris 1997).

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Fig. 5.8 Area of the caudal lobe (A and C) and cephalic lobe (B and D) of an adult white-crowned sparrow, Zonotrichia leucophrys gambelii. Note cells arranged in cords. From Matsuo, S.-I., Vitums, A., King, J. R. and Farner, D. S. (1969). Zeitschrift für Zellforschung 95: 143-176. Springer-verlag, Berlin, Fig. 3.

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Fig. 5.9 In situ hybridization for LH b-subunit mRNA (right) and immunocytochemistry of LH (left) in anterior pituitaries of White-crowned Sparrows, Zonotrichia leucophrys gambelii. SD = short day (16L 8D) LD = long day (20L 4D). From Kubokawa, K., Ishii, S. and Wingfield, J. C. (1994). General and Comparative Endocrinology 95: 42-51. Academic Press, New York, Fig. 6.

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5.4.2

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Hormones of the Pars Distalis and their Targets

There are three major types of hormones in the pars distalis: the glycoproteins, the pro-opiomelanocortin derived peptides, and the polypeptides. Some have direct actions on morphology, physiology and behavior, whereas others have tropic effects on peripheral endocrine glands. A detailed account of all three types is beyond the scope of this review and we will focus on gonadotropins and prolactin. There are three glycoprotein hormones secreted by the pars distalis and all have both early and late (trophic) cell responses. Luteinizing hormone (LH) regulates sex steroid production as well as ovulation. Follicle-stimulating hormone (FSH) promotes spermatogenesis and ovarian follicle maturation. It also regulates secretion of a gonadal protein inhibin, that feeds back negatively to regulate FSH secretion. Thyroid-stimulating hormone (TSH) regulates thyroid hormone synthesis and secretion (see Goldsmith and Follett 1980; Norris 1997). All three hormones share a characteristic structure of two subunits, a and b. The a subunit is shared by all three hormones whereas the b subunit is specific. For example, if subunits are disassembled, it is possible to recombine an FSH a subunit with a TSH b subunit and still get good TSH biological activity. The b subunit provides specificity, but association of a and b subunits is essential for binding to receptor and biological action (e.g. Gorbman et al. 1983; Baeulieu and Kelly 1990; Norris 1997). There are two related polypeptides secreted by the pars distalis, one is growth hormone that regulates growth (through release of insulin-like growth factors—IGFs), as well as lipid metabolism. The other is prolactin that has a vast spectrum of actions including reproductive function, lipid metabolism and osmoregulation (Goldsmith and Follett 1980; Norris 1997).

5.4.3

The Hypothalamo-Adenohypophysial-Gonad Axis

A summary of the interrelationships among the brain, anterior pituitary and gonad is presented in Fig. 5.6. This follows the general vertebrate system (e.g. Gorbman et al. 1983; Wingfield and Farner 1993). The scheme can be referred to as a summary guide and quick reference for the complex relationships with morphology, physiology and behavior discussed below. Synthesis and release of the gonadotropin LH is regulated by a hypothalamic releasing factor—gonadotropin-releasing hormone, GnRH (also called luteinizing hormone releasing hormone, LHRH, see Dunn and Millam 1998; Sharp and Ciccone 2005 for reviews). Hypothalamic control of FSH secretion is less clear because injections of GnRH have little effect, at least in chickens (Kirby et al. 2005). Evidence for GnRH in birds comes from measurements of biological activity and content of the avian hypothalamus by determining the ability of hypothalamic extracts to release LH in vitro. In some cases the released LH was measured by a second bioassay (e.g. 32P uptake in chick testis (e.g. Erickson 1975) in White-crowned sparrows (Zonotrichia leucophrys gambelii) or by radioimmunoassay either in the

'& Reproductive Biology and Phylogeny of Birds incubation medium (e.g. Bicknell and Follett 1975) in Japanese quail (Coturnix japonica), or in blood after injection of hypothalamic extracts in vivo in Z. l. gambelii (Wingfield and Farner 1993). In the 1980s, it was found that the hypothalamus of the domestic fowl had two distinct GnRH molecules, designated chicken GnRH-I and GnRH-II (cGnRH-I and -II) that differed from mammalian GnRH by one and three amino acid substitutions (King and Millar 1982a,b; Miyamoto et al. 1984). It has since been shown that the two chicken GnRHs are also found in the hypothalamus of wild species, European starling, and Song sparrow (Melospiza melodia) with cGnRH-I predominating (Sherwood et al. 1988). The cDNA encoding for the pre-pro-GnRH molecule has been sequenced for the domestic fowl (Dunn et al. 1993). Both have potent effects on release of LH in male and female Song sparrows in a more or less dose dependent fashion. However, in the domestic fowl, cGnRH-I and -II had different potencies in releasing LH (Sharp et al. 1987). In general, cGnRH-I is located in the pre-optic region of the brain with axonal projections (fibers) to the median eminence, where the terminal fields are located. The oculomotor complex of the midbrain is the location of the cGnRH-II neurons, which are typically less polar in shape than the cGnRH-I neurons and as a result have few processes extending from the cell bodies (for distribution see Millam et al. 1993, Millam et al. 1998; Teruyama and Beck 2000). Despite a long-held acceptance of the notion that cGnRH-II neurons are not hypophysiotropic, there have been some reports in quail that cGnRH-II irfibers are present in the POA, the lateral septum (Millam et al. 1993), and the median eminence (D’Hondt et al. 2001; Millam et al. 1998; van Gils et al. 1993; Clerens et al. 2003). However, there has only been one published report of cGnRH-II in the ME of songbirds (Stevenson and MacDougall-Shackleton 2005), while others find no evidence of GnRH-II in the same area (Meddle et al. 2006), and experiments in chickens suggest that the median eminence is a site of release of cGnRH-I, but not cGnRH-II, into the hypophysial portal vasculature (Sharp et al. 1990). Thus, the distribution and function(s) of cGnRH-II have remained somewhat enigmatic, although it should be noted that central administration of cGnRH-II, but not cGnRH-I enhances copulation solicitation in female white-crowned sparrows (Maney et al. 1997). Thus, cGnRH-II might act in a neurotransmitter role to influence reproductive behaviors, as has also been postulated for some mammalian species (Temple et al. 2003; Kaufmann and Rissman 2004). Hypothalamic hormones released from the median eminence circulate in the portal system and are quickly degraded before they reach the peripheral circulation. Avian LH and FSH were first purified from adenohypophyses of the domestic fowl by Stockell-Hartree and Cunningham (1969), and later by Furuya and Ishii (1974), and Sakai and Ishii (1980). Bioassay studies show that as in mammals, LH acts primarily on the endocrine gonad and FSH on the gametogenic gonad (e.g. Brown et al. 1975; Furuya and Ishii 1976; Brown and Follett 1977; Maung and Follett 1977; Follett et al. 1978). FSH acts in synergy with testosterone on spermatogenesis. Note that in Zonotrichia l.

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gambelii and many other photoperiodic songbirds testis size shows large variation in size with season (e.g. Wingfield and Farner 1993). In the regressed, non-breeding testis, semiferous tubules are inactive and regressed and cells of Leydig are undifferentiated. However, in the reproductively active males, seminiferous tubules are greatly enlarged and have bunches of spermatozoa. Leydig cells are active but not conspicuous between the greatly enlarged tubules. The epididymis, deferent duct (vas deferens) and seminal glomus show marked development in the breeding season. The cloacal protuberance (vent) can also become greatly enlarged, but this varies with species (see Birkhead and Møller 1992 for review). Using purified avian gonadotropins, specific radioimmunoassays (RIAs) have been developed for chicken LH (Follett et al. 1972), Turkey LH (Burke et al. 1979); and for chicken FSH (Scanes et al. 1977; Sakai and Ishii 1985). With the exception of the FSH assay developed by Scanes et al. (1977) these gonadotropin RIAs have proved to be applicable to plasma samples from a wide range of avian species (see Follett et al. 1978; Goldsmith and Follett 1980; Sakai and Ishii 1985, 1986). Isolation of LH and FSH from truly wild species appears to be restricted to the ostrich in which LH and FSH fractions have potent activity in both avian and mammalian bioassy systems (Bona-Gallo et al. 1983). Furthermore, LH from the ostrich is far more potent than FSH in stimulating secretion of testosterone from immature Mallard (Anas platyrhynchos) testes suggesting that as in domesticated species, increased androgen secretion is highly specific for LH (Chase 1982). Putative cDNA sequences for chicken LH b-subunit and Japanese quail LH b- and a-subunits have been cloned (Noce et al. 1989; Ando and Ishii 1994). The predicted amino acid sequences reveal some homology with mammalian forms of LH including the positions of proline residues which Ishii (1988, 1990) suggests may be related to species specificity of LH and its receptor. The chicken cDNA clone has been used to measure changes in mRNA for b-LH subunit during photostimulated gonadal growth in Zonotrichia l. gambelii (Kubokawa et al. 1994). The cDNA sequence encoding the extracellular component of LH receptor in testes of Japanese quail has also been characterized. This domain of the receptor is thought to be important for binding to the LH molecule. The predicted sequence of amino acids was about 70% homologous with mammalian LH receptor sequences, although the positions of cysteine residues and potential N-linked glycosylation sites were virtually identical to those of mammals (Akazome et al. 1994). Such conservation suggests extensive homology of molecular conformation of the LH receptors across phylogeny. A number of investigations (summarized in Lofts and Murton 1968, 1973; Wingfield and Farner 1993) as well as others, indicate that cells of Leydig are present in all phases of the annual cycle of periodically breeding species, and that their numbers and activity correlate reasonably well with sexual behavior. This conclusion is consistent, with few exceptions, with measured plasma levels of LH and testosterone, e.g. in Zonotrichia l. gambelii (Wingfield

 Reproductive Biology and Phylogeny of Birds and Farner 1978a,b), European starling (Ball and Wingfield 1984), and others (see Wingfield and Farner 1993; Wingfield et al. 1999 for reviews). Steroid biosynthesis in avian gonads is essentially similar to that of mammals. In all steroid secreting cells the precursor molecule is cholesterol (Furr and Pope 1970). The actions of LH after binding to a receptor on a steroid synthesing cell is to activate adenylate cyclase resulting in an increase in cAMP levels in the cell (e.g. Kubokawa et al. 1994). cAMP then activates protein kinases resulting in phosphorylations of proteins associated with mobilization of cholesterol droplets with the cell. The critical first step is enzymatic cleavage of the cholesterol side chain to give pregnenolone (e.g. Ozon 1972a). Next is the conversion of the hydroxyl group at carbon 3 to a ketone group to give progesterone under the action of an enzyme D5, 3bhydroxysteroid-dehydrogenase (HSD). Histochemical methods to visualize activity of this enzyme were used widely in the past as an indication of steroid synthesis because this enzyme catalyzes an important step from biologically inactive to active steroid hormones. Progesterone may, in some circumstances and in specific cells, be secreted itself. In other cases it it hydroxylated at carbon 17 to give 17ahydroxyprogesterone (e.g. Furr and Pope 1970; Ozon 1972a) that is then converted to androstenedione by cleavage of the side chain and dehydrogenase to give a ketone group at carbon 17. Simple hydroxylation at carbon 17 then gives testosterone that may be secreted (but note that it can be converted to various metabolites in the periphery, see below). Both androstenedione (since it is so easily converted to testosterone) and testosterone may also be substrate for an enzyme aromatase that results in aromatization of ring A (with loss of carbon 19, and hydroxylation at carbon 3) to give estradiol-17b (e.g. Ozon 1972b). Aromatase is widespread in avian tissues including gonads, steroid sensitive secondary sex characters, and brain (e.g. Schlinger and Arnold 1991,1992). The most common metabolite of estradiol-17b is estrone. The latter is generally regarded as being much less potent than estradiol, although plasma levels of estrone are sometimes measured. The avian ovary is unusual in that only one develops in the vast majority of species. This is usually the left ovary, although in Apterygidae both ovaries are apparently functional (see Lofts and Murton 1973; Welty and Baptista 1988). A cohort of ovarian follicles is recruited each breeding season and they develop to a stage in which they contain white yolk, but are still far from reaching an ovulatory state (King et al. 1966; Kern 1972; Lofts and Murton 1973). For each laying sequence (or clutch) a sub-cohort of these partially developed follicles begin sequestering yellow yolk from blood. This process is called vitellogenesis. Vitellogenin is synthesized in the liver and is stimulated by estradiol-17b. It is a dimer molecule of about 480 kDa containing complexes of protein-bound phosphorus and lipoproteins. The liver also produces other lipid rich proteins that are precursors to yolk (Griffin et al. 1984). Note that once these yolk precursors reach the ovary they must pass through the capillary wall as well as the basal lamina of the granulosa and granulosa

Endocrinology of Reproduction



cells themselves that surround the oocyte. Yolk precursors appear to be transported by receptor mediated endocytosis (Griffin et al. 1984). Ovarian follicles grow rapidly as yolk is deposited in layers (Kern 1972; Lofts and Murton 1973; Griffin et al. 1984) in a sequence according to size and they are ovulated one at a time as they mature (Wells and Gilbert 1984). Usually one follicle is ovulated each day (although there is great variation in ovulation times—see Welty and Baptista 1988). The ovulated egg is “captured” by the ostium (also known as infundibulum—not to be confused with the infundibulum of the hypothalamus) of the oviduct. Fertilization and deposition of the first layer of albumen occur here. The ovum passes down the oviduct through highly differentiated regions that have specific functions. Further albumen is laid down in the magnum, and membranes surround the developing egg in the isthmus. On reaching the shell gland (also known as uterus) a shell and pigment is deposited (e.g. Solomon 1983). Finally oviposition occurs through the vagina and cloaca. Note that since there is only one ovary and one oviduct, further ovulations cannot occur unless oviposition has occurred (e.g. Sharp 1980). It is possible that medullary and cortical interstitial cells of the theca are structurally identical, separate designations for them may be useful only because they may be functional at different times. Studies of the ovary show follicles as well as the theca and granulosa layers of follicles having steroid secreting capacity. Interstitial cells in the ovarian stroma also may secrete steroids. Atretic follicles also occur because not all follicles of a cohort selected for development are actually ovulated. Once the clutch is complete, other follicles with white yolk that have never entered the final maturation pathway, degenerate. Typically they collect lipid droplets and regress. Histochemical techniques that visualize 3bHSD (the enzyme that regulates conversion of pregnenolone to progesterone—and thus the conversion of a biologically inactive steroid to an active one) show potential steroid synthesizing activity of theca, granulosa and stromal cells (for reviews see Gilbert 1971; van Tienhoven 1983; Wells and Gilbert 1984; Johnson and Tilly 1990). Evidence suggests that interstitial cells of the theca interna and/or theca granulosa of the follicle secrete estrogens, progesterone, and testosterone. Indeed, Huang et al. (1979) have proposed a model that attributes an interaction between these two types of cells in the secretion of these hormones involving an interaction of LH and FSH on theca and granulosa cells respectively. The sex steroids testosterone, and also estradiol provide negative feedback signals to the gonadotropes in the pars distalis as well as GnRH neurons in the hypothalamus to inhibit GnRH and gonadotropin secretion. There is a marked decrease in sensitivity to steroid feedback at least in male Zonotrichia l. gambelii during gonadal maturation that allows a general increase in gonadotropin and sex steroid secretions (Matt 1983).



Reproductive Biology and Phylogeny of Birds

5.4.4 Actions of FSH on Follicular Development —Ovarian Growth Factors Inhibin, a peptide first demonstrated in the Sertoli cells of seminiferous tubules, is under the control of FSH and can inhibit the secretion of FSH from the pars distalis. There is also evidence that granulosa cells may secrete a substance similar to inhibin. This action seems to be specific for inhibition of FSH release while LH secretions remains unchanged. Since then there is now extensive evidence for an inhibin-activin system in the regulation of ovarian folliculogenesis in birds. They are members of a conserved transforming growth factor b super family that play intragonadal roles via autocrine and paracrine actions. As yet, there is no evidence that activins act on the pituitary (Knight et al. 2005). There is also evidence for paracrine effect of a growth factor (GDF9) from the oocyte itself that also plays an important role in oocyte development—possibly during the yolk uptake phase (Johnson et al. 2005).

5.5

PINEAL BODY (EPIPHYSIS CEREBRI)

The pineal gland (epiphysis) is located in the dorsal brain and made up of a stalk and rudimentary photoreceptors complete with visual pigments (rhodopsins). In birds there is a reduction in sensory neurons, but some sympathetic aminergic innervation may remain. This innervation is involved in the regulation of melatonin secretion—the main product of the pineal gland. Melatonin is an indoleamine synthesized from serotonin by the action of Nacetyltransferase to N-acetylserotonin and by hydroxyindole-Omethyltransferase (HIOMT) to melatonin (e.g. Gorbman et al. 1983; Norris 1997). At least in diurnal species melatonin is secreted at night (i.e. light inhibits release) and injections of melatonin can reverse activity rhythms. Transplants of pineal glands from individuals entrained to one rhythm can induce a similar rhythm in the recipient suggesting some autonomy. Indeed, pineals cultured in vitro can retain rhythmicity and even entrain to changing photoperiod (Cassone and Menaker 1984). Note, however, that the retina can also produce melatonin and thus the eyes may also be important. The avian pineal body is a circumventricular organ that shows great morphological diversity (e.g. Quay and Renzoni 1963; Quay 1965; Renzoni 1965), and in general consists of a stalk that arises from the roof of the diencephalon in the region between the habenular and posterior commissures, expanding dorsally to become the pineal organ. It thus lies close to the interior surface of the dorsal wall of the brain case in a triangular area delimited by the posterior margins of the cerebral hemispheres and the anterior margin of the cerebellum. In some species such as owls (Strigidae), the pineal gland is lacking or is a degenerate remnant (Breucker 1967; Renzoni 1968). The avian pineal has been studied in no less than 100 species of 13 orders (for cytological details on some selected wild species (see Quay and Renzoni 1963; Oksche and Kirschstein 1969; Ueck 1979).

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Few investigations addressed annual changes in pineal function, but in the House sparrow (Passer domesticus) Ralph and Lane (1969) found no evidence for seasonal changes. However, Barfus and Ellis (1971) found that the activity of pineal hydroxyindole-O-methyl transferase of P. domesticus was lowest during the breeding season. Similar results were obtained in the Baya weaver (Ploceus phillipinus) (Saxena et al. 1979) and the Indian tree pie (Dendrocitta vagabunda) (Chandhuri and Maiti 1989). Although direct illumination apparently does not alter electrical activity of the avian pineal body (Ralph and Dawson 1968; Morita 1975; Homma et al. 1980), secretion of melatonin is an endogenous circadian function that is entrained by photoperiod (e.g. Ralph et al. 1967; Ralph 1976; Binkley et al. 1977; Menaker et al. 1978; Cassone and Menaker 1984). Chicken pineal cells in culture show an endogenous oscillator that controls the circadian rhythm of serotonin N-acetyltransferase activity and contain a rhodopsin-like photoreceptor (Deguchi 1982). Although pineal activity may be affected by light in some species, whether this can be viewed as evidence that this organ is directly photosensitive in all has been questioned (Oliver and Baylé 1982) and there is now evidence suggesting extensive interspecific differences (cf. Cassone and Menaker 1984). There are several investigations claiming both anti- and progonadal effects of the avian pineal (for critical review see Binkley 1988). Unlike in mammals, there is little evidence of a significant direct role of the pineal in the regulation of gonadal function in birds. However, recently published findings point to the possibility that melatonin is, to some degree, involved in regulation of reproduction [see later section on gonadotropininhibitory hormone (GnIH)]. Pinealectomy of male Baya weavers resulted in increased rate of photoperiodically induced testicular growth (Balasubramanian and Saxena 1973), and when mature males were transferred to short days, the pinealectomized group showed partial testicular regression in comparison with complete regression in the sham operated group. The authors interpreted these results as effects of removal of an inhibitory effect of melatonin on the hypothalamo-hypophysial-gonadal axis. However, because the pineal body is a component of the avian circadian system (Cassone and Menaker 1984), it is possible that a shift in phase angle between a daily cycle in photosensitivity and the daily photoperiod could explain the results (e.g. Gwinner et al. 1981). Other components of the avian circadian system (Gaston and Menaker 1968) include the suprachiasmatic nucleus, retina, and deep encephalic (hypothalamic) photoreceptors (e.g. Cassone and Menaker 1984; Underwood and Siopes 1985), all have a common developmental origin in the diencephalon, and they are photoreceptive or receive photic information (Menaker 1982; Cassone and Menaker 1984; Underwood and Siopes 1985). More recently there has been a renaissance of interest in the possibility that melatonin is involved in seasonal processes in birds, including regulation of the reproductive axis. In a study involving the use of melatonin antiserum, Ohta et al. (1989) concluded that melatonin is involved in at least the initial stages of photoperiodism in quail (i.e., early in the dark phase), and that the

" Reproductive Biology and Phylogeny of Birds timing of suppression of plasma melatonin is critical to gonadal development. In a contradictory study, injecting melatonin into quail exposed to long day lengths (and thereby simulating a short-day melatonin signal) did not inhibit the long-day-induced gonadal growth (Juss et al. 1993). The fact that one study (Ohta et al. 1989) took the approach of shortening the melatonin signal during short days and another (Juss et al. 1993) lengthened the melatonin signal during long days might prove fundamental to the differences in results, especially if it is true that the timing of the melatonin signal is critical to the ensuing effect. It is possible that studies on melatonin and the reproductive axis in birds have been hampered by the modes and timing administration of melatonin, making interpretation of the findings difficult at best. At any rate, it seems obvious that the role of melatonin in reproduction in birds is somewhat more complex than its role in mammalian reproduction. Whatever its actions are, it is likely that melatonin affects a suite of physiological systems in addition to the reproductive system. For example Guyomarc’h et al. (2001) suggest that melatonin supplementation might affect food intake and fat deposition and thereby be responsible for the slight inhibition of sexual development that they observed in European quail (Coturnix coturnix). Perhaps the most convincing, and least confusing, effects of melatonin upon the avian reproductive system stem from two recent studies. The first study indicates that in chickens the effects of melatonin on plasma LH can be described very simply (Rozenboim et al. 2002). These authors demonstrated very large reductions in plasma LH (up to 70%) after injections of relatively high doses of melatonin into castrated white leghorn roosters. Additional experiments in that study indicated that melatonin administration inhibited plasma LH in a time- and dose-dependent manner, and that the effects persisted for as long as melatonin was administered (Rozenboim et al. 2002) . The second study is described in section 5.7.2 (Ubuka et al. 2005), and is concerned with the dose-dependent regulation of gonadotropin-inhibitory hormone by melatonin.

5.6 5.6.1

SEASONAL BREEDING, THYROID HORMONES AND PROLACTIN Photoperiodism and Photorefractoriness

In northern temperate zones, most bird species have evolved mechanisms to coincide their breeding with periods when environmental conditions are optimal for the raising of young. Often this depends on seasonal food supply and/or climatic conditions. In general, the further north an individual lives, the more imperative it becomes to coincide sexual maturity and raising of young with favorable conditions and to avoid breeding during less suitable periods. Any environmental factor that predictably signals the onset of a change in ecological conditions can act as a “proximate factor”—as long as it can be perceived and measured by the organism. The most reliable proximate factor is the changing length of photoperiod throughout the annual cycle. Other proximate factors as rainfall, food availability and temperature may

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advance or delay the breeding period to varying degrees (Gibb 1950; Kluijver 1951; Immelman 1971; Wingfield 1980, 1983; Wingfield et al. 1983; Wingfield and Farner 1993). Thus, most non-tropical birds have a very well-defined annual breeding season which is regulated precisely by changing photoperiod. This was first demonstrated in the junco (Junco hyemalis) by Rowan in 1925. These types of environmental signals have been called initial predictive information (such as photoperiod that initiates the reproductive process and sustains it) and local predictive information that acts as an inhibitor or accelerator (Fig. 5.3, see Wingfield et al. 1999). Photoperiodism in birds involves a physiological system that regulates not a change in responsiveness to day length, as it is often perceived, but a change in the quality of the response to changing day length (Nicholls et al. 1988; Wilson and Donham 1988). At one time of the year the neuroendocrine system responds to the appropriate long day length with physiological cascade that includes a dramatic increase in gonadotropin secretion, gonadal growth, and a range of hormone-dependent processes, including behavior. This state of responsiveness to long day lengths in birds is known as photosensitivity. However, temperate-zone birds develop a change in response to these stimulatory day lengths over time, so that at other times of the year the same day length maintains a state of reproductive inactivity (for reviews see Nicholls et al. 1988; Wilson and Donham 1988; Wingfield and Farner 1993; Dawson et al. 2001). In birds, this state is known as photorefractoriness following which GnRH cell bodies in the brain shrink and fibers emanating from these towards the median eminence show a marked decrease in immunocytochemical staining (Foster et al. 1987; Goldsmith et al. 1989; Boulakoud and Goldsmith 1991). This has also been demonstrated in the House sparrow (Hahn and Ball 1995). Thus, pituitary release of the gonadotropins luteinising hormone (LH) and follicle-stimulating hormone (FSH) is reduced to a minimum (Dawson and Goldsmith 1982, 1983) and the gonads undergo marked regression (see Fig. 5.10). Parry and Goldsmith (1993) have shown that increased synaptic input to the GnRH neurons is coincident with the long-term photorefractory state. Other physiological effects associated with photorefractoriness are a peak in plasma prolactin levels and a post-nuptial moult (Goldsmith and Nicholls 1984a; b). Despite the lack of gonadal activity and reproductive behavior during photorefractoriness, long day lengths maintain this inactivity. Without a long day stimulus, photorefractory birds develop what is termed photosensitivity, so that they are once again able to grow their gonads in response to a long day stimulus. Seen this way, it seems that the terms “photosensitive”, “photostimulated” and “photorefractory” are somewhat misleading, given that the reproductive system of photoperiodic birds is sensitive to, and exhibits a response to, the ambient photoperiod no matter what it is; it is just the nature of the response that changes from one time of year to another. It might be envisaged that changing photoperiod determines when birds can feed and how much they can take it. A decrease in food intake could be

$ Reproductive Biology and Phylogeny of Birds

Fig. 5.10 A. Immunocytochemically-localized gonadotropin-releasing hormone (GnRH) neurons in the hypothalamus of Rufous-collared sparrows, Zonotrichia capensis, from non-breeding (left panel) and breeding (right panel) individuals. Note the increased number of neuronal processes and larger, more intensely immunoreactive neurons in the breeding individual as compared to the nonbreeding individual. This reflects increased GnRH synthesis, transport and release. B. Testes from non-breeding and breeding European starlings, Sturnus vulgaris. Note the large change in testis volume (this mirrors the GnRH changes in the brain) and the change in color (reason unknown but possibly a result of circulating lymphocytes invading the regressing testis and depositing melanin). Original.

responsible for such a dramatic change in reproductive state from one time of year to another, but this is not the case. For example, in the European starling (Sturnus vulgaris), photorefractoriness ensues as day length is still increasing (Dawson and Goldsmith 1982) and therefore the opportunity to further increase food intake still exists as photorefractoriness occurs. Also, experimental manipulation with ad libitum food supply contradicts such this theory. An example of this is a study carried out by Dawson et al. (1985). They demonstrated that if non-photorefractory (i.e. photosensitive) starlings are exposed to only seven “long” days, then this is often sufficient to provide the photoperiodic drive for photorefractoriness to ensue some weeks later. It has also been shown that the rate of onset of photorefractoriness is proportional to the length of the photoperiod (Hamner 1971; Dawson and Goldsmith 1983). A long day is that in which the duration of the light period is over and above a “critical day length”, that being the length of day below which photorefractoriness cannot be induced. The experiment by Dawson et al. (1985) also indicates that even though photorefractoriness does not become apparent for several weeks after exposure to long days, it is initiated rapidly and the reproductive system continues to proceed towards a photorefractory state regardless of subsequent photoperiod once initiation is complete. More recently, Dawson (2001) provided evidence that a single long day can initiate

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progression towards photorefractoriness. In the same way, a single long day causes activation of the reproductive system in quail (Nicholls et al. 1983; Meddle and Follett 1995), and song sparrows (Wingfield 1993). Photorefractoriness is gradually dissipated by short days (i.e. day length below the critical photoperiod), and “photosensitivity” is acquired. That is, the bird’s gonads grow in response to long days again. In the wild, photosensitivity is acquired in the autumn, when day length falls below approximately 11.5 hr—resulting in some species what has been termed “autumnal sexuality” (Murton and Westwood 1977; Lincoln et al. 1980; Dawson 1983; Wingfield et al. 1987). Birds that breed within the tropics (where 60% of birds live) have traditionally been considered to be non-photoperiodic, since the annual change in photoperiod is slight (Dittami and Gwinner 1985). Nevertheless, tropical birds have been shown to be photoperiodic, e.g. African stonechats (Saxicola torquata) (Gwinner and Scheuerlein 1999) and Zebra finches (Bentley et al. 2000), although the experimental amplitude in photoperiod used in these studies exceeded that of the tropics. However, two other studies suggest that Spotted antbirds (Hylophylax naevioides) can respond to tropical changes in photoperiod (Hau et al. 1998; Beebe et al. 2005). The strategy adopted by tropical birds appears to be to remain in a physiological state of “readiness to breed” for a large proportion of the year, and then to use non-photoperiodic cues, such as rainfall or food abundance, to determine the exact time of breeding. In zebra finches, for example, breeding can occur at any time of the year, but breeding bouts are more intense during periods of rainfall (Zann et al. 1995). Stonechats may respond to low light intensity as a predictive cue for rainfall (Gwinner and Scheuerlein 1998). There is evidence that the HPG axis of such birds remains for much of the year in a state somewhere between that characteristic of photosensitivity and photostimulation in non-tropical seasonally breeding birds, and that full functionality is triggered by the relevant proximate cues (Perfito et al. 2006). Photorefractoriness does not seem to occur in spotted antbirds (Beebe et al. 2005). Gonadal regression and molt frequently follow breeding, but the extent to which the GnRH system is “switched off” during this period is mostly unknown. One study on Ecuadorean rufous-collared sparrows (Zonotrichia capensis) which experience only 3 min photoperiod fluctuation over the year suggests that there are changes in the GnRH system of a magnitude similar to that seen in obligately photoperiodic birds (Moore et al., 2006). Non-tropical northern hemisphere opportunistic breeders, such as crossbills can breed at most times of the year (mainly between January and August). Crossbills (Loxia curvirostra) feed on conifer seeds, the availability of which appears to synchronize reproductive effort. However, these species do have a fixed non-breeding period during fall, when the HPG axis appears to be photorefractory and when molt occurs (Hahn et al. 1995b; Hahn 1998). Even if opportunistic breeders are photoperiodic to a degree (Bentley et al. 2000), the relative importance of non-photoperiodic cues in “fine-tuning” the

& Reproductive Biology and Phylogeny of Birds timing of breeding varies and appears to be greater than that in obligately photoperiodic species (Wingfield 1980).

5.6.2

Absolute and Relative Photorefractoriness

In birds that exhibit absolute photorefractoriness, e.g. European starling, (Nicholls et al. 1988), and European rook (Corvus frugilegus) (Lincoln et al. 1980), breeding attempts are halted at a specific time of year and at a consistent time after initial photoperiodic stimulation. At this point, the gonads spontaneously regress, moult is initiated and hormonal changes occur concurrently. There is a rise in plasma prolactin levels coincident with the onset of moult (Dawson and Goldsmith 1982, 1983); plasma LH and FSH levels decrease to a minimum as photorefractoriness occurs (Goldsmith and Nicholls 1984a). Such changes occur even though day lengths are longer than those which were originally stimulatory and often still increasing. In other species where absolute photorefractoriness occurs—e.g. American crowned sparrows (Zonotrichia sp.), Mallard (Anas platyrhynchos), Canary (Serinus canarius)—changes associated with photorefractoriness happen when day length is decreasing but is still longer than those at the time of reproductive stimulation. Such a condition is termed absolute photorefractoriness because no increase in day length, even up to 24 hr light per day, will cause a change in the reproductive state once a photorefractory condition has been induced. A condition similar to absolute photorefractoriness is relative photorefractoriness. The main difference is that once relative photorefractoriness has been induced and the gonads have regressed, a subsequent substantial increase in day length will once more initiate reproductive maturation— without the need for a “sensitization”, or photosensitive, stage (Robinson and Follett 1982). For example, if Japanese quail experience day lengths of over 11.5 hr, rapid gonadal development occurs. After about 3 months, and when (in the wild) day length decreases below 14.5 hr, complete gonadal regression occurs—in a similar manner to absolute photorefractoriness (Nicholls et al. 1988). However, if the day length is subsequently artificially increased further, a full return to reproductive maturity occurs. Indeed, if quail are maintained on any constant long day length, no form of photorefractoriness will be elicited unless they experience a decrease in day length, for example, from 23 hr to 16 hr (Nicholls et al. 1988). This suggests a shift in the critical day length in birds which are relatively photorefractory—a shift which appears to depend on the photoperiodic history of such birds (Robinson and Follett 1982). There does not seem to be any change in critical day length in birds that exhibit absolute photorefractoriness, regardless of their photoperiodic history (Dawson 1987; Follett and Nicholls 1984, 1985) The fact that there is such an apparent difference between bird species that exhibit absolute photorefractoriness and those that exhibit relative photorefractoriness, and that breeding seasons vary so widely among bird species to suit their ecological needs has led to two schools of thought as to

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how mechanisms controlling the termination of the breeding season have evolved. The first school is that which argues for the separate evolution of similar mechanisms in different bird groups (Farner 1964; Farner et al. 1977). The idea behind such views is that although circadian periodicities became established very early in evolution, the photoperiodic systems that have been imposed upon them evolved much later and from multiple origins. The alternative view is that proffered by Follett and Nicholls (1984, 1985), who suggest that photorefractoriness has a single origin, but simple modifications have made the phenomenon appear to be very different in separate bird species. It is now generally accepted that relative photorefractoriness in quail is closely related to absolute photorefractoriness in starlings, and that both types of photorefractoriness involve long day-induced alterations to the photoperiodic response; which will not occur without a functional thyroid gland (Follett and Nicholls 1984, 1985). This explanation obviously dismisses the idea of a change, or “shift”, in the critical day length. If critical day length is shifted to a higher threshold during the onset of relative photorefractoriness (i.e. more hours of light are required per day for gonadal maturation), then gonadal regression is not long day-induced—as, by definition, critical day length is the boundary between “long” and “short” days, with long and short days having completely different photoperiodic effects. Similarities also exist between photorefractoriness in juvenile and adult Red-legged partridge (Alectoris graeca chukar), just as there are between juvenile and adult starlings (Dawson et al. 1987; Williams et al. 1987; Creighton 1988). Homology has also been suggested for the neural mechanisms controlling seasonal breeding in Welsh mountain ewes and starlings (Nicholls et al.. 1988b). In addition, if birds that exhibit absolute photorefractoriness are placed under short days whilst in a photorefractory state, photosensitivity is regained over a period of weeks. Before photosensitivity is fully regained, however, they are found to be in a state of relative photorefractoriness (Hamner 1968; Nicholls et al. 1988a). This demonstrates that the regaining of photosensitivity is a gradual process, and it can be shown that the rate of recovery is proportional to the “shortness” of short days, and to the length of time held on short days (Vaugien 1955; Farner and Follett 1966; Steel et al. 1975; Turek 1975; Nicholls and Storey 1977; Gwinner et al. 1988; Dawson 1991, Boulakoud and Goldsmith 1994, 1995).

5.6.3 Mechanisms Involved in the Detection and Transduction of Day Length Unlike other vertebrates, seasonally breeding mammals generally use a combination of their eyes (retina) and pineal gland for photoreception and transduction of the light signal (see Foster et al. 1989 for review). For example, if sexually mature hamsters are blinded whilst held under long days, then their gonads regress—even though they are still receiving the long-day light stimulus (Reiter 1978). This indicates that ocular photoreceptors are responsible for light detection and transduction. Although it has been unclear

 Reproductive Biology and Phylogeny of Birds as to the nature of the photoreceptors that fulfill this function (Foster et al. 1991), it appears that melanopsin is most likely involved (Hattar et al. 2003). Non-mammalian vertebrates, on the other hand, transduce seasonal changes in photoperiod via photoreceptors that are extraretinal and extrapineal (see Groos 1982 for review). The first evidence for avian extraretinal photoreceptors was shown by Benoit (1935a,b), who demonstrated that simultaneous photostimulation of blinded and sighted ducks resulted in equal testicular growth rates. Furthermore, the testicular response could be abolished by covering the duck’s head with a black cap. Since then, further experiments involving ducks (Benoit and Ott 1944), house sparrows (Menaker and Keatts 1968) and American tree sparrows, Spizella arborea (Wilson 1991) have demonstrated convincingly that an extraretinal photoreceptor participates in the reproductive responses of birds, and that there is little or no retinal involvement in reproductive responses (Underwood and Menaker 1970; see also Foster and Follett 1985). Thus in birds, the detection of light for reproductive purposes occurs via an extraretinal, hypothalamic pathway (Menaker 1971). In an elegant experiment, Yokoyama et al. (1977) used fiber optics to illuminate discrete areas of the hypothalamus and found that the infundibular nucleus appeared to contain most photoreceptors in Zonotrichia l. gambelii. Despite numerous studies, it is still unclear precisely as to where the photoreceptors lie in the avian brain. There is some evidence that extraretinal photoreceptors may reside in the infundibular region of the hypothalamus (Oliver and Baylé 1982) and in the parolfactory lobe of quail (Sicard et al. 1983). Additionally, Foster et al. (1985) demonstrated the involvement of a rhodopsin-like photopigment, which has a maximum spectral photosensitivity at 492 nm (Foster and Follett 1985). More recent research involving immunostaining of opsin (a protein involved in signal transduction by light activation) has added weight to the idea that the deep brain photoreceptors are located in the hypothalamus (in this case, the tuberal hypothalamus of the Ring dove (Streptopelia risoria), duck and quail—Silver et al. 1988; Saldanha et al. 2001). Thus, it appears that in birds, light must pass through the skull and into the hypothalamus for transduction of its signal to occur. For a “long day” signal to result in photostimulation, day length must be measured with a high degree of accuracy. Birds, unlike mammals, do not seem to require the melatonin as a time signal (from the pineal gland) to control reproductive changes in response to day length (but see later section on gonadotropin-inhibitory hormone, or GnIH). The physiological mechanism(s) underlying measurement of day length are still unknown, but there are two main models for which there are some supporting experimental data. The first is the “hour-glass” model, in which it is hypothesized that an “hour-glass”like timer is set in motion by the onset of dusk or dawn, and a photochemical accumulates during either the light or the dark phase. If a sufficient amount of the photochemical accumulates, then a photoperiodic response is initiated. The fundamental property of the hour-glass model is that it requires constant

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resetting by the light/dark cycle. Thus, it would not operate under conditions of constant light or constant dark. There is evidence that such a system operates in some insects (for examples see Beck 1968; Lees 1973; Vaz Nunes and Veerman 1984, 1986; Veerman et al. 1988). The second theory, which has been shown to hold true for birds, was first proposed by Bünning in 1936 (see also Pittendrigh and Minis 1964, 1971; Follett 1973; Elliott 1976). It assumes that there is a circadian rhythm of responsiveness to light, i.e. for the first part of the cycle (subjective day) the organism is insensitive to light; whereas during the second part (subjective night) it becomes “photoinducible”. Should light fall during the subjective night, then a “long-day” photoperiodic response is initiated. Evidence that this “external coincidence model” applies to avian photoperiodicity was first supplied by night-interruption experiments on the House finch (Carpodacus mexicanus) (Hamner 1963, 1964), which has been reinforced by experiments on other bird species (for examples see Follett and Sharp 1969; Turek 1974; Follett et al. 1992). Another model of how photoperiodic responses are initiated by day length is termed the “internal coincidence model” (Pittendrigh and Minis 1964). This model assumes that photoperiod entrains two circadian oscillators, one entrained by dusk and the other by dawn, with photoperiodic time measurement a function of the phase relationship between the two clocks. Thus, when the phase of the two oscillators coincides, photoinduction occurs. Although there is little experimental evidence to help distinguish between the latter two hypotheses, Follett et al. (1974) in an elegant experiment looking at a surge in plasma LH induced by the coincidence of light with the photoinducible phase in Zonotrichia l. gambelii, showed that the responsiveness to light had a strong circadian component with peak LH responses between 12 and 18 hours after the subjunctive dawn. This is consistent with the external coincidence model. Other experiments on Zonotrichia sp suggested that circadian rhythms in corticosterone and prolactin represented internal coincidence because injections of these hormones at different times of day had different effects on breeding, fattening and migration (Meier et al. 1965; Meier 1972; Meier and McGregor 1972). Evidence against the external coincidence model comes from Bentley et al. (1998), in an experiment where light intensity, rather than length of photoperiod, was manipulated. Few investigations have incorporated manipulation of light intensity rather than day length to affect reproductive responses since Bissonnette’s study in 1931. It is difficult to draw definitive conclusions from most early experiments on this subject, as mixtures of natural and artificial light were used in the same days of treatment (Bissonnette 1931); others omitted to control for different wavelengths of light from sources of differing intensities (e.g. Bissonette and Wadlund 1933) [the importance of wavelength in the avian photoperiodic response has been established for some time (Ringoen 1942; Benoit and Ott 1944)]. There are even reports of both low and high light intensities causing convulsions and death in the African weaver finch (Rollo and Domm 1943)! So far, the most



Reproductive Biology and Phylogeny of Birds

comprehensive experiment of this type to have been carried out was by Bartholomew (1949), in which house sparrows were subjected to different light intensities under the same photoperiod. His study concluded that light intensity can indeed modify the reproductive response of sparrows to day length, but that there is a minimum intensity below which no photoperiodic response can be evoked. In addition, there is an upper intensity threshold above which there is no increase in rate of response, and intensity cannot be substituted for day length per se. Work carried out on mammals indicates that there are also light intensity effects on the mammalian endocrine system, such as in ferrets (Marshall and Bowden 1934) and pigs (Griffith and Minton 1992). In the experiment by Bentley et al. (1998), photosensitive starlings transferred from short days to long days of different light intensities underwent graded reproductive responses according to the light intensity they experienced. Testes size in the group in the lowest intensity (3 lux) increased faster than that in controls on short days of normal intensity, but they did not become photorefractory. Testes size increased in the groups on 13, 45, and 108 lux and subsequently became photorefractory. However, the 13- and 45-lux groups required more time to become photorefractory than did the 108-lux group. The responses observed were similar to those seen in starlings exposed to different photoperiods (e.g. 11 hr light:13 hr dark [11L:13D], 13L:11D, 16L:8D, 18L:6D), even though all were on the same 18L:6D photoperiod. Initially, the results appear to challenge the external coincidence model for photoperiodic time measurement, but consideration of the phase response curve of the circadian rhythm of photoinducibility in starlings and the way in which it might be affected by low light intensities refute this challenge and reinforce the external coincidence model (see Bentley et al. 1998 for discussion). For further details on recent advances in our understanding of the molecular and cellular intricacies of the avian circadian system, see Dawson et al. 2001; Brandstatter 2003).

5.6.4 Physiological Mechanisms that have been Suggested to be Involved in the Control of Photorefractoriness A seasonal change in hypothalamic sensitivity to gonadal steroid feedback (Cusick and Wilson 1972; Sharp and Moss 1977; Stokkan and Sharp 1980a,b) considers the idea that as gonadal steroid feedback increases under long days, the hypothalamus becomes increasingly sensitive to this feedback and “shuts down” completely, resulting in photorefractoriness. This was shown not to be the case because gonadectomised Zonotrichia l. gambelii and European starlings became photorefractory in the same way, and within the same timecourse as intact birds (Wingfield et al. 1980; Dawson and Goldsmith 1984). The conclusion was drawn that gonadal steroid feedback is unimportant in the induction and maintenance of photorefractoriness. Also, long days override the inhibitory effect of gonadal steroids on hypothalamic GnRH. There is no decrease in hypothalamic sensitivity to the inhibitory effects of steroids (inhibition must increase as steroid output from the gonads increases

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with long days) but the “photoperiodic drive” is increased to a greater level than the inhibition by the steroids, producing a net positive effect (Matt 1983; Nicholls et al. 1984). Furthermore, Wilson (1985) has demonstrated that in American tree sparrows, hypothalamic sensitivity to inhibition by gonadal steroids decreases, rather than increases, following exposure to long days. It is possible that an inhibitory hormone terminates reproduction and prolactin was thought to be a strong candidate for this (Dawson and Goldsmith 1982; Ebling et al. 1982; Goldsmith 1983; Dawson and Goldsmith 1983), as there is always a peak in the level of plasma prolactin coinciding with the physical manifestations of photorefractoriness. This theory is unlikely, however, as the administration of exogenous prolactin does not on its own cause the onset of photorefractoriness (Goldsmith 1985), although this experiment was carried out using heterologous prolactin. Further evidence against the idea that prolactin is involved in the control of photorefractoriness is that if photosensitive starlings are thyroidectomized and transferred from short to long days, photorefractoriness does not occur, and there is no rise in plasma prolactin levels (Dawson and Goldsmith 1984). This indicates that prolactin probably does not have a causal role in the instigation of photorefractoriness, as the pituitary is not damaged by thyroidectomy and hence the prolactin-secreting cells are intact (Goldsmith and Nicholls 1984b; Dawson et al. 1985). However, this approach does not take into account the fact that thyroid hormones may be required as “permissive” agents for the release of prolactin as a result of photostimulation, thus resulting in photorefractoriness. It can be shown that prolactin is not responsible in itself for gonadal regression. If photosensitive starlings are kept under a photoperiod of 11 hours of light and 13 hours of darkness per day (11L:13D), slow but complete reproductive maturity ensues (Hamner 1971; Goldsmith and Nicholls 1984c). With 11L, photorefractoriness does not occur because this is below the critical day length for this species. When the birds are subsequently transferred back to short days (6-8L), the gonads regress in size, but there is no increase in plasma prolactin levels (Goldsmith and Nicholls 1984c). If such 11L birds are put in to long days (18L) instead of being returned to short days, photorefractoriness occurs, along with the associated rise in plasma prolactin. Thus, prolactin is not responsible for gonadal regression in short days, but is associated somehow (perhaps only temporally) with regression during photorefractoriness. The timing of high levels of circulating prolactin is also closely linked to plumage moult, and, where it occurs, premigratory fattening and migration (Meier and MacGregor 1972; Meier 1972; Dawson and Goldsmith 1983). A clear-cut experiment demonstrating that prolactin is not a causative agent of photorefractoriness was performed by Dawson and Sharp in 1998. European starlings were actively immunized against vasoactive intestinal polypeptide (VIP), the prolactin releasing hormone in birds, or against prolactin, during a photo-induced breeding cycle. VIP-immunized birds became photorefractory but the rate of gonadal regression was markedly

" Reproductive Biology and Phylogeny of Birds slowed, and the photo-induced increase in prolactin was completely suppressed in 50% of these birds. Molt was prevented in those birds in which prolactin release was completely suppressed. In those VIP-immunized birds in which the photo-induced increase in prolactin was inhibited but not completely suppressed, gonadal regression was delayed, but molt occurred as normal. The same was true for prolactin-immunized birds. There were no significant differences in concentrations of plasma thyroxine between treatment and control groups, indicating that the effects of immunization on gonadal regression were not mediated by the induction of hypothyroidism. Thus, in European starlings the associated increase in prolactin accelerates gonadal regression during the onset of photorefractoriness but does not itself cause photorefractoriness. In addition, the increase in prolactin associated with photorefractoriness is required for the induction of the postnuptial molt (Dawson and Sharp 1998). Following on from this, Dawson et al. (2002) demonstrated that VIP as measured by radioimmunoassay did not change over a photoperiod-induced cycle in European starlings. Therefore, photorefractoriness in terms of prolactin secretion is not similarly related to a decrease in basal hypothalamic VIP. An exhaustion of some component of the hypothalamo-hypophysial axis (Nicholls and Storey 1976) is a third hypothesis. The idea that some component of the hypothalamo-hypophysial axis has, once stimulated, a limited period of activity and then requires a recovery period to “build up reserves” does fit in many ways with the annual reproductive cycle of birds that exhibit photorefractoriness. One suggestion was, broadly, that the hypothalamic GnRH-synthetic cells have a constant rate of GnRH synthesis and that it is only the secretion rate that fluctuates during the annual change in photoperiod (Nicholls and Storey 1976). The proposal is that secretion is only permitted under long days (but some “leaks” out when the neurosecretory GnRH cells are “full”—permitting photosensitivity), and that under long days, secretion occurs at a greater rate than synthesis. Thus, the rapid secretion of GnRH allows photorefractoriness to occur. A time lag is subsequently needed (under short days, so that there is no secretion) for GnRH reserves to “ build up” prior to exposure to long days in the spring. They can then be released in sufficient quantities to elicit a reproductive response. The above hypothesis can be seen to tally with the responses shown by starlings to long days when in a relatively photorefractory state - i.e. when photorefractory birds are put into short days and after total photorefractoriness has been lost, but before full photosensitivity has been acquired. What is seen is that photorefractoriness is induced more rapidly than in fully photosensitive birds, so it could be construed that GnRH levels have not had time to build up to their maximum. Also, the fact that the rate of onset of photorefractoriness is proportional to the duration of the prevailing photoperiod (Hamner 1971; Dawson and Goldsmith 1983) could be assumed to agree with this hypothesis. But since it has been shown that changes in

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hypothalamic content of GnRH occur after gonadal regression has been initiated, it seems improbable that the exhaustion hypothesis is tenable (Nicholls et al. 1988). In addition, there is the fact that thyroidectomy permits sexual maturity to be maintained, perhaps indefinitely on long days (Wieselthier and van Tienhoven 1972; Goldsmith and Nicholls 1984b; Dawson et al. 1985). This appears to demonstrate that without a higher neural “switching” process occurring, GnRH continues to be synthesized and secreted at a high rate during appropriate photoperiodic stimulation. Other experiments involving the use of exogenous thyroxine or a thyroid-dependent neurotrophin to mimic the effect of long days in reducing hypothalamic GnRH content (Boulakoud and Goldsmith 1991) also dispute the “exhaustion” hypothesis (Bentley et al. 1997a,b). Other experiments in which GnRH-I was injected into photorefractory male Zonotrichia l. gambelii resulted in a dramatic, but short-lived, increase in LH also argues against the exhaustion hypothesis (Wingfield et al. 1979). Similar results were obtained following injection of the excitatory amino acid agonist NMDA injected into photorefractory Zonotrichia l. gambelii (Meddle et al. 1999). The state of photorefractoriness can also be “broken”, or dissipated, by the removal of circulating thyroid hormones—i.e. thyroidectomy. This technique has shown that both the initiation and maintenance of photorefractoriness are dependent upon the presence of thyroxine (Wieselthier and van Tienhoven 1972; Goldsmith and Nicholls 1984b). Termination of the breeding condition by thyroidectomy has also been observed in other vertebrates—e.g. tree sparrows (Wilson and Reinert 1993; 1995a,b), sheep (Moenter et al. 1991; Dahl et al. 1994; Parkinson and Follett 1994; Parkinson et al. 1995) and red deer (Shi and Barrell 1992). Although the presence of thyroxine is required, and plasma thyroxine levels rise markedly upon a starling’s transfer from short to long days (Dawson 1984), and also in quail (Sharp and Klandorf 1981), it is likely that thyroid hormones are not the cause of photorefractoriness. It appears that thyroxine is a permissive factor allowing the mechanisms that cause the reproductive system to “switch off” under long days to function, and may be involved in the perception of day length (Dawson 1989a,b; Bentley et al. 1997a,b). Similar suggestions have been made for the termination of breeding in sheep (Dahl et al. 1995). It has not been clear as to whether the long dayinduced rise in circulating thyroxine is a necessary precursor of the photorefractory response, or if this rise is associated with other long day responses, such as increased metabolic rate. Importantly, thyroidectomy does not affect the daily or circadian pattern of circulating melatonin concentrations (Dawson and King 1994). At another level, it has been shown that in photostimulated Zonotrichia l. gambelii, glial cell process do not interpose between axon terminals and capillaries of the portal system in the median eminence, but they do in photorefractory birds (see Fig. 5.11, from Bern et al. 1966; Farner et al. 1967; Mikami et al. 1978) The same appears to be true for Japanese quail (Yamamura et al. 2004).

$ Reproductive Biology and Phylogeny of Birds

5.7 A NEW LEVEL OF CONTROL OF AVIAN REPRODUCTION: NOVEL PEPTIDES 5.7.1

A Third Type of GnRH

A recent study (Bentley et al. 2004) presented evidence for the presence of an immunoreactive third GnRH in songbirds that is clearly hypophysiotropic and has gonadotropin-releasing capabilities. This third GnRH, immunoreactive (ir)-lamprey GnRH-III, possibly has multiple functions, as suggested by its widespread distribution. In addition, ir-lamprey GnRH-III is present in abundance in telencephalic areas, including the hippocampal formation and the song control system. In no vertebrate has a GnRH been localized in these “higher” control regions before, although fragments of mammalian GnRH have been detected in primate forebrain, and their functions are unknown (Terasawa et al. 2001). In fact, we are aware of only two studies that have investigated the GnRH system concurrently with the oscine song control system (MacDougall-Shackleton et al. 2001; Marsh et al. 2002), most likely because the distribution of ir-chicken GnRH-I and -II are so distant from telencephalic areas (Juss et al. 1992; Millam et al. 1993). The finding that a third GnRH is likely involved in regulation of reproductive function in songbird species has implications for the way in which we envisage the avian brain processes environmental cues and transduces them into endocrine signals. We will understand this putative peptide more when it is isolated and sequenced from the songbird brain.

5.7.2 Gonadotropin-Inhibitory Hormone (GnIH) Since the molluscan cardioexcitatory neuropeptide Phe-Met-Arg-Phe-NH2 (FMRFamide) was found in the ganglia of the venus clam (Price and Greenberg 1977), neuropeptides that possess the RFamide motif at their Ctermini (i.e., RFamide peptides) have been characterized in various invertebrates (see Tsutsui and Ukena 2006 for review). Subsequently, many immunohistochemical studies that used the antiserum against FMRFamide suggested that vertebrate nervous systems possess some unknown neuropeptides similar to FMRFamide (Raffa 1988; Rastogi et al. 2001). Immunohistochemical findings indicated that some of the FMRFamide-like immunoreactive neurons projected to the hypothalamic region close to the pituitary gland, and thus were expected to play an important role in the regulation of pituitary function (see Tsutsui and Ukena 2006 for review). Tsutsui et al. (2000) therefore looked for a novel RFamide peptide in the avian brain. Amino acid sequence analysis of a novel isolated substance revealed the following sequence: Ser(62)-Ile(252)-Lys(233)-Pro(226)-Ser(38)-Ala(194)Tyr(173)-Leu(148)-Pro(104)-Leu(108)-Arg(45)-Phe(52) with the detected amount (pmol) of each amino acid indicated in parentheses. The isolated native peptide in the quail brian was confirmed as a 12 amino acid sequence (SIKPSAYLPLRFamide) with RFamide at the C-terminus (Tsutsui et al. 2000). This neuropeptide had not been previously reported in vertebrates, although

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Fig. 5.11 A. and B. Posterior median eminence (PME) with a primary capillary (PCp) of a photostimulated male white-crowned sparrow, Zonotrichia leucophrys gambelii. The capillary is separated from the median eminence by two layers of basement membrane (Bm) and is covered by perivascular cells (PVC) and/or pericytes (Pc). Note that glial cells (Gc) do not interpose between axons with neurosecretory material and the basement membranes and endothelial cells (En). From Mikami et al. (1970) Zeitschrift für Zellforschung 106: 155-174, Springerverlag, Berlin, Fig. 3; see also Bern et al. 1966. Zeitschrift für Zellforschung 69: 198227, Springer-verlag, Berlin, Fig. 13. C. Electron micrograph of the ventral region of the posterior median eminence of a photorefractory male white-crowned sparrow. Axons contain small vesicles (Ve) and a few dense granules (Gr). A layer of glial processes (Gl) is interposed between the axons and the basement membrane region (Bm) of the portal capillary. An erythrocyte is at lower left. Mi = mitochondria. From Farner, D. S., Wilson, F. E. and Oksche, A. (1967). Pp. 529-582. In L. Martini and W. F. Ganong (eds.), Neuroendocrinology, vol. 2. Academic Press, New York, Fig. 10.

the C-terminal LPLRFamide was identical to chicken pentapeptide LPLRFamide peptide (Dockray et al. 1983). Subsequently, the isolated novel peptide was shown to be located in the quail hypothalamo-hypophysial system and to decrease gonadotropin release from cultured anterior pituitary in a dose-dependent manner (Tsutsui et al. 2000). This novel RFamide peptide was therefore named as gonadotropin-inhibitory hormone (GnIH) (Tsutsui et al. 2000). Using immunohistochemistry, clusters of distinct GnIH-immunoreactive (-ir) neurons were found in the paraventricular nucleus (PVN) in the hypothalamus. In addition to the PVN, some scattered small cells were

& Reproductive Biology and Phylogeny of Birds immunoreactive in the septal area (Tsutsui et al. 2000; Ukena et al. 2003; Ubuka et al. 2003). In contrast to the highly-localized clusters of cell bodies, GnIH-ir nerve fibers were widely distributed in the diencephalic and mesencephalic regions. Dense networks of immunoreactive fibers were found in the ventral paleostriatum, septal area, preoptic area, hypothalamus, and optic tectum. The most prominent fibers were seen in the median eminence (ME) of the hypothalamus, and in the dorsal motor nucleus of the vagus in the medulla oblongata. We further investigated GnIH localization in the brain of seasonallybreeding songbird species (Bentley et al. 2003; Osugi et al. 2004). Dense populations of GnIH-ir neurons were also found in the PVN of Song sparrows, House sparrows, White-winged crossbills (Loxia leucoptera), Pine siskins (Carduelis pinus), Redpolls (Carduelis flammea), Rufous-collared sparrows (Zonotrichia capensis), Rufous-winged sparrow (Aimophila carpalis) and Gambel’s white-crowned sparrows. The PVN was the only location where immunoreactive neurons were located, regardless of sex or species (data only published for four species in this list: Bentley et al. 2003; Osugi et al. 2004; Deviche et al. 2006). Thus the presence of GnIH in the PVN appears to be a conserved property among several families and at least two orders of birds (Galliformes and Passeriformes). In addition to the dense population of GnIHir neurons within the hypothalamus of all the avian species studied so far, there were extensive networks of branching beaded fibers emanating from those cells, presumably transporting GnIH. Some of the fibers extended to terminals in the ME, consistent with a role for GnIH in pituitary gonadotropin regulation. In house sparrows, song sparrows and Gambel’s white-crowned sparrows, other fibers extended through the brain caudally at least as far as the brainstem and possibly into the spinal cord, consistent with multiple regulatory roles for GnIH (see Fig. 5.12). Tsutsui’s group further examined the precursor polypeptide for GnIH and localization of its transcript, which would provide key information as to the regulation of the mature GnIH peptide, along with confirmation of brain area(s) that synthesize this novel peptide. A cDNA that encoded the GnIH precursor polypeptide was identified in the quail brain by a combination of 3' and 5' rapid amplification of cDNA ends (3'/5' RACE) (Satake et al. 2001). The deduced GnIH precursor consisted of 173 amino acid residues that encoded one GnIH and two putative GnIH-related peptide (GnIH-RP-1 and GnIH -RP2) sequences that included -LPXRF (X = L or Q) at their C-termini. All of these peptide sequences were flanked by a glycine C-terminal amidation signal and a single basic amino acid on each end as an endoproteolytic site. In collaboration with Tsutsui’s group, we also cloned a cDNA that encoded GnIH in the brain of Gambel’s white-crowned sparrow (Osugi et al. 2004). The deduced sparrow GnIH precursor also consisted of 173 amino acid residues, encoding one sparrow GnIH and two sparrow GnIH-related peptides (sparrow GnIH-RP-1 and GnIH-RP-2) that included -LPXRFamide (X = L or Q) at their C-termini. Although the homology of sparrow and quail GnIH precursors was approximately 66%, the C-terminal structures of GnIH, GnIH-

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Fig. 5.12 A. Sagittal section from female house sparrow, Passer domesticus, brain with dense GnIH immunoreactivity in the PVN. B. Higher magnification of the PVN of male house sparrow brain in sagittal section showing individual GnIH-ir neurons and fibers emanating from the cell bodies. C. Sagittal section showing GnIH-ir fibers in the fascisculus longitudinalis medialis (FLM) of the brainstem of a female house sparrow. Arrows indicate GnIH-ir fibers. Cb= cerebellum. D. Higher magnifications of C. Scale bars, 100 mm. Original.

RP-1 and GnIH-RP-2 were all identical in two species (Satake et al. 2001; Osugi et al. 2004). Subsequently, a cDNA encoding GnIH and GnIH-RPs was also reported in the chicken from a gene database (see Tsutsui et al. 2005 for review). The chicken LPLRFamide peptide (Dockray et al. 1983) is considered to be a fragment of GnIH and GnIH-RP-1) (see Tsutsui et al. 2005 for review). In situ hybridization further revealed the cellular localization of GnIH mRNA solely in the PVN of quail and sparrow hypothalami (Ukena et al. 2003; Osugi et al. 2004). As already discussed, immunohistochemical analysis using the quail and sparrow also showed that quail and sparrow GnIH-ir cell bodies and terminals were localized in the PVN and ME, respectively. Thus only the PVN expresses GnIH and, in birds, the immunoreactive peptide found in fibers in multiple brain areas appears to originate from the PVN only (Ukena et al. 2003; Osugi et al. 2004).

5.7.3

Relative Distributions of GnIH and GnRH

The cell bodies for each population of neurons containing chicken GnRH-I (cGnRH-I), GnIH or cGnRH-II are in discrete locations. The preoptic area

 Reproductive Biology and Phylogeny of Birds (POA) contains cGnRH-I-ir neurons, the PVN contains GnIH-ir neurons and the cGnRH-II neurons are located in the midbrain (Millam et al. 1993). Thus we can confidently distinguish between two forms of GnRH based upon their location in the brain and their appearance. Closer inspection of each area indicates close proximity of GnIH fibers to the cGnRH-I neurons and fibers in the POA in songbirds. The PVN also contains cGnRH-I fibers which pass directly through and in close proximity to the population of GnIH neurons and fibers as they project to the ME. GnIH-ir fibers are also in close proximity to cGnRH-II neurons in the midbrain (Bentley et al. 2003). Further, confocal microscopy indicates that the cGnRH-I and GnIH peptides are located within the same 0.2 micron optical plane and possibly in contact with one another, although electron microscopy will be necessary to determine contact conclusively. Contact appears to occur between GnIH fibers and GnRH-I and -II neurons, and between GnIH and GnRH-I fibers in the ME (Bentley et al. 2003). Taken together, there is potential for GnIH to influence the GnRH system at the neuron and fiber terminal levels. Furthermore, when song sparrows were subjected to a simulated annual cycle of changing photoperiod, GnIH-ir neuron area was significantly greater at the onset of photorefractoriness (long days) when compared to photosensitive (short days) or photostimulated (long days) birds. Thus there is potential for dynamic interactions of GnIH and GnRH peptides in different reproductive states and neuroanatomical locations. It is clear that further study is needed to elucidate the seasonal dynamics of GnIH synthesis and release and its relation to seasonal changes in GnRH in songbirds.

5.7.4 Actions of GnIH on Gonadotropin Synthesis and Release in Galliformes Using cultured quail anterior pituitaries, GnIH significantly inhibited LH release, after 100-min incubation (Tsutsui et al. 2000). The inhibitory effect on LH release was dose-dependent and its threshold concentration ranged between 10–9 and 10–8 M. There was no effect of GnIH on prolactin release. Addition of a physiological dose (10–7 M) of GnIH to short-term (120 min) cultures of diced pituitary glands from adult cockerels suppressed common a and FSH b subunit mRNAs, with no effect on LH b subunit mRNA (Ciccone et al. 2004). The suppressive effect of GnIH on gonadotropin mRNA was associated with an inhibition of both LH and FSH release in the adult chicken (Ciccone et al. 2004). When administered intraperitoneally to quail in vivo via osmotic pumps, GnIH significantly reduced gonadotropin common a and LH b subunit mRNAs in a dose-dependent manner as well as reducing plasma LH (Ubuka et al. 2006). To elucidate the mode of action of GnIH, a novel G protein-coupled receptor for GnIH was identified in quail (Yin et al. 2005). The identified GnIH receptor was expressed in the pituitary and specifically bound to GnIH in a concentration-dependent manner (Yin et al. 2005). Southern blotting analysis

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of reverse-transcriptase-mediated PCR products revealed the expression of GnIH receptor mRNA in the pituitary and several brain regions including the hypothalamus in the quail (Yin et al., 2005). These results indicate that GnIH acts directly on the pituitary via GnIH receptor to inhibit gonadotropin synthesis and release. GnIH may also act on the hypothalamus to inhibit the activity of GnRH neurons. To clarify the functional significance of GnIH and its potential role as a key neuropeptide involved in avian reproduction, GnIH actions on gonadal development and maintenance were investigated in the male quail (Ubuka et al. 2006). Continuous administration of GnIH to mature quail via osmotic pumps decreased plasma testosterone concentrations (Ubuka et al. 2006). Interestingly, administration of GnIH to mature quail induced testicular apoptosis and decreased spermatogenic activity in the testis (Ubuka et al. 2006). GnIH administration to immature quail also suppressed normal testicular growth and rise in plasma testosterone concentrations (Ubuka et al. 2006). These results indicate that GnIH inhibits gonadal development and maintenance through the decrease in gonadotropin synthesis and release. GnIH may participate not only in neuroendocrine function but also in behavioral and autonomic mechanisms. Because GnIH neurons are localized in the PVN, with their fibers visible in multiple brain locations including the ME and brainstem, the action of GnIH on feeding behavior was further investigated in chicks (Tachibana et al. 2005). Intracerebroventricular (i.c.v.) injection of GnIH stimulated food intake in chicks. The chicken pentapeptide LPLRFamide, a degraded C-terminus of GnIH did not stimulate feeding thereby demonstrating the importance of the N-terminus of GnIH for the orexigenic effect. Anti-GnIH antiserum suppressed appetite induced by fasting. These data suggest that GnIH acts as an endogenous orexigenic factor in the brain of chicks. Food restriction induces not only an increase of appetite but also a reduction of the HPG axis activity (Richard-Yris et al. 1987). These changes also occurred after GnIH treatment, since GnIH stimulated feeding behaviour (Tachibana et al. 2005) and inhibited the HPG axis (Tsutsui et al. 2000; Osugi et al. 2004; Ciccone et al. 2004; Ubuka et al. 2006).

5.7.5 Seasonal Dynamics and Functional Significance of GnIH in Passerines Once we had identified the White-crowned sparrow GnIH precursor, we were able to synthesize putative White-crowned sparrow GnIH (“sparrow” GnIH) with the deduced amino acid sequence SIKPFSNLPLRFamide. We used this peptide for in vivo analysis on gonadotropin release in field-caught Whitecrowned sparrows during their breeding season in northern Alaska. Birds that were injected with GnIH had lower plasma LH at 2 min than the salineinjected group (Osugi et al. 2004). In contrast, saline-injected birds had high plasma LH (approx 5 ng/ml) 2 min post-injection. This difference was shortlived and had dissipated by 10 min.

Reproductive Biology and Phylogeny of Birds

As already mentioned, the distribution of GnIH in the avian brain suggests that it has not only hypophysiotropic actions but also unknown behavioral actions. GnIH fibers are present in the ME, and are in apparent contact with chicken GnRH (cGnRH) -I and -II neurons and fibers. In birds, cGnRH-I regulates pituitary gonadotropin release, whereas cGnRH-II is known to enhance copulation solicitation in estradiol-primed females exposed to male song (Maney et al. 1997). In a recent study (Bentley et al. 2006), we determined the effects of GnIH administered centrally to female white-crowned sparrows. A physiological intracerebroventricular (i.c.v.) dose of GnIH rapidly reduced circulating LH and inhibited copulation solicitation, without affecting locomotor activity. Using rhodaminated GnIH delivered i.c.v., putative GnIH binding sites were seen in the ME close to GnRH-I fiber terminals, and in the midbrain on or close to GnRH-II neurons. These data demonstrate direct effects of GnIH upon reproductive physiology and behavior, possibly via separate actions on two forms of GnRH. Taken together, our results indicate that, despite amino acid sequence differences, sparrow GnIH and quail GnIH (see Osugi et al. 2004) have similar inhibitory effects on the reproductive axis in wild sparrow species. Overall, GnIH appears to be a modulator of gonadotropin release in vivo as well as in vitro. Taking the immunohistochemical data into consideration as well, there is potential for GnIH to act on the reproductive axis at the level of the GnRH neurons, the ME, and the pituitary, and thus possibly over different timeframes. Furthermore, the wide distribution of GnIH terminal fields coupled with the fact that GnIH can influence reproductive behavior implies that there are as yet unknown actions of GnIH or GnIH-related peptides upon physiology and behavior.

5.7.6

Regulation of GnIH Expression in the Brain

Until now, a regulatory mechanism(s) governing GnIH expression has remained unclear. We have already mentioned that although many bird species are photoperiodic, a dogma has existed that birds do not use seasonal changes in melatonin secretion to time their reproductive effort, and a role for melatonin in birds has remained enigmatic (Wilson 1991; Juss et al. 1993). Despite the accepted dogma, there is strong evidence that melatonin is involved in regulation of several seasonal processes, including gonadal activity and gonadotropin secretion (Ohta et al. 1989; Bentley et al. 1999; Bentley and Ball 2000; Bentley 2001; Guyomarc’h et al. 2001; Rozenboim et al. 2002). In light of these reports and considering GnIH’s inhibitory effects on gonadotropin secretion (Tsutsui et al. 2000; Osugi et al. 2004), we investigated the action of melatonin on GnIH expression in the quail brain (Ubuka et al. 2005). Pinealectomy combined with orbital enucleation (Px+Ex) decreased the expression of GnIH precursor mRNA and the mature peptide GnIH in the diencephalon including the PVN and ME. Melatonin administration to Px+Ex birds caused a dose-dependent increase in expression of GnIH precursor mRNA and production of mature peptide. The expression of GnIH was photoperiodically controlled and increased under short day photoperiods

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(Ubuka et al. 2005), when the duration of melatonin secretion increases (Cockrem and Follett 1985; Kumar and Follett 1993). Finally, Mel1c, a melatonin receptor subtype was expressed in GnIH-ir neurons in the PVN (Ubuka et al. 2005). Thus melatonin appears to act directly on GnIH neurons via its receptor to induce GnIH expression.

5.8

CONCLUSIONS

In conclusion we hope we have demonstrated, at least in part, how complex are the mechanisms that control avian reproduction. Readers of this chapter should also bear in mind that the recent discoveries of new hormones in this field highlight that although a great deal is already known about neural integration of environmental information, there is a great deal still waiting to be discovered. This latter comment could easily be applied to other vertebrate classes. New tools at our disposal, such as those in molecular biology and proteomics will undoubtedly hasten our discovery of novel peptides and control mechanisms. However, the avian Class is so diverse, and their reproductive systems and strategies so varied that future avian biologists will undoubtedly be puzzling over some of the same questions that we have raised in this chapter. Comparative studies on different species in different vertebrate classes, such as those described in this chapter will enable us to gain a better understanding vertebrate reproductive biology as a whole. For example, studies on birds have uncovered several “firsts” in reproductive biology; the discoveries of GnRH-II and GnIH are two prime examples. Both of these neuropeptides appear to be relevant to reproduction in all vertebrates studied, including humans. Another “first” is the dramatic adult neurogenesis and neuroplasticity in the song control system of songbirds (Nottebohm 1981). Again, this paved the way for exciting insights into adult neuroplasticity in mammals, once believed not to exist.

5.9

ACKNOWLEDGMENTS

Preparation of this review was supported by grants IBN-0317141 and OPP9911333, from the National Science Foundation and by R01 MH65974-01 from the National Institutes of Health and by 15207007, 16086206 and 18107002 from the Ministry of Education, Science and Culture, Japan.

5.10

LITERATURE CITED

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Singh, K.B. and Dominic, C. J. 1975. Anterior and posterior groups of portal vessels in the avian pituitary, incidence in forty nine species. Archives d’Anatomie Microscopique et de Morphologie Experimentale (Paris) 64: 359-374. Solomon, S. E. 1983). Oviduct. Pp. 379-420. In B.M. Freeman (ed.), Phsiology and Biochemistry of the Domestic Fowl. Academic Press, London. Steel, E., Follett, B. K. and Hinde, R. A. 1975. The role of short days in the termination of photorefractoriness in female canaries (Serinus canarius). Journal of Endocrinology 64: 451-464. Stevenson, T.J. and Macdougall-Shackleton, S.A. 2005. Season- and age-related variation in neural cGnRH-I and cGnRH-II immunoreactivity in house sparrows (Passer domesticus). General and Comparative Endocrinology 143:33-39. Stockell-Hartree, A. and Cunningham, F. J. 1969. Purification of chicken pituitary follicle-stimulating hormone and luteinizing hormone. Journal of Endocrinology 43: 609-619. Stokkan, K.-A. and Sharp, P. J. 1980a. The roles of daylength and the testes in the regulation of plasma LH levels in photosensitive and photorefractory willow ptarmigan (Lagopus lagopus lagopus). General and Comparative Endocrinology 41: 520-526. Stokkan, K.-A. and Sharp, P. J. 1980b. The development of photorefractoriness in willow ptarmigan (Lagopus lagopus lagopus) after the suppression of photoinduced LH release with implants of testosterone. General and Comparative Endocrinology 41: 527-530. Tachibana, T., Sato, M., Takahashi, H., Ukena, K., Tsutsui, K. and Furuse, M. 2005. Gonadotropin-inhibiting hormone stimulates feeding behavior in chicks. Brain Research 1050: 94-100. Temple, J. L., Millar, R. P. and Rissman, E. F. 2003. An evolutionarily conserved form of gonadotropin-releasing hormone coordinates energy and reproductive behavior. Endocrinology 144: 13-19. Terasawa, E., Busser, B. W., Luchansky, L. L., Sherwood, N. M., Jennes, L., Millar, R. P., Glucksman, M. J. and Roberts, J. L. 2001. Presence of luteinizing hormonereleasing hormone fragments in the rhesus monkey forebrain. Journal of Comparative Neurology 439: 491-504. Teruyama, R. and Beck, M. M. 2000. Changes in immunoreactivity to anti-cGnRH-I and -II are associated with photostimulated sexual status in male quail. Cell and Tissue Research 300: 413-426. Tixier-Vidal, A. and Follett, B. K. 1973. The adenohypophysis. Pp. 110-182. In D. S. Farner and J. R. King (eds.), Avian Biology. Vol. 3. Academic Press, New York. Tsutsui, K., Bentley, G. E. and Ciccone, N. 2005. Structure, action and functional significance of GnIH. Pp. 73-82. In: A Dawson and P. J. Sharp (eds.) Functional Avian Endocrinology, Narosa Publishing House, New Delhi. Tsutsui, K., Li, D., Azumaya, Y., Muneoka, Y., Minakata, H. and Nomoto, K. 1997. Demonstration, localization, and development of galanin receptors in the quail oviduct. Journal of Experimental Zoology 277: 57-65. Tsutsui, K., Li, D., Ukena, K., Kikuchi, M. and Ishii, S. 1998. Developmental changes in galanin receptors in the quail oviduct and the effect of ovarian sex steroids on galanin receptor induction. Endocrinology 139: 4230-4236. Tsutsui, K., Saigoh, E., Ukena, K., Teranishi, H., Fujisawa, Y., Kikuchi, M., Ishii, S. and Sharp, P. J. 2000. A novel avian hypothalamic peptide inhibiting gonadotropin release. Biochemical and Biophysical Research Communications 275: 661-667.

" Reproductive Biology and Phylogeny of Birds Tsutsui, K. and Ukena, K. 2006. Hypothalamic LPXRF-amide peptides in vertebrates: Identification, localization and hypophysiotropic activity (review). Peptides 27: 1121-1129. Turek, F. W. 1974. Circadian rhythmicity and the initiation of gonadal growth in sparrows. Journal of Comparative Physiology 92: 59-64. Turek, F. W. 1975. The termination of the avian photorefractory period and the subsequent gonadal response. General and Comparative Endocrinology 26: 562564. Ubuka, T., Bentley, G. E., Ukena, K., Wingfield, J. C. and Tsutsui K. 2005. Melatonin induces the expression of gonadotropin-inhibitory hormone in the avian brain. Proceedings of the National Academy of Sciences USA 102: 3052-3057. Ubuka, T., Ueno, M., Ukena, K. and Tsutsui, K. 2003. Developmental changes in gonadotropin-inhibitory hormone in the Japanese quail (Coturnix japonica) hypothalamo-hypophysial system. Journal of Endocrinology 178: 311-318. Ubuka, T., Ukena, K., Sharp, P. J., Bentley, G. E. and Tsutsui, K. 2006. Gonadotropininhibitory hormone inhibits gonadal development and maintenance by decreasing gonadotropin synthesis and release in male quail. Endocrinology 147: 1187-1194. Ueck, M. 1979. Innervation of the vertebrate pineal. Progress in Brain Research 52: 45-88. Ukena, K., Ubuka, T. and Tsutsui, K. 2003. Distribution of a novel avian gonadotropin-inhibitory hormone in the quail brain. Cell and Tissue Research 312: 73-79. Underwood, H. and Menaker, M. 1970. Photoperiodically significant photoreception in sparrows: Is the retina involved? Science 167: 299-301. Underwood, H. and Siopes, T. 1985. Melatonin rhythms in quail: regulation by photoperiod and circadian pacemakers. Journal of Pineal Research 2: 133-143. van Gils, J., Absil, P., Grauwels, L., Moons, L., Vandesande, F. and Balthazart, J. 2003. Distribution of luteinizing hormone-releasing hormones I and II (LHRH-I and -II) in the quail and chicken brain as demonstrated with antibodies directed against synthetic peptides. Journal of Comparative Neurology 334: 304-323. van Tienhoven, A. 1983. Reproductive Physiology of Vertebrates. Cornell University Press. 491 pp. Vaz Nunes, M. and Veerman, A. 1984. Light-break experiments and photoperiodic time measurement in the spider mite Tetranychus urticae. Journal of Insect Physiology 30: 891-897. Vaz Nunes, M. and Veerman, A. 1986. A “dusk” oscillator affects photoperiodic induction of diapause in the spider mite Tetranychus urticae. Journal of Insect Physiology 32: 605-614. Vaugien, L. 1955. Sur les réactions testiculaires du jeune moineau domestique illuminé à diverses époques de la mauvais saison. Bulletin de Biologie de la France et de la Belgique 89:218-244. Veerman, A., Beckman, M. and Veenedaal, R. L. 1988. Photoperiodic induction of diapause in the large white butterfly, Pieris brassicae—evidence for hour glass time measurement. Journal of Insect Physiology 34: 1063-1069. Vitums, A., Mikami, S.-I., Oksche, A. and Farner, D. S. 1964. Vascularization of the hypothalamo-hypophysial complex in the white-crowned sparrow, Zonotrichia leucophrys gambelii. Zeitschrift für Zellforschung und Mikroskopische Anatomie? 64: 541-569.

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Vitums, A., Ono, K., Oksche, A., Farner, D. S. and King, J. R. 1966. The development of the hypophysial portal system in the white-crowned sparrow, Zonotrichia leucophrys gambelii. Zeitschrift für Zellforschung und Mikroskopische Anatomie? 73: 335-366. Wells, J. W. and Gilbert, A. B. 1984. Steroid hormone production by the ovary. Pp. 323-344. In B. M. Freeman (ed.), Physiology and Biochemistry of the Domestic Fowl. Vol. 5. Academic Press, London. Welty, J. C. and Baptista, L. 1988. The Life of Birds. Fourth edition. Saunders College Publishing, New York, NY. 768 pp. Wieselthier, A. S. and van Tienhoven, A. 1972. The effect of thyroidectomy on testicular size and on the photorefractory period in the starling (Sturnus vulgaris L.). Journal of Experimental Zoology 179: 331-338. Williams, T D., Dawson, A., Nicholls, T. J. and Goldsmith, A. R. 1987. Short days induce premature reproductive maturation in juvenile starlings, Sturnus vulgaris. Journal of Reproduction and Fertility 80: 327-333. Wilson, F. E. 1985. Androgen feedback-dependent and independent control of photoinduced LH secretion in male tree sparrows (Spizella arborea). Journal of Endocrinology 105: 141-152. Wilson, F. E. 1991. Neither retinal nor pineal photoreceptors mediate photoperiodic control of seasonal reproduction in American tree sparrows (Spizella arborea). Journal of Experimenal Zoology 117-127. Wilson, F. E. and Donham, R. S. 1988. Daylength and control of seasonal reproduction in male birds. Pp. 101-120. In M. H. Stetson (ed.), Processing of Environmental Information in Vertebrates. Springer-Verlag, Berlin. Wilson, F. E. and Reinert, B. D. 1993. The thyroid and photoperiodic control of seasonal reproduction in American tree sparrows (Spizella arborea). Journal of Comparative Physiology B 163: 563-573. Wilson, F. E. and Reinert, B. D. 1995a. The photoperiodic control circuit in euthyroid American tree sparrows (Spizella arborea) is already programmed for photorefractoriness by week 4 under long days. Journal of Reproduction and Fertility 103: 279-284. Wilson, F. E. and Reinert, B. D. 1995b. A one-time injection of thyroxine programmed seasonal reproduction and postnuptial moult in chronically thyroidectomised male American tree sparrows Spizella arborea exposed to long days. Journal of Avian Biology 26: 225-233. Wingfield, J. C. 1980. Fine temporal adjustment of reproductive functions. Pp. 367389. In A. Epple and M. H. Stetson (eds), Avian Endocrinology. Academic Press, New York. Wingfield, J. C. 1983. Environmental and endocrine control of reproduction: an ecological approach. Pp. 205-288. In S. I. Mikami and M. Wada (eds), Avian Endocrinology: Environmental and Ecological Aspects. Japanese Scientific Societies Press, Tokyo, and Springer-Verlag, Berlin. Wingfield, J. C. 1993. Control of testicular cycles in the song sparrow, Melospiza melodia: Interaction of photoperiod and an endogenous period? General and Comparative Endocrinology 92: 388-401. Wingfield, J. C. 2004. Allostatic load and life cycles: implications for neuroendocrine mechanisms. Pp. 302-342. In J. Schulkin (ed.), Allostasis, Homeostasis and the Costs of Physiological Adaptation. Cambridge University Press, Cambridge. Wingfield, J. C. 2005. Flexibility in annual cycles of birds: implication for endocrine control mechanisms. Journal of Ornithology 146: 291-304.

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Wingfield, J. C. 2006. Communicative behaviors, hormone-behavior interactions, and reproduction in vertebrates. Pp. 1995-2040. In J. D. Neill (ed.), Physiology of Reproduction. Academic Press, New York. Wingfield, J. C. and Farner, D. S. 1978a. The endocrinology of a naturally breeding population of the white-crowned sparrow (Zonotrichia leucophrys pugetensis). Physiological Zoology 51:188-205. Wingfield, J. C. and Farner, D. S. 1978b. The annual cycle in plasma irLH and steroid hormones in feral populations of the white-crowned sparrow, Zonotrichia leucophrys gambelii. Biology of Reproduction 19: 1046-1056. Wingfield, J. C. and Farner, D. S. 1993. The endocrinology of wild species. Pp. 163327. In D.S. Farner, J. R. King and K. C. Parkes (eds), Avian Biology. Vol 9. Academic Press, New York. Wingfield, J. C. and Jacobs, J. D. 1999. The interplay of innate and experiential factors regulating the life history cycle of birds. Pp. 2417-2443. In N. Adams and R. Slotow (eds), Proceedings of the 22nd International Ornithological Congress. BirdLife South Africa, Johannesburg. Wingfield, J. C., Crim, J. W., Mattocks P. W. Jr. and Farner, D. S. 1979. Responses of photosensitive and photorefractory white-crowned sparrows (Zonotrichia leucophrys gambelii) to synthetic mammalian luteinizing hormone releasing hormone (Syn-LH-RH). Biology of Reprodction 21: 801-806. Wingfield, J. C., Follett, B. K., Matt, K. S. and Farner, D. S. 1980. Effect of day length on plasma FSH and LH in castrated and intact white-crowned sparrows. General and Comparative Endocrinology 42: 464-470. Wingfield, J. C., Jacobs, J. and Hillgarth, N. 1997. Ecological constraints and the evolution of hormone-behavior interrelationships. Annals of the New York Academy of Sciences 807: 22-41. Wingfield, J. C., Jacobs, J. D., Tramontin, A. D., Perfito, N., Meddle, S., Maney, D. L. and Soma, K. 1999. Toward an ecological basis of hormone-behavior interactions in reproduction of birds. Pp. 85-128. In K. Wallen and J. Schneider (eds), Reproduction in Context. M.I.T. Press, Cambridge, Massachusetts. Wingfield, J. C., Moore, M. C. and Farner, D. S. 1983. Endocrine responses to inclement weather in naturally breeding populations of white-crowned sparrows (Zonotrichia leucophrys pugetensis). Auk 100: 56-62. Wingstrand, K. G. 1951. The Structure and Development of the Avian Pituitary. Gleerup, Lund. 316 pp. Wingstrand, K. G. 1954. The ontogeny of the neurosecretory system in chick embryos. Pubblicazione Stazione Zoologica di Napoli 24: 27-31. Yamamura T., Hirunagi K., Ebihara S. and Yoshimura T. 2004. Seasonal morphological changes in the neuro-glial interaction between gonadotropin-releasing hormone nerve terminals and glial endfeet in Japanese quail. Endocrinology 145: 4264-4267. Yin, H., Ukena, K., Ubika, T. and Tsutsui, K. 2005. A novel G protein-coupled receptor for gonadotropin-inhibitory hormone in the Japanese quail (Coturnix japonica): identification, expression and binding activity. Journal of Endocrinology 184: 257-266. Yokoyama, K., Oksche, A., Darden, T. R. and Farner, D. S. 1978. The sites of encephalic photoreception in photoperiodic induction of the growth of the testes in the white-crowned sparrow, Zonotrichia leucophrys gambelii. Cell and Tissue Research 189: 441-467. Zann, R. A., Morton, S. R., Jones, K. R. and Burley, N. T. 1995. The timing of breeding by zebra finches in relation to rainfall in central Australia. Emu 95: 208-222.

CHAPTER

6

Ovarian Dynamics and Follicle Development A.L. Johnson and Dori C. Woods

6.1

INTRODUCTION

The avian ovary represents a truly dynamic organ system capable of fostering the annual development of one or more broods of viable eggs, then undergoing nearly complete regression followed by eventual recrudescence (see also Chapter 4 and Volume 6B, Chapter 13). The seasonal initiation of ovarian recrudesence may be driven by endogenous circannual rhythmicity and synchronized by environmental cues, the most important of which is photoperiod and to a more limited extent, light intensity (Gwinner 2003). Such cues are translated into neuroendocrine and endocrine signals, the primary factors being hypothalamic gonadotropin releasing hormone (GnRH) and pituitary gonadotropins, respectively. Subsequent events, which include nesting, follicle maturation and ovulation, are generally initiated by factors other than light (e.g., rainfall, temperature, food availability, male behaviors). A unique morphological and functional aspect of the reproductively active avian ovary, as compared to the mammalian counterpart, is that follicles at all stages of development, from resting primordial and primary follicles to the fully differentiated preovulatory stage, exist simultaneously during egglaying. As a consequence, the sequential selection of one undifferentiated follicle into the final rapid growth stage of development provides for ovulation of an oocyte from a fully differentiated follicle on an approximate daily basis (the interval between ovulations is species-dependent). The ovarian follicular hierarchy is a reflection of oviparity, and is a feature held in common with avian predecessors, the reptiles and apparently some dinosaurs (Sato et al. 2005). Department of Biological Sciences, The University of Notre Dame, Notre Dame, IN 46556, USA

"" Reproductive Biology and Phylogeny of Birds While there are some excellent field studies documenting seasonal changes in reproductive hormones and comparing ovarian dynamics among free ranging species (e.g., see review Wingfield and Farner 1993), the majority of information pertaining to cellular and molecular mechanisms regulating follicle growth and differentiation has been derived primarily from domesticated birds. This is largely due to the quantity and ready availability of tissues required for detailed studies, but nevertheless leaves open the question as to what extent models of ovarian organization and function developed from genetically-manipulated avian models maintained under well-controlled environmental and nutritional conditions directly pertain to wild birds. Accordingly, the following represents a discussion of the avian ovary literature from a comparative and integrative perspective. A primary objective is to highlight some of the many unresolved questions that can assist both field and bench biologists in developing more refined working models of ovarian function to better understand the remarkably diverse and successful reproductive strategies attributed to avian species.

6.2

OVARY MORPHOLOGY AND DEVELOPMENT

The ovary is loosely attached to the peritoneal cavity by the mesovarian ligament (the hilus) at the cephalic end of the kidney, and is suspended from the dorsal body wall by a peritoneal fold, the mesovarium. As in mammals, the avian ovary is covered by a single layer of surface epithelium that displays characteristics associated with the potential for rapid proliferation, and conversely, cell death by apoptosis (Chalana and Guraya 1979). These accessory tissues accommodate the annual growth and regression of the ovary, and in particular, the rapid growth of preovulatory follicles followed by reabsorption of the postovulatory follicle. Unlike the mammalian ovary, where developing follicles largely remain embedded within the cortex, avian follicles beyond the primordial stage of development protrude from ovarian stromal tissue much like grapes in a bunch (Figs. 6.1-6.3). This pendulate architecture enables large amounts of yolk to be incorporated within the follicle during the rapid growth phase. The most prominent features of a reproductively active ovary are the developing follicles that serve to: 1) incorporate liver-derived yolk within the oocyte; 2) provide structural support for growth of the large oocyte (especially compared to that in therian mammals); 3) synthesize and secrete innumerable paracrine and autocrine factors that maintain oocyte viability and promote follicle differentiation (including steroidogenesis), or alternatively promote apoptotic cell death; and 4) release the oocyte in a timely fashion at ovulation. The potential for annual nutritional investment required for follicle growth and maintenance during development is illustrated by the 32 to 43% increase in body mass of the Common eider (Somateria mollissima) from prior to ovarian recrudescence compared to that during peak seasonal reproductive activity. In a few instances, an important consequence of this significant investment

Ovarian Dynamics and Follicle Development

"#

Fig. 6.1 Left ovary of the domestic hen (Gallus gallus) illustrating the hierarchal structure of ovarian follicles. Under environmentally controlled conditions the laying hen will continue to lay eggs on a daily basis for a year or longer with a typical clutch size of 6 to 20+ eggs. F1-5 represent preovulatory follicles 12 to 40 mm in diameter) that have been selected to enter the rapid growth phase of development. SYF, small yellow prehierarchal follicles (6 to 8 mm diameter); SWF, small white prehierarchal follicles (1 to 5 mm). POF, postovulatory follicle. Not readily visible are primordial and primary follicles (<1 mm). Reprinted from Johnson, A.L. 1990. Critical Reviews in Poultry Biology 2: 319-346. Fig. 1.

towards ovarian growth during the reproductive season may be temporary flightlessness and increased risk of predation (Guillemette and Ouellet 2005). The ovary itself increases in weight by greater than 100-fold from the beginning of recrudescence to the peak of lay, and virtually all of this weight represents the growth of yolk-filled follicles (Gilbert 1979).

6.2.1

Development and Asymmetry

The left and right ovaries are present during the early embryonic stages of presumably all birds, but in many species only a functional left ovary is maintained post-hatch. By comparison, some species commonly maintain both gonads (e.g., a number of falcons, eagles, vultures, and the kiwi [Apteryx]; Fig. 6.3), yet these functional ovaries may be asymmetrical in size (the left

"$ Reproductive Biology and Phylogeny of Birds

Fig. 6.2 Ovary from the Gentoo penguin (Pygoscelis papua) at an early and late stage of the breeding cycle. Clutch size in the gentoo is typically two eggs, laid with an interval of 3 to 4 days. A. Prehierarchal (non-vitellogenic) follicles prior to follicle selection. B. Preovulatory follicle, with prehierarchal follicles destined to become atretic. Reprinted from Valencia, J. and Leyton, V. 1992. Gonadal cycles of Pygoscelis penguins of the South Shetland Islands. Pp. 198-207. In W.C. Hamlett (ed.), Reproductive Biology of South American Vertebrates. Springer-Verlag, New York, Fig. 14-3.

Ovarian Dynamics and Follicle Development

"%

characteristically being the larger). It has been reported that in various birds, including sparrows, gulls, doves and pigeons, a small percentage of individual specimens may maintain two active ovaries through adulthood (Kinsky 1971). The left ovary consists of intermingling medullary and cortical tissues (the ovarian stroma), and this supportive tissue is abundantly supplied with blood vessels. Primordial and early developing primary follicles are clustered in distinct masses of cortical tissue. The undeveloped right ovary is composed mainly of medullary tissue with small patches of cortex, and as such, resembles a testis. Removal of the dominant left ovary or exposure to endocrine disruptors (particularly estrogenic compounds) during early embryo development can result in the development of a testis or ovotestis, both of which may be capable of producing spermatozoa (Gilbert 1979; Yoshimura and Fujita 2005). In birds, the female represents the heterogametic sex, with sex chromosomes designated as ZW, while the male is homogametic (ZZ). Although the basic mechanism for genetic sex determination in birds is still unknown, there is evidence that candidate regulatory genes reside on both the W and Z chromosomes (Smith and Sinclair 2004) (see also Chapter 25). For instance, a functional W chromosome is required for regulating aromatase enzyme expression during early embryogenesis (day 5 to 6 of incubation in Gallus domesticus). Activity of this enzyme results in the synthesis of estrogen, and is prerequisite for the development of a functional left ovary (Bruggeman et al. 2002). The effects of estrogen are mediated by estrogen receptors, which are selectively expressed on the left, but not right, gonad in females by day 7. A phenotypic sex reversal in genetic males is affected by treatment, in ovo, with estrogen, while administration of an aromatase inhibitor to genetic females before day 7 will promote the differentiation of testes.

6.2.2 Primordial Germ Cells, Migration and Proliferation During Early Embryogenesis Primordial Germ Cells (PGCs) originate in the epiblast within the central zone of the area pellucida at the early somite stage of chicken embryo development. Following their translocation to the germinal crescent region, the number of PGCs has been estimated at 200-250 (Tsunekawa et al. 2000). Avian PGCs migrate from the germinal crescent to blood vessels by diapedesis and are proposed to exit ovarian blood vessels in response to chemotactic signals from the germinal ridge (Kuwana et al. 1986). This route of PGC transport distinctly differs from that of amphibians and mammals, where PGCs reach the germinal ridge via an extravascular migration. In the incubating pheasant (Phasianus colchicus) embryo the number of circulating PGCs is maximal by 64 h of development (somite number, 21-23), and dramatically decreases thereafter. This decline in circulating PGCs is directly correlated with an increase in the number of germ cells populating the embryonic ovary (Kim et al. 2005). Once PGCs take up residence within the germinal ridge of the ovary, they become

"& Reproductive Biology and Phylogeny of Birds

Fig. 6.3 Paired ovaries from the adult North Island brown kiwi (Apteryx mantelli). Although the left ovary is normally larger and contains a greater number of developing follicles than the right, the number and size of follicles in both ovaries is often similar early in the breeding season. Clutch size in the kiwi is typically one egg, but a second and occasionally a third egg may be laid after intervals of about 25 days. LO, left ovary; LOV, left oviduct; RO, right ovary. Reprinted from Kinsky F.C. 1971. Journal of Ornithology 112: 334-357, Fig. 1, with permission.

Ovarian Dynamics and Follicle Development

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mitotically active oogonia. Although specific factors implicated in PGC survival, chemotaxis and proliferation have, to date, been insufficiently studied in any avian species, a recent report has directly implicated the chemokine, stromal cell-derived factor-1 (SDF-1), in the latter stages of PGC migration (Stebler et al. 2004). In teleost, amphibian and some reptilian species that are capable of spawning hundreds to millions of ova within a reproductive season, the ovarian oogonia constitute self-renewing stem cells that continue to proliferate post-hatch and post-puberty. By comparison, in avian and mammalian species that produce considerably fewer eggs, the oogonia divide a finite number of times and produce the maximum available number of egg precursor cells by the time of hatch or birth. The resulting cells progress through the first meiotic prophase to the diplotene stage, at which point they are called primary oocytes, and are maintained in an arrested state until puberty. The final germ cell number is apparently dependent upon the local hormonal environment, as follicle stimulating hormone (FSH) and estradiol treatments during embryonic development have been reported to delay meiotic arrest and increase the overall size of the oocyte pool (Xie et al. 2004). Resumption of meiosis and completion of the first meiotic division occurs only shortly before the fully matured ovarian follicle is destined to ovulate. The total number of primary oocytes within the domestic chick ovary has been estimated to increase from some 28,000 on the ninth day of incubation to 680,000 on the seventeenth day, then abruptly decrease to 480,000 by the time of hatching or shortly thereafter. Only a small fraction of these primary oocytes (200-500) subsequently develops to the preovulatory stage within the lifespan of most domesticated species, and considerably fewer mature to this stage in wild species. Ultimately, the fate for the vast majority of these potential eggs, both prior and subsequent to hatch, is death via apoptosis (see section 6.3.4). Current dogma dictates that by the time of, or shortly after, hatch, no additional number of germ cells can be attained or generated by the avian ovary. A recent publication, however, has challenged this concept in mammals by providing evidence that female germ line stem cells also take up residence in the bone marrow, and that these stem cells continually migrate to the ovary via the circulatory system over much of the female’s reproductive lifespan (Johnson, Bagley et al. 2005). Should this concept of germ cell renewal prove true in the avian ovary, there would exist numerous, significant implications towards the potential for enhancing the propagation of threatened and endangered species.

6.2.3

Sexual Maturation

Many species of wild birds hatch their eggs in spring or early summer, and the offspring attain adult body size within a few weeks. In many respects, the ovary of such fully-grown juveniles appears comparable in size and morphology to that of photorefractory adults (Williams et al. 1987). Nevertheless, these

# Reproductive Biology and Phylogeny of Birds offspring do not attain sexual maturity until the following breeding season presumably because their photoperiodic responsiveness is comparable to the photorefractory state of adult birds. Note that opportunistic breeders represent an exception to this paradigm (see section 6.2.5). By comparison, some larger, long-lived birds, including albatrosses, condors and penguins, may take 5 to 10 years to reach sexual maturity.

6.2.4

Seasonal Reproduction

The onset of each reproductive season in free-range birds can typically be separated into two distinct phases of ovarian growth, each with different environmental cues controlling their onset (Jacobs and Wingfield 2000). In photoperiodic species such as the sparrow (Zonotrichia), the first, or proximate, phase is initiated by a lengthening photoperiod that increases hypothalamic secretion of GnRH, followed by increases in circulating levels of FSH. The initial growth of the ovary culminates after approximately six weeks, when follicles reach a size of 2 to 3 mm in diameter but are as yet undifferentiated (‘sub-functional’). Female sparrows will generally not progress to the second phase of rapid growth and final differentiation until appropriate supplementary information is provided. While the source of this second set of supplementary cues is dependent upon species and latitude at which it breeds, such ultimate factors likely include: the availability of food, suitable nesting conditions, and interactions with a male. For example, exposure to male song alone has been reported to augment the rate of follicle development in White-crowned sparrows (Zonotrichia leucophrys) (Morton et al. 1985). During the reproductive season, avian species will develop and ovulate a characteristic number of eggs (a clutch) prior to the initiation of incubation. While there is the tendency for larger birds to lay fewer eggs, the evolutionary forces that dictate clutch size remain elusive. For instance, clutch size is often smaller in tropical and sub-tropical birds compared to those in temperate northern climates, despite the prediction that abundant food availability in tropical regions should provide for optimal egg laying. A recent study by Martin et al. (2000) attempted to reconcile clutch size as a function of food availability compared to the risk of predation. Their data support the hypothesis that higher rates of nest predation in subtropical regions serve as a constraint to the rate at which parent birds can provide food to the nestlings, thereby limiting the number of young they can feed, and by implication, clutch size. Alternative, and not necessarily mutually exclusive, hypotheses implicate physiological constraints (e.g., large clutches place more physiological strain on females than small clutches), plus seasonal and habitat constraints as factors regulating clutch size. In lesser snow geese (Anser caerulescens), it has been proposed that the clutch size can be adjusted to the female’s nutrient reserve plane at a time just before or at the onset of laying. This adjustment can occur by the reabsorption of the smallest developing follicles, and assures adequate development of the remaining follicles plus a

Ovarian Dynamics and Follicle Development

#

favorable nutrient plane for the nesting female. The loss of potential offspring is offset by a higher probability of successful fledging (Hamann et al. 1986). Many species will produce a single clutch per season (determinant layers; e.g., white pelican, Pelecanus erythrorrhynchos, blue-winged teal, Anas discors, barn swallow, Hirundo rustica). Others may produce more than one clutch and/or extend laying within a clutch, particularly if the clutch is destroyed or the first eggs of the clutch are removed (indeterminant layers; e.g., European starling, Sturnis vulgaris, song sparrow, Melospiza melodia, common flicker, Colaptes auratus, barn owl, Tyto alba). There is evidence that the extension of a clutch in indeterminant breeders can be accomplished by the rescue of small preovulatory follicles from atresia, rather than the selection of additional follicles into the preovulatory hierarchy (Challenger et al. 2001). Finally, species that invest a prolonged period of time to raise their young may not breed every year (e.g., king penguin, Aptenodytes patogonica; some species of albatrosses, Diomedeidae). This strategy is typically associated with advanced age of sexual maturity (albatrosses as late as 10 years of age) and longevity of adult survival (Jouventin and Dobson 2002).

6.2.5

Opportunistic Breeders

Free-ranging birds that live in areas where environmental changes are cyclical and highly predictable typically exhibit regular phases of annual ovarian recrudescence, regression and refractoriness. By comparison, those species that live in regions where environmental changes are less predictable or entirely unpredictable, lack appropriate proximal factors to cue ovarian recrudescence in a timely fashion. Accordingly, such species must opportunistically respond to ultimate factors as rapidly as possible to insure the coincidence of fertility with optimal breeding and rearing conditions. One example of an opportunistic breeder is the Zebra finch (Taeniopygia (=Poephila) guttata) that has adapted to the semi-arid and arid regions of Australia. Female finches are reported to undergo ovarian growth beginning shortly after hatch and continuing throughout the first 100 days. Thereafter, the developing ovarian follicles enter an indefinite resting stage where, following an appropriate ultimate cue (e.g., rainfall), selected follicles can enter the rapid growth phase and be ovulated within one to two weeks (Sossinka 1980). In contrast to photoperiodic species (McNaughton et al. 1992), the precocious ovarian development observed in the zebra finch following hatch is attributed to the absence of any detectable refractory period. Moreover, the White-winged crossbill (Loxia leucoptera) inhabits an Alaskan region where its primary food source, conifer seeds, can vary in quantity from year to year. Results from endocrinological studies (including the seasonal content of hypothalamic GnRH) support the view that these birds have the potential to respond to ultimate factors and to breed throughout the year depending upon food availability, except for the period of fall molt when elevated levels of prolactin presumably suppress ovarian activity (Deviche and Sharp 2001).

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These two examples further illustrate the adaptiveness of birds to extreme and variable environmental conditions, but also raise important questions as to how the ovary maintains an appropriate milieu to support long-term prehierarchal follicle viability (e.g., prevent cellular apoptosis and consequent atresia over a period from weeks to months), and how the presence of ultimate factors becomes translated into appropriate cellular signals to initiate follicle selection into the rapid growth phase. Such questions can be appropriately addressed only in free-ranging birds, and the mechanisms will likely be species-dependent.

6.2.6

Reproductive Aging and Ovarian Senescence

Reproductive aging can be defined as any decline in reproductive performance, and includes a decrease in fitness of the offspring (Saino et al. 2002). By comparison, ovarian senescence more specifically constitutes a significant decline or loss of the primary follicle reserve within the ovary, and/or the significant loss of oocyte quality (capacity for fertilization). Reproductive senescence is often preceded by a gradual decline in egg production. Associated with this decline in the domestic hen is an overall increase in atresia specifically in follicles within the slow growth phase, and a decreased rate of follicle selection into the preovulatory hierarchy (Waddington et al. 1985). Holmes et al. (2003) have described female reproductive aging in birds according to three paradigms. The first is characterized by a short life span with relatively rapid declines in fertility (e.g., many Galliformes, including domesticated species). A second consists of moderately slow aging birds with a long life span (greater than 10 years) where the slow decline in reproductive success correlates closely with increased mortality (e.g., passerine songbirds, including the Great tit, Parus major, and small raptors like the European sparrowhawk, Accipiter nisus). The third model is characterized by long life spans, slow aging, and negligible reproductive declines in either sex. For instance, neither the Common tern (Sterna hirundo; Nisbet et al. 1999) nor the Nazca boobie (Sula granti; Anderson and Apanius 2003) appears to show an appreciable decline in reproductive effort even after 20 years of reproductive activity. Some terns and gulls have been reported to actually increase the number of offspring they successfully fledge with advancing age, though this may be more related to post-hatch survival and parental behaviors than to enhanced ovarian fecundity. These proposed models, though not yet rigorously tested, raise important questions regarding the role of the ovarian environment in promoting long-term germ cell survival and viability. Unfortunately, there appears to be no published information on primary oocyte reserves with aging in species which have differing rates of reproductive longevity, nor of cellular mechanisms that might account for the apparent variability in the onset of ovarian senescence among avian species.

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Vascularization and Nervous Innervation of the Ovary

The entire process of follicle growth can be conveniently divided into: 1) the organization of primordial follicles and the early growth of primary follicles, the latter of which may persist for years; 2) the recruitment of primary follicles into the slow growth phase (the prehierarchal or previtellogenic stage) lasting weeks to perhaps months; and 3) the selection and rapid growth of preovulatory follicles which occurs during the final days preceding ovulation. The continued viability and development of ovarian follicles within a breeding season is critically dependent upon endocrine factors (e.g., gonadotropins) circulated by the blood, paracrine and autocrine factors produced within the ovary itself, plus neurochemical and neurohumoral factors expressed by the nervous system. Blood supply to the avian ovary generally originates from the renal artery and exits the ovary directly into the caudal vena cava. Prior to the initiation of growth, primary follicles remain embedded within the highly vascular stromal tissue and receive blood from adjacent capillaries. To service growing follicles that begin to extrude from the stromal tissue, the ovarian artery supplies two to four smaller arteries that enter the follicle via the follicular stalk (or pedicle). While small arteries and arterioles extend throughout the follicle theca layers, there is no direct blood supply to the inner granulosa cell layer (Fig. 6.4). Not unexpectedly, blood flow is proportionately greatest to the most mature follicles within the preovulatory hierarchy. In most species, an avascular stigma region forms where the follicle fuses with the surrounding surface epithelium, and serves as the site of rupture at ovulation. Lymphocytes and macrophages circulate throughout ovarian stromal tissue and the follicle theca layer (Barua and Yoshimura 1999), and these cells represent a potential source of cytokines that can influence follicle viability (see section 6.3.4.2). Immune competence of tissues within growing and postovulatory follicles is implied by the reported expression of major histocompatibility complex (MHC) molecules within both the theca and granulosa layers of follicles throughout development (Subedi and Yoshimura 2005). The ovary is well innervated by both adrenergic and cholinergic fibers. Nerve fibers enter through the pedicle and radiate through the theca (but not granulosa) layer. The number of neurons within the theca layer increases as a follicle progressively matures. It is clear that such innervation provides a variety of neurochemicals (e.g., catecholamines) and neurohumoral factors (e.g., neurotropins, vasoactive intestinal peptide, substance P) to the ovary and developing follicles. Such factors are proposed to function in diverse roles such as regulation of steroidogenesis in the embryonic ovary (MullerMarschhausen et al. 1988), the organization and recruitment of primary follicles (Gilbert 1979), and the growth and differentiation of growing follicles in the adult (Johnson et al. 1994; Jensen and Johnson 2001). More generally, it is likely that ovarian innervation plays an integral role in the seasonal

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Fig. 6.4 Organization of domestic hen ovarian follicles during development. Primary oocytes enclosed by the vitelline membrane become organized into a primordial follicle (up to ~80 m diameter) following the recruitment of presumptive granulosa cells, and the perivitelline membrane is subsequently formed by granulosa cells. The initiation of primordial follicle growth to the primary follicle stage is associated with the formation of the theca layer (from mesenchymal cells), which is separated from the granulosa layer by the basal lamina. Primary follicles range in size from .08 to 1 mm in diameter. Further growth to the prehierarchal follicle stage (1-8 mm) entails the accumulation of lipoprotein-rich, white yolk, plus the differentiation of the theca into interna and externa layers. Vasculature and nervous innervation reaches the follicle through the pedicle and radiates through the theca layer. Following selection into the preovulatory hierarchy, preovulatory follicles rapidly grow from 9 mm to 40 mm over the course of days. Granulosa cells from preovulatory follicles facilitate the uptake of large amounts of vitellogenin and very low density lipoprotein, except within the germinal disc region. Ovulation of the largest preovulatory follicle eventually occurs at the region of the comparatively avascular stigma region. See text for additional details.

recrudescence of the ovary. Of interest is one report that functional innervation persists in the regressed right ovary of the hen (Ohmori et al. 1994). While the physiological significance of such innervation has not been established, it may contribute to the generation of testicular tissue following abnormal loss of the functional left ovary during early development.

6.3 STEROIDOGENESIS AND CELLULAR MECHANISMS MEDIATING FOLLICLE GROWTH AND DIFFERENTIATION Unfortunately, the majority of information pertaining to cellular and molecular mechanisms regulating follicle growth, selection and final differentiation is derived from a limited number of domesticated species (e.g., Gallus gallus, Coturnix japonica, Meleagris gallopavo). Nevertheless, such studies have begun to provide an understanding of ovarian processes common to the vertebrate

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archosaur and synapsid lineages, plus those unique to the oviparous bird. It is emphasized, however, that the working models presented will undoubtedly require modifications to accommodate both reproductive seasonality under free-ranging conditions, and the diversity of reproductive strategies among birds. Additional, detailed discussions of circulating hormonal profiles relative to the ovulatory cycle, cell signaling, steroidogenesis, and follicle selection can be found elsewhere (Wingfield and Farner 1993; Johnson 1990, 2000; Woods and Johnson 2005).

6.3.1

Primordial and Primary Follicles

Beginning shortly after hatch, primary oocytes become organized into primordial follicles following the recruitment of a single somatic (presumptive granulosa) cell layer (Fig. 6.4); oocytes that fail to become enclosed die via apoptosis. Granulosa cells of primordial follicles are cuboidal, densely packed and coupled via tight junctions. Such multifunctional cells have been implicated in a number of critical processes during follicle growth and differentiation, including meiotic arrest, perivitelline membrane formation, gonadotropin-induced hormone production, mediation of yolk protein (vitellogenin) uptake, and transfer of maternal RNAs to the egg. Primordial follicles do not increase in size appreciably and have the potential to survive in this arrested state of development for years. The transition to a primary follicle represents the stage when primordial follicles are recruited from the resting state to begin slow growth. While a variety of growth factors have been implicated in initiating (e.g., stem cell factor, basic fibroblast growth factor) or inhibiting (anti-Müllerian hormone) follicle recruitment in mammals (Skinner 2005), these have apparently not been investigated in the avian ovary. Mesenchymal cells are subsequently recruited by one or more factors presumably secreted from granulosa cells (possibly including stem cell factor), to form a peripheral theca layer. The granulosa and theca layers are separated by an acellular basement membrane (basal lamina) that effectively isolates the granulosa layer from receiving a direct supply of blood and nervous tissue (Fig. 6.4). The morphology of the basal lamina and its relationship to the granulosa layer have recently been described by Asem et al. (2000). The single layer of theca will eventually differentiate into morphologically and functionally distinct theca interna and externa layers. While primary follicle growth is proposed to occur independent of gonadotropins, follicle viability remains dependent upon the support of both gonadotropin and growth factors (Johnson et al. 1996b). This earliest growth phase can progress over several years, and in the turkey the overall growth is from approximately 80 mm to 1 mm in diameter, primarily via an increase in the size of the oocyte without an increase in the germinal vesicle (Carlson et al. 1996). Primordial follicles and early developing primary follicles remain embedded within the ovarian stromal tissue, and thus lack a surrounding epithelial cell layer. The collective stromal tissue containing such follicles

#$ Reproductive Biology and Phylogeny of Birds expresses both FSH and luteinizing hormone (LH) receptors, and is steroidogenically active when challenged with LH, in vitro, as short term incubations produce progestin, androstenedione and estradiol (Levorse and Johnson 1994). Moreover, isolated primary follicles (<1 mm) produce both androgen (dehydroepiandrosterone and androstenedione) and estradiol in response to LH treatment (Robinson et al. 1988). While the amount of gonadotropin-induced steroid production at this stage of development is not particularly impressive on a per follicle basis, the cumulative steroid production is considered physiologically significant based upon the large total number of such follicles present within the ovary. The expression of gonadotropin receptors or steroidogenic enzymes within the seasonally regressed ovary has apparently not been evaluated in any species, however it is predicted that resting ovarian follicles remain competent to produce some level of steroids, even during the nonbreeding season.

6.3.2

Prehierarchal Follicles

A second, comparatively more rapid phase of growth lasting for a period of weeks to perhaps months occurs as previtellogenic (1-5 mm diameter prehierarchal follicles) follicles begin to incorporate small amounts of a lipoprotein-rich, white yolk. More follicles than are ultimately required to establish the preovulatory hierarchy are recruited into this growth phase, thus the fate of many prehierarchal follicles is atresia (see section 6.3.4). Although the exact rate of atresia among all prehierarchal follicles has not been precisely estimated, it is assumed that it is rather high, perhaps up to 20% (Gilbert et al. 1983). Conditions that can promote atresia at this stage include inadequate growth factor and gonadotropin support of survival, and genetic or morphological defects. As the follicle continues to grow it protrudes from the surface of the stromal tissue and becomes enveloped by the ovarian epithelium. Internally, an acellular perivitelline layer (homologous to the zona pellucida of mammals) forms from the granulosa cells and appears as electron-dense granules. The granulosa cells themselves are densely packed and form projections that extend into the oocyte plasma (vitelline) membrane. The theca has now differentiated into morphologically and functionally distinct externa and interna layers. By the end of this prehierarchal phase, follicles in Gallus and Zonotrichia reach the size of 6 to 8 mm and 2 to 3 mm, respectively. In freeranging birds, the termination of this slow growth phase corresponds to the ‘sub-functional’ level of development as described by Jacobs and Wingfield (2000). Much of the lipoprotein-rich yolk incorporated during the early phase of prehierarchal follicle growth may transverse the granulosa cell layer either by transcytosis or, perhaps more importantly, by a paracellular transport route. Nevertheless, the amount of yolk transported is limited at this stage of development by the tight junctions (and a major tight junction protein, occludin) that serve to connect adjacent granulosa cells and restrict paracellular transport (Schuster et al. 2004). Occludin expression is regulated

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by the stimulatory effects of FSH and the transforming growth factor-b (TGFb) superfamily member, activin. Eventually, the levels of occludin associated with tight junctions gradually decrease such that progressively greater amounts of yolk, including the lipid-rich yellow yolk, transverse the granulosa cell layer to reach the oocyte. Thus, 6 to 8 mm prehierarchal follicles begin to assume a yellow color (often referred to as small yellow follicles). A single receptor type has been implicated in the uptake of the two major yolk precursors, very low-density lipoprotein (VLDL) and vitellogenin (VTG), by the oocyte (Schneider 1996). This oocyte-specific VTG/VLDL receptor is initially localized within the central portion of the cell in slow growing follicles, and rapid yolk uptake occurs only following its redistribution to the oocyte vitelline membrane (Shen et al. 1993).

6.3.2.1

Granulosa and theca cells

The theca interna of prehierarchal follicles becomes both innervated and vascularized, and initiates pregnenolone and androgen (largely androstenedione) production. Structural support of the follicle is provided by sheets of collagen fibers and fibronection plus smooth muscle located within the theca externa, which gives this layer its stratified appearance (Fig. 6.4). The externa layer contains groups of cells selectively expressing aromatase activity, and represents the sole site within the follicle for the conversion of androgens to estradiol (Nitta et al. 1991). Collectively, the theca layers express comparatively high levels of both LH receptor (Johnson et al. 1996a) and FSH receptor (You et al. 1996) mRNA. While treatment of whole prehirearchal follicles or isolated theca layers with either LH or FSH, in vitro, promotes secretion of progestin, androgen and estrogen, LH is comparatively more potent (Robinson et al.,1988; Kowalski et al. 1991). In contrast, granulosa cells from prehierarchal follicles are incapable of synthesizing significant levels of the progesterone precursor, pregnenolone, due to the absence of cytochrome P450 side-chain cleavage (P450scc) enzyme activity. In addition, these cells lack detectable levels of the cholesterol transport protein, steroidogenic acute regulatory (StAR) protein, required for the transfer of cytoplasmic cholesterol from the outer to the inner mitochondrial membrane (the site of P450scc activity), plus cytochrome P450 17a-hydroxylase (P450-17a-OH) required for androgen production. Granulosa cells at this stage of development express FSH receptor (but not readily detectable LH receptor) mRNA and produce the second messenger, cyclic adenosine monophosphate (cAMP), in response to FSH treatment, in vitro. Because StAR, P450scc and P450-17a-OH expression are primarily dependent upon the cAMP/protein kinase A cell signaling pathway, the absence of steroidogenesis in prehierarchal follicle granulosa cells, in vivo, has been proposed to be the result of tonic inhibitory cell signaling at a site distal to cAMP production (Woods et al. 2005). Nevertheless, it is important to note that treatment of such granulosa cells with FSH, in vitro, initiates StAR, P450scc and P450-17a-OH expression plus production of modest amounts of progesterone and androstenedione (Li and Johnson 1993; Johnson et al.

#& Reproductive Biology and Phylogeny of Birds 2002b). Finally, granulosa cells from prehierarchal follicles demonstrate the capacity for proliferation (Tilly et al. 1992), and this is promoted by one or more members of the epidermal growth factor (EGF) family, FSH, and possibly the TGFb superfamily member, growth differentiation factor 9 (GDF9). EGF ligands represent paracine/autocrine factors produced within the oocyte, the granulosa or theca layers of the follicle or ovarian stromal tissue, whereas GDF9 is produced specifically by the oocyte (Yao and Bahr 2001; Johnson, Dickens et al. 2005; Woods et al. 2005). With regard to free-ranging species, plasma levels of estrogen in Zonotrichia are observed to increase shortly after the females arrive at the breeding area (Wingfield and Farner 1993). This increase in estrogen production occurs at a time when the ovary presumably contains primarily prehierarchal follicles (e.g., prior to when granulosa cells become steroidogenically active). Accordingly, the capacity of the theca layer from prehierarchal follicles to produce estrogen no doubt represents a mechanism to provide for the stimulation of VTG/VLDL synthesis required for the imminent rapid growth phase.

6.3.2.2

Follicle selection into the preovulatory hierarchy

Follicle selection represents the process that ultimately determines the characteristic number of eggs to be ovulated (clutch size) by any given species. A prehierarchal follicle selected into the preovulatory hierarchy immediately begins the process of growth and final differentiation, and this is reflected by both the ability of the follicle to rapidly incorporate yellow yolk, and of the granulosa layer to initiate increasingly large (mg) amounts of progesterone production. While the ultimate signal(s) and cellular events that provide a single follicle the selective advantage to initiate rapid growth and final differentiation have yet to be described in birds (or for that matter, any vertebrate), the consequent differentiation that occurs within the granulosa layer within the hen follicle immediately following selection has been the subject of recent investigations. Among the endocrine, paracrine and autocrine factors that have been implicated in directly initiating steroidogenesis and/or enhancing gonadotropin (FSH and LH) receptor expression in granulosa cells from prehierarchal follicles are included FSH and the TGFb superfamily members, TGFb and activin. Both TGFb and activin signal via type I plus type II receptormediated phosphorylation of the Smad2 regulatory protein, and a primary action in granulosa cells is to enhance FSH receptor expression (Johnson et al. 2004). The actions of activin, but not TGFb, can be modulated by the presence of an activin-binding protein, follistatin. However, as noted above, evidence supports the proposal that tonic inhibition of FSH-mediated StAR, P450scc, and P45017a-OH expression, in vivo, renders granulosa cells from prehierarchal follicles steroidogenically incompetent. Such inhibitory signaling is also proposed to prevent any increase in FSH receptor number and to preclude premature LH receptor expression, thus maintaining all but one prehierarchal follicle in an

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undifferentiated state despite the continued, daily exposure of all follicles to circulating concentrations of FSH. In fact, FSH-induced P450scc and StAR expression plus progesterone production, in vitro, is completely blocked by EGF family ligands (including EGF, transforming growth factor-a and betacellulin) signaling via the mitogen activated protein kinase (MAPK)/extracellular regulated kinase (Erk) pathway. Moreover, MAPK/Erk signaling blocks TGFb- and activin-induced receptor signaling by preventing the phosphorylation required for Smad signaling. Significantly, inhibition of Erk signaling using the selective MAP kinase inhibitor, U0126, not only promotes a synergistic enhancement of StAR expression and progesterone production compared to FSH treatment alone, but also promotes greatly enhanced expression of the FSH receptor plus the initiation of LH receptor expression (Woods and Johnson 2005). Accordingly, a model depicting events initiated within the granulosa layer immediately following the selection of a prehierarchal follicle into the preovulatory hierarchy is presented in Fig. 6.5.

6.3.3 Preovulatory Follicles, Ovulation and Postovulatory Follicles The preovulatory phase is the shortest, and perhaps most characteristic of the avian ovary, and culminates with ovulation. In general, its duration tends to be associated with species size, weight of the egg and precociousness of the newly hatched bird. For instance, the rapid-growth phase typically lasts between 6 and 11 days in the domestic hen and barn owl, approximately 12 days in the Canada goose (Branta canadensis) and up to 16 days in some penguins, but only 5 to 7 days in the Ring-necked dove (Streptopelia capicola) and Japanese quail (Coturnix japonica) (Gilbert 1979). By comparison, the Brown kiwi (Apteryx mantelli) has one of the largest egg-to-body ratios and the most yolk as a percentage of total egg weight (61%) among birds. Accordingly, the rapid growth phase is estimated to occur over a period of some 15 to 17 days and results with an egg size of some 80+ mm. Combined with a comparatively long, 70- to 90- day incubation period (conducted exclusively by the male), these adaptive characteristics enable the kiwi embryo to hatch at an advanced state of development (Jensen and Durrant 2006). Considerably more work has been conducted to characterize cellular mechanisms mediating rapid growth (e.g., yolk incorporation) and final differentiation (e.g., steroidogenesis) in preovulatory follicles compared to those follicles prior to selection, and only an updated summary will be provided here.

6.3.3.1

Preovulatory follicles

Rapid growth of preovulatory follicles occurs largely due to the enhanced capacity for VTG and VLDL uptake by receptor-mediated endocytosis. A second receptor, with preference for binding LDL, is localized to granulosa cells and the theca externa layer in preovulatory follicles, and selective expression of this LDL receptor is associated with supplying LDL-derived cholesterol as precursor for steroid production (Hummel et al. 2003). While the

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Fig. 6.5 Proposed model for cellular events mediating the initiation of steroidogenesis and gonadotropin receptor expression in granulosa cells immediately subsequent to follicle selection. A. Both type I and type II TGFb plus Fig. 6.5 Contd. ...

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majority of the oocyte surface is actively engaged in the uptake of yellow yolk, the 2 to 3 mm diameter germinal disc (the portion of oocyte containing the nucleus, most cellular organelles and the largest amount of cytoplasm) remains visible as a white plaque due to the comparatively lower expression of VTG/VLDL receptors. The comparative reduction of yolk incorporation at this site presumably facilitates the migration of pronuclei and the process of fertilization. As rapid follicle growth proceeds, granulosa cells distal to the germinal disc undergo remodeling to become squamous in appearance and less densely packed (Fig. 6.4) due to a reduction of cell-to-cell connections (tight junctions). This latter change facilitates the paracellular transport of some 2 g of yellow yolk per day into domestic hen preovulatory follicles. Accordingly, follicle mass increases from approximately 0.15 g in 6 to 8 mm prehierarchal follicles to some 12 to 14 g just prior to ovulation. Increased production of the yolk precursors, VTG and VLDL, by the liver occurs in response to increasing estrogen concentrations produced by the developing follicles. The total energetic costs of yolk formation can account for 40% to 50% of the bird’s daily energy budget, and the extra energy required results in a 13% to 41% increase in basal metabolic rate in passerines, to more than a 200% increase in some waterfowl (Meijer and Drent 1999). Given that such an investment of protein synthesis and the production of yolk precursors are energetically expensive, it can be predicted that the supply of yolk precursors should closely match the demand for vitellogenin uptake by preovulatory follicles. In fact, plasma levels of vitellogenin in female zebra Fig. 6.5 Contd. ...

activin receptors are expressed within the granulosa cell layer. While the autocrine/ paracrine factor, TGFb, can bind to its respective receptors, the bioactivity of activin is modulated by follistatin (produced within granulosa cells). It is propoed that active induction of FSH receptor (FSH-R) expression by TGFb (and possibly activin) is precluded in all but one prehierarchal follicle per day by one or more EGF family ligand binding to one or more ErbB receptor and the tonic activation of MAPK/Erk signaling. The sites of MAPK/Erk-mediated tonic inhibition include the absence of Protein Kinase A/cAMP signaling pathway both prior and subsequent to cAMP formation, and the prevention of Smad2 phosphorylation (Smad2-P). The absence of phosphorylation prevents transport of regulatory (r-) Smad2 to the nucleus via the co-regulatory Smad, co-Smad4. B. Following selection, a transient attenuation of MAPK signaling enables TGFb (and activin) signaling via Smad, and results in enhanced FSH-R expression (1). Elevated FSH-R expression subsequently facilitates FSH-induced LH-R expression (2), P450scc and StAR expression, and as a consequence, progesterone production (3). Mechanisms proposed to attenuate MAPK/Erk signaling, in vivo, in the single follicle selected into the preovulatory hierarchy per day include the rapid upregulation of Erk-specific phosphatases. For additional details, see text and Woods and Johnson (2005); Woods et al. (2005). Reprinted, with modifications, from Woods, D.C. and Johnson, A.L. 2005. Biology of Reproduction 72: 643-650, Fig. 9.

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finches (Taeniopygia guttata) are undetectable in non-breeders as well as following termination of a six-egg clutch. In contrast, circulating vitellogenin is dramatically increased at the onset of the rapid yolk development stage, highest at the one- and three-egg stage of the clutch (in excess of 1.4 mg vitellogenin per ml plasma), and significantly decreased by the fifth-egg stage in advance of clutch completion (Salvante and Williams 2002). This ability to precisely regulate yolk precursor availability according to the changing mass of developing follicles as a clutch progresses has similarly been documented for the European starling (Challenger et al. 2001). In addition to yolk, a developing follicle accumulates both maternal messenger RNA (mRNA) and hormones, factors that have been proposed to influence oocyte and early zygote viability plus embryo development. In preovulatory follicles of the Japanese quail, two distinct pools of maternallyderived mRNA have been described. One pool exists within the oocyte germinal disc region, while a second, larger pool is localized in the cytoplasmic layer surrounding the yolk (Malewska and Olszanska 1999). The amount of RNA in the germinal disc region prior to fertilization varies little from that in the blastoderm at oviposition, but RNA content at the vitelline membrane is found to decrease by 80% from fertilization to oviposition. Although the exact timing of genome activation in the avian embryo is not known, it is proposed to occur subsequent to ovulation and the early cleavage stages but prior to oviposition. A variety of transcripts, presumably of maternal origin, have been identified in the unfertilized blastodisc, including those encoding pro- and anti-apoptotic proteins (Muscarella et al. 1998). In particular, the expression and translation of maternally-derived antiapoptotic genes prior to the initiation of embryonic transcription may provide enhanced resistance of the early zygote to environmental stressors, such as fluctuations in temperatures that can occur during the laying of a clutch and the incubation period. Similarly, there is considerable information regarding yolk steroid and thyroid hormones of maternal origin. Exposure to such hormones during embryo development has been associated with the programming of reproductive organs, early somatic growth, development of the brain, sexual behavior and immune function, even long after fledging (Groothuis et al. 2005). For example, in the Japanese quail the yolk content of thyroxine (T3) and triiodothyronine (T4) has been positively associated with accelerated cartilage growth and differentiation in the developing embryo (Wilson and McNabb 1997), while levels of yolk corticosterone have been negatively associated with the growth rate of the growing chick (Hayward and Wingfield 2004). The incorporation of maternally-derived progesterone, testosterone and estradiol has been described within the yolk of Dark-eyed junco (Junco hyemalis) and Red-winged blackbird (Agelaius phoeniceus) eggs (Lipar et al. 1999). Patterns of steroid deposition conform to the concentric rings of yolk incorporated by the follicle on a daily basis, with concentrations of progesterone highest in the most peripheral layers, estradiol concentrations highest near the center of the

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yolk, and testosterone concentrations highest in the intermediate layers. This pattern of steroid deposition largely reflects the temporal changes in steroid production by preovulatory follicles during development (Bahr et al. 1983) and may result in the embryo being exposed to varying levels of hormones during embryonic development. Nevertheless, despite the potential adaptive value of this non-genetic mechanism of inheritance, a critical question is whether the biased allocation of hormones and other factors into the yolk of developing follicles represents a flexible maternal strategy to influence the phenotype of offspring, or alternatively, a passive consequence of maternal physiology at the time of follicle development.

6.3.3.2

Preovulatory follicle granulosa and theca cells

A significant change in gonadotropin receptor expression occurs in the granulosa layer during the transition of a selected prehierarchal follicle into preovulatory stage of development. Granulosa cells from preovulatory follicles demonstrate predominant expression of LH receptor, while FSH receptor mRNA declines to low, but still detectable, levels. Progesterone production is controlled by LH, and LH-induced cAMP formation increases dramatically as a follicle approaches the time of ovulation. Fully potentiated production of progesterone, androgens and estradiol by preovulatory follicles requires the participation of granulosa cells combined with cells located in both the theca interna and externa layers (Nitta et al. 1991). The portion of the granulosa layer distal to the germinal disc is the predominant source of progesterone that serves as the primary steroid involved in potentiating the preovulatory surge of LH that precedes and subsequently induces ovulation. In addition, granulosa-derived progesterone serves as precursor for androstenedione, and testosterone, synthesis by the theca layer, and to a lesser extent by granulosa cells. Steroid production by the theca layer is also regulated almost exclusively by LH, despite the continued expression of FSH receptor mRNA. Significantly, LH receptor mRNA levels and LH-induced cAMP plus androstenedione formation dramatically declines specifically within the theca from the largest preovulatory follicle (Johnson et al. 1996a; Marrone and Hertelendy 1985). At the same time, granulosa cells from the largest preovulatory follicle become capable of producing considerably greater amounts of progesterone, both in vivo and in vitro, compared to those from the second largest follicle. This enhanced progesterone production is associated with significantly increased mitochondrial activity and/or number of mitochondria together with increased levels of P450scc mRNA, but is not directly related to enhanced LH-induced StAR protein expression (Dive et al. 1992; Johnson et al. 2002b). Presumably, the combination of these events ensures maximal progesterone secretion from the largest preovulatory follicle at the time of the preovulatory LH surge. It is important to note, however, that granulosa cells differ not only in morphology relative to their proximity to the germinal disc (see section

$" Reproductive Biology and Phylogeny of Birds 6.3.3.1), but also phenotype. Preovulatory follicle granulosa cells localized adjacent to the germinal disc region (animal pole) remain mitotically active, whereas those cells distal to the germinal disc (vegetal pole) become nonmitotic (Perry et al. 1978; Tilly et al. 1992). Moreover, cells at the vegetal pole express comparatively less EGF (ErbB1 and ErbB4) receptor mRNA but more LH receptor mRNA and produce more progesterone in response to LH compared to those at the animal pole (Tischkau et al. 1997; Yao and Bahr 2001). Finally, germinal disc-derived factors (including one or more EGF family members) can stimulate granulosa proliferation and inhibit progesterone production, in vitro (Tischkau and Bahr 1996). Accordingly, these findings are consistent with the model proposed for the initiation of granulosa cell differentiation following follicle selection (see section 6.3.3.2), in that granulosa cells are maintained in a proliferative and undifferentiated state until released from tonic inhibition. It is further speculated that processes leading to the eventual differentiation of granulosa cells within the vegetal pole are those described for prehierarchal follicles following selection (Fig. 6.5). Finally, granulosa and theca cells produce additional endocrine/ paracrine/autocrine factors whose functions in follicle development are currently being investigated. For instance, the TGFb superfamily members, inhibins (in particular, inhibin A) are produced by granulosa cells from preovulatory (particularly the largest) follicles (Johnson, Brooks et al. 2005), and are proposed to negatively modulate pituitary FSH secretion. However, a negative relationship between plasma levels of inhibin A and FSH during the ovulatory cycle has yet to be conclusively demonstrated. Inhibins may also competitively antagonize the actions of activin by its association with betaglycan and the activin type II receptor (Knight et al. 2005).

6.3.3.3

Ovulation and postovulatory follicle

Final maturation of the oocyte and ovulation of the largest preovulatory follicle is triggered primarily by a surge of pituitary-derived LH that, in domesticated birds (e.g., Gallus gallus, Coturnix japonica, Meleagris gallopavo) precedes ovulation by 4 to 6 hours. The initiation of germinal vesicle breakdown occurs coincident with peak concentrations of circulating LH, and condensation of chromatin, extrusion of the first polar body, and formation of the second maturation spindle are completed 2.5 to 1 h prior to ovulation. At the same time, interdigitations and the remaining tight junctions between granulosa cell cytoplasmic projections and the oocyte plasma membrane dissociate, and a perivitelline space develops as a result of fluid accumulation (Yoshimura et al. 1993). Significantly, such changes are not induced within the second largest preovulatory follicle. Although the cellular mechanisms providing for this selectivity are not yet clear, it is reasonable to propose a role for the local, high concentrations of progesterone produced within the largest preovulatory follicle. At the time of ovulation, rupture of the follicle occurs along the stigma by a combination of factors: 1) activation of proteases (including collagenase) that

Ovarian Dynamics and Follicle Development

$#

reduce tensile strength in the follicle wall; 2) increased intrafollicular pressure due to increased blood flow to the follicle; 3) induction of smooth muscle contractions in the follicle wall; 4) a weakening of granulosa cell support following the retreat of microvilli from the surface of the oocyte; and 5) a selective loss of cells within the stigma region via apoptosis. The structure remaining after release of the oocyte is the postovulatory follicle (POF), and contains both granulosa and theca tissues. This structure is not homologous to the mammalian corpus luteum in that the POF remains steroidogenically active for, at most, a few days following ovulation. Cell death via apoptosis is initiated shortly after ovulation, and the POF is largely reabsorbed within several days. Regression of the POF may be further aided by the presence of immunocompetent cells expressing the MHC class II antigen (Barua et al. 2001). Importantly, the most recent POF of the domestic hen exerts a direct or indirect influence on the oviduct, as removal of the largest (but not next most recent) POF is associated with delayed oviposition of the egg derived from that follicle. This rapid loss of functional postovulatory tissue presumably insures that a direct influence on oviposition is limited to that egg present within the reproductive tract. The POF, like the mammalian corpus luteum, reportedly expresses relaxin (Brackett et al. 1997), yet its role in oviposition, if any, has yet to be described.

6.3.4

Follicle Atresia

Vertebrate follicle atresia represents the death of an organized follicle at any time during development beginning with the primordial stage through the ovulatory stage. Atresia is generally considered a normal physiological process by which surplus or non-viable growing follicles are rapidly reabsorbed by a non-inflammatory process, though it can also be abruptly induced by environmental or physiological perturbations. It is significant to note, however, that among vertebrate species the overall proportion of ovarian follicles lost by atresia, as well as the time during follicle development at which atresia can occur, differs according to reproductive strategy. In particular, those animals that produce thousands to perhaps millions of mature gametes within a reproductive cycle (spawners; e.g. fish) show a very low overall incidence of atresia, and most of the oogonia produced during the lifespan of the female are ovulated as mature oocytes. By comparison, those groups that produce a considerably more limited number of offspring (many reptiles, birds, mammals) demonstrate a proportionately higher incidence of atresia. Furthermore, within a normal reproductive cycle a major difference between reptiles and birds compared to mammals is that in the former species, atresia occurs prior to selection into the preovulatory hierarchy, whereas in mammals atresia of subdominant follicles occurs following selection of the dominant follicle(s) (Rothchild 2003). The following sections will first address stages of the avian reproductive cycle when follicle atresia can occur, then some cellular mechanisms mediating this process.

$$ Reproductive Biology and Phylogeny of Birds

6.3.4.1

Avian follicle atresia

In birds, the loss of follicles by atresia can occur any time during the active breeding season, at the termination of breeding (following the onset of photorefractoriness or at the start of incubation), and during seasonal molt (Erpino 1973), and the overall rate of atresia has been observed to increase with advancing age (Waddington et al. 1985). Under optimal breeding conditions atresia rarely occurs in preovulatory follicles, but can be induced at this final stage of development by inappropriate environmental conditions (e.g., during forced molt induced by food deprivation and/or reduction in photoperiod) and a decline in circulating gonadotropins. By contrast, follicle atresia frequently occurs in prehierarchal follicles (Gilbert et al. 1983). Thus, it has been proposed that a transition from atresia-susceptibility to atresiaresistance is initiated coincident with follicle selection into the preovulatory hierarchy (Johnson 2003). A variety of factors contributes to the survival of preovulatory follicles. It is clear that the integrity of the germinal disc region is required for follicle viability given that its destruction results in the rapid onset of atresia in the affected follicle (Yoshimura and Bahr 1995; Yao et al. 1998). The critical supportive factors produced by the oocyte have not been unequivocally identified, but likely include one or more growth factor(s), including those from the EGF family (Yao and Bahr 2001). Continued support by circulating gonadotropins is a second factor, as hypophysectomy leads to the loss of all follicles within the hierarchy. Conversely, in vivo treatment of chicken or quail hens with FSH or equine chorionic gonadotropin (eCG) decreases the rate of atresia and increases the overall number of prehierarchal and preovulatory follicles. Interestingly, Challenger et al. (2001) provide evidence that the clutch size in starlings is increased, not by the selection of additional prehierarchal follicles, but by rescue of the smallest, most recently selected preovulatory follicles from atresia. This suggests that in at least some indeterminant layers (including the barn owl, Durant et al. 2004, and Eurasian kestrel, Meijer et al. 1989), there are more follicles selected into the preovulatory hierarchy than will normally be fully developed and ovulated. In the event that eggs are lost during the early portion of the clutch, a prolonged or increased secretion of survival factors (e.g., FSH and LH from the pituitary and/or growth factors from ovarian tissues) may serve to support the continued growth of supplemental follicles. The comparatively high incidence of atresia that occurs in prehierarchal follicles that have yet to enter the rapid growth phase represents a reproductive strategy that maximizes the availability of healthy prehierarchal follicles readily available for selection into the preovulatory hierarchy, yet minimizes the loss of energetically expensive maternal resources that would otherwise be invested in extensive VTG/VLDL deposition. There is evidence that non-growing follicles embedded within cortical tissue of the mature ovary also undergo atresia, but the overall incidence has not been reported.

Ovarian Dynamics and Follicle Development



%$Underlying causes of follicle loss at this early stage of development include genetic or meiotic anomalies.

6.3.4.2 Apoptosis as modulator of ovarian function and as a proximal cause of follicle atresia It has been recognized for some time that apoptosis is a proximal cause of follicle atresia in mammals and birds (Tilly et al. 1991). In particular, apoptosis is observed first in isolated granulosa cells, and subsequently progresses throughout the granulosa layer. This rapid progression of apoptosis is facilitated by an increase in the number of gap junctions that provide for cell-to-cell communication (Krysko et al. 2004) and the transfer of apoptosis-inducing factors. Signals initiating granulosa cell apoptosis can variably originate from the oocyte, from the granulosa layer itself, or from tissues peripheral to the basement membrane (theca or circulatory system). As atresia continues, apoptosis eventually progresses throughout the theca layers. Additionally, during the late stages of embryogenesis and the early post-hatch period, there are data to suggest that apoptosis occurs in ovarian cortical follicles, and that the incidence of apoptosis decreases to negligible levels by day 14 post-hatch (Yoshimura and Nishikori 2004). These latter observations can explain the decline in available germ cell number during the latter stages of embryo development (see section 6.2.2). Finally, the reabsorption of postovulatory follicle tissues also occurs via apoptosis, and is initiated within the first 24 h after ovulation (Tilly et al. 1991). Accordingly, apoptosis represents a fundamental process by which the avian ovary accomplishes: 1) the regression of the right ovary early in ontogeny; 2) a reduction of germ cell number by the time of hatch; 3) the selection of a limited number of the most viable follicles for ovulation; 4) the rapid elimination of postovulatory tissues as a clutch progresses; and 5) the reduction of ovarian mass that accompanies the non-breeding season. There are several reviews detailing cellular mechanisms involved in the regulation of apoptosis specifically in the avian ovary (Johnson 2003; Johnson and Bridgham 2002), and only an updated summary will be provided here. Apoptosis involves a complex and well-coordinated sequence of intracellular signaling events that results in the elimination of affected cells without the initiation of an inflammatory reaction. The entire process can be divided into phases of initiation, execution and termination, with the first of these two phases dependent upon the activation of one or more cysteine proteases (caspases). Terminal endpoints include membrane blebbing, cell shrinkage, the formation of membrane-enclosed vesicles (apoptotic bodies), proteolytic cleavage of numerous structural plus functional proteins, and ultimately DNA fragmentation (oligonucleosome formation). In the domestic hen, the comparatively high incidence of follicle atresia in prehierarchal follicles, in vivo, correlates with susceptibility of the granulosa cell layer to undergo apoptosis as observed, in vitro (Johnson et al. 1996b). In light of this observation, and because apoptosis in granulosa cells represents

$& Reproductive Biology and Phylogeny of Birds the earliest event described during the onset of atresia, most studies of the ovary have focused primarily on this cell type. The progression and amplification of the apoptotic process in granulosa cells is initiated by either intrinsic or extrinsic pathways (Fig. 6.6). Activation of an intrinsic pathway can occur by a variety of factors, including cellular (e.g., oxidative) stress, and/or the withdrawal of growth factor support. Either of these stimuli may be initiated by the lack of adequate food or the withdrawal of growth factor and/or gonadotropin support, as typically occurs with photorefractoriness. At the cellular level, such stimuli cause perturbations in mitochondrial function that result in the release of mitochondrial proteins, including cytochrome C, into the cytoplasm. Cytosolic cytochrome C (cytC) promotes the formation of an apoptosome complex consisting of Apoptosis Protease Activating Factor-1 (Apaf-1) and the initiator caspase, caspase-9. In turn, activated caspase-9 activates the executioner caspase, caspase-3, a primary mediator of the terminal endpoints mentioned above, together with the proteolysis of many structural and catalytic proteins (e.g., Poly-(ADP-ribose)-polymerase, PARP). By comparison, an extrinsic pathway can be activated by a variety of cytokines that bind to one or more receptors containing intracellular death domains (DD) (Fig. 6.6). Such receptors belong to an evolutionarily conserved family of death receptors known as the Tumor Necrosis Factor Receptor Superfamily (TNFRSF) (Bridgham et al. 2003). Ligand-activated death receptors recruit cytoplasmic adaptor proteins (e.g., Fas-Associated Death Domain, FADD) by the homodimerization of death domains (DD). In turn, FADD recruits and activates an alternative initiator caspase, caspase-8, through the homodimerization of their respective death effector domains (DED). Activated caspase-8 cleaves the cytoplasmic protein, BH3 Interacting Domain (Bid) death agonist, that promotes release of mitochondrial cytC, as well as activation of the executioner caspase, caspase-3. From this point, the downstream events converge with those described for the intrinsic pathway. No less than five DD-containing receptors from this death receptor superfamily are expressed by hen granulosa cells: TNFRSF1 (TNF receptor type 1); TNFRSF6 (Fas); TNFRSF10B (DR5); TNFRSF16 (p75 Nerve Growth Factor receptor); and TNFRSF23 (Bridgham and Johnson 2004). Several known cytokines are produced in a paracrine or autocrine fashion by one or more cell types within the ovary, and these include Tumor Necrosis Factor a (TNFa; binds to TNFRSF1), TNF-Related Apoptosis Inducing Ligand (TRAIL; binds to TNFRSF10B), and Fas Ligand (TNFRSF6). Moreover, potential deathinducing ligands can be trafficked to the follicle via nervous innervation (e.g., NGF; binds to TNFRSF16) or by immune cells from the vasculature (e.g., TNFa, TRAIL). Intuitively, the finding that no less than five death domain-containing receptors are expressed within this cell type implies that multiple ligands may regulate cell viability under varying environmental and physiological conditions. To date, TNFa has been demonstrated to promote hen granulosa cell apoptosis, in vitro, in granulosa collected from atresia-susceptible

CMYK $'

Fig. 6.6 Simplified model of pro- and anti-apoptotic pathways in hen granulosa cells. Apoptotic cell death can be initiated by either an intrinsic or extrinsic pathway, both of which converge at the activation of caspase-3. Activation of this executioner enzyme is irreversible, and results in the cleavage of numerous structural and functional proteins (e.g., PARP) and the internucleosomal cleavage of genomic DNA. Cell death pathways are opposed by anti-apoptotic proteins that protect mitochondrial membrane integrity (Bcl-x, Bcl-2), block caspase activity (IAPs) or prevent signaling via death receptors (the adaptor protein, FLIP). Expression of these anti-apoptotic proteins is regulated by gonadotropins (FSH and LH) and locally produced growth factors (IGF-I and EGF family ligands) via cell survival signaling pathways. The absence of sufficient anti-apoptotic protein expression (dotted green lines) is proposed to tip the balance in favor of activating proapoptotic pathways (solid red lines). See text for further details. Abbreviations: aCasp.-3, -7, -8, activated caspases; Apaf-1, Apoptosis Protease Activating Factor-1; Bid and tBid, intact or truncated BH3 Interacting Domain death agonist; cytC, cytochrome C; DD, death domain; DED, death effector domain; EGF, epidermal growth factor; FADD, Fas-Associated Death Domain; FLIP, Flice-like inhibitory protein; FSH, follicle-stimulating hormone; Inhibitor of Apoptosis Protein, IAP; IGF-I, insulin-like growth factor I; LH, luteinizing hormone; PARP, Poly-(ADP-ribose)polymerase; PKA/cAMP, protein kinase A/cyclic AMP signaling pathway; PKB/Akt, protein kinase B/Akt signaling pathway. Each of the proteins and cell signaling pathways depicted has been characterized in cultured hen granulosa cells. For a more complete description with citations, see Bridgham et al. 2003; Johnson 2003 and Johnson and Bridgham 2002.

CMYK

CMYK

CMYK

Ovarian Dynamics and Follicle Development

% Reproductive Biology and Phylogeny of Birds prehierarchal follicles (Witty et al. 1996). Studies are ongoing to elucidate the ability of additional cytokines (e.g., TRAIL and Fas) to promote granulosa cell apoptosis, and the conditions under which this can occur. The finding that cellular components of both the intrinsic and extrinsic cell death pathways are constitutively expressed, clearly indicates that mechanisms exist to counteract their actions. Not surprisingly, granulosa cells express a variety of anti-apoptotic proteins that prevent initiator and effector caspase activity (Inhibitor of Apoptosis Proteins, IAPs) or prevent mitochondrial membrane perturbations (Bcl-2, Bcl-x). In addition, a decoy adaptor protein that mimics caspase-8 but lacks caspase activity (Flice-Like Inhibitory Protein, FLIP) can block death receptor activity by preventing the recruitment of caspase-8 (Fig. 6.6). In fact, immediately subsequent to follicle selection, granulosa cells from atresia-resistant preovulatory follicles have been demonstrated to express significantly higher levels of Bcl-x, Bcl-2 and IAP compared to granulosa from atresia-susceptible prehierarchal follicles (Johnson et al. 1997, 1998, 1999). Furthermore, the activation of cell survival signaling pathways, such as the protein kinase A pathway, enhances expression of IAP and Bcl-x protein in hen granulosa cells while PKB signaling promotes cell cycle progression and survivin expression (Johnson et al. 2002a). To a large extent, it is by tipping the balance in favor of such antiapoptotic mechanisms that the gonadotropins (FSH and LH) and locally produced growth factors (EGF family members, insulin-like growth factor-I) are proposed to support granulosa cell, and by implication follicle, viability throughout the preovulatory stage of development.

6.3.5

Avian Clock Genes and Reproduction

Finally, it is clear from the above discussion that circannual and circadian rhythmicity is a fundamental component of seasonal reproduction. A central site of seasonal (periodic) time measurement in birds has been localized to the mediobasal hypothalamus (MBH), and lesions within the avian MBH can block photoperiodic-induced increases in gonadotropin secretion and gonadal growth. Such photoperiodic responsiveness is mediated at the molecular level by oscillations in the expression of clock genes, and each of the known avian clock genes required to retain a steady state photoinducible phase have now been localized to the MBH (e.g., Clock, Per, Bmal, Cry, E4bp4) (Yasuo et al. 2003). Significantly, circadian clock genes are also expressed in the ovary of the rat, and their expression is influenced by the preovulatory LH surge (Karman and Tischkau 2006). Accordingly, it is predicted that each of the phases within the avian reproductive cycle (seasonality, follicle recruitment, follicle selection, and ovulation-oviposition cycles) can be explained at a molecular level by the ebb and flow expression of clock genes. A challenge for the future is to expand our knowledge of molecular clocks to the avian ovary in an effort to elucidate the means by which seasonal and daily fluctuations of environmental cues (e.g., photoperiod) are ultimately translated and/or modified at the level of the ovary to influence follicle recruitment, selection and the timing of ovulation.

Ovarian Dynamics and Follicle Development

6.4

%

ACKNOWLEDGMENTS

The authors thank Drs. Jamie T. Bridgham and Tom Jensen for their many contributions to portions of the data discussed herein, and Morgan Haugen for help with the preparation and editing of the manuscript. We acknowledge the National Science Foundation (IBN31185, IOB45949) and the National Institutes of Health (HD-36095) for recent research support.

6.5

LITERATURE CITED

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Johnson, A. L., Bridgham, J. T. and Wagner, B. 1996a. Characterization of a chicken luteinizing hormone receptor (cLH-R) complementary deoxyribonucleic acid, and expression of cLH-R messenger ribonucleic acid in the ovary. Biology of Reproduction 55: 304-309. Johnson, A. L., Bridgham, J. T., Witty, J. P. and Tilly, J. L. 1996b. Susceptibility of avian ovarian granulosa cells to apoptosis is dependent upon stage of follicle development and is related to endogenous levels of bcl-xLong gene expression. Endocrinology 137: 2059-2066. Johnson, A. L., Bridgham, J. T., Witty, J. P. and Tilly, J. L. 1997. Expression of bcl-2 and nr-13 in hen ovarian follicles during development. Biology of Reproduction 57: 1096-1103. Johnson, A. L., Bridgham, J. T. and Woods, D. C. 2004. Cellular mechanisms and modulation of Activin A- and transforming growth factor b-mediated differentiation in cultured hen granulosa cells. Biology of Reproduction 71: 18441851. Johnson, A. L., Langer, J. S. and Bridgham, J. T. 2002a. Survivin as a cell cycle-related and antiapoptotic protein in granulosa cells. Endocrinology 143: 3405-3413. Johnson, A. L., Li, Z., Gibney, J. A. and Malamed, S. 1994. Vasoactive intestinal peptide-induced expression of cytochrome P450 cholesterol side-chain cleavage and 17 alpha-hydroxylase enzyme activity in hen granulosa cells. Biology of Reproduction 51: 327-333. Johnson, A. L., Solovieva, E. V. and Bridgham, J. T. 2002b. Relationship between steroidogenic acute regulatory protein expression and progesterone production in hen granulosa cells during follicle development. Biology of Reproduction 67: 1313-1320. Johnson, J., Bagley, J., Skaznik-Wikiel, M., Lee, H. J., Adams, G. B., Niikura, Y., Tschudy, K. S., Tilly, J. C., Cortes, M. L., Forkert, R., Spitzer, T., Iacomini, J., Scadden, D. T. and Tilly J. L. 2005. Oocyte generation in adult mammalian ovaries by putative germ cells in bone marrow and peripheral blood. Cell 122: 303-315. Johnson, P. A., Brooks, P. A. and Davis, A. J. 2005. Pattern of secretion of immunoreactive inhibin/activin subunits by avian granulosa cells. General and Comparative Endocrinology 141: 233-239. Johnson, P. A., Dickens, M. J., Kent, T. R. and Giles, J. R. 2005. Expression and function of growth differentiation factor-9 in an oviparous species, Gallus domesticus. Biology of Reproduction 72: 1095-1100. Jouventin, P. and Dobson, F. S. 2002. Why breed every other year? The case of albatrosses. Proceedings of the Royal Society, London B 269: 1955-1961. Karman, B. N., Tischkau, S. A. 2006. Circadian Clock Gene Expression in the Ovary: Effects of Luteinizing Hormone. Biology of Reproduction 2006. Jun 28; [Epub ahead of print] Kim, J. N., Lee, Y. M., Park, T. S., Jung, J. G., Cho, B. W., Lim, J. M. and Han, J. Y. 2005. Detection and characterization of primordial germ cells in pheasant (Phasianus colchicus) embryos. Theriogenology 63: 1038-1049. Kinsky F. C. 1971. The consistent presence of paired ovaries in the Kiwi (Apteryx) with some discussion of this condition in other birds. Journal of Ornithology 112: 334-357. Knight, P. G., Gladwell, R. T. and Lovell, T. M. 2005. The inhibin-activin system and ovarian folliculogenesis in the chicken. Pp 323-337. In A. Dawson and P.J. Sharp (eds.), Functional Avian Endocrinology. Narosa Publishing House, New Delhi.

%" Reproductive Biology and Phylogeny of Birds Kowalski, K. I., Tilly, J. L. and Johnson, A. L. 1991. Cytochrome P450 side-chain cleavage (P450scc) in the hen ovary. I. Regulation of P450scc messenger RNA levels and steroidogenesis in theca cells of developing follicles. Biology of Reproduction 45: 955-966. Krysko, D. V., Mussche, S., Leybaert, L. and D’Herde, K. 2004. Gap junctional communication and connexin43 expression in relation to apoptotic cell death and survival of granulosa cells. Journal of Histochemistry & Cytochemistry 52: 11991207. Kuwana, T., Maeda-Suga, H. and Fujimoto, T. 1986. Attraction of chick primordial germ cells by gonadal anlage in vitro. Anatomical Record 215: 403-406. Levorse, J. M. and Johnson, A. L. 1994. Regulation of steroid production in ovarian stromal tissue from 5- to 8-week-old pullets and laying hens. Journal of Reproduction and Fertility 100: 195-202. Li, Z. and Johnson, A. L. 1993. Regulation of cholesterol side-chain cleavage messenger ribonucleic acid expression and progesterone production in hen granulosa cells. Biology of Reproduction 49: 463-469. Lipar, J. L., Ketterson, E. D., Nolan, V. and Casto, J. M. 1999. Egg yolk layers vary in the concentration of steroid hormones in two avian species. General and Comparative Endocrinology 115: 220-227. Malewska, A. and Olszanska, B. 1999. Accumulation and localisation of maternal RNA in oocytes of Japanese quail. Zygote 7: 51-59. Marrone, B. L. and Hertelendy, F. 1985. Decreased androstenedione production with increased follicular maturation in theca cells from the domestic hen (Gallus domesticus). Journal of Reproduction and Fertility 74: 543-550. Martin, T. E., Martin, P. R., Olson, C. R., Heidinger, B. J. and Fontaine, J. J. 2000. Parental care and clutch sizes in North and South American birds. Science 287: 1482-1485. McNaughton, F. J., Dawson, A. and Goldsmith, A. R. 1992. Puberty in birds: the reproductive system of starlings does not respond to short days until birds are fully grown. Journal of Endocrinology 132: 411-417. Meijer, T. and Drent, R. 1999. Re-examination of the capital and income dichotomy in breeding birds. Ibis 141: 399-414. Meijer, T., Masman, D. and Daan, S. 1989. Energetics of reproduction in female kestrels. The Auk 106: 549-559. Morton, M. L., Pereya, M. E. and Baptista, L. 1985. Photoperiodically-induced ovarian growth in the white-crowned sparrow (Zonotrichia leucophrys gambelii) and its augmentation by song. Comparative Biochemistry and Physiology 80A: 93-97. Muller-Marschhausen, U., Grothe, C., Kaveri, S., Strosberg, A. D., Verhofstad, A. A. and Unsicker, K. 1988. Catecholamine nerves in the embryonic chick ovary: colocalization with beta 2-adrenoceptor-bearing steroidogenic cells. Cell and Tissue Research 254: 1-9. Muscarella, D. E., Rachlinski, M. K. and Bloom, S. E. 1998. Expression of cell death regulatory genes and limited apoptosis induction in avian blastodermal cells. Molecular Reproduction and Development 51: 130-142. Nitta, H., Yoshio, Y. and Bahr, J. M. 1991. Multiple steroidogenic cell populations in the thecal layer of preovulatory follicles of the chicken ovary. Endocrinology 129: 2033-2040. Nisbet, I. C., Finch, C. E., Thompson, N., Russek-Cohen, E., Proudman, J. A. and Ottinger, M. A. 1999. Endocrine patterns during aging in the common tern (Sterna hirundo). General and Comparative Endocrinology 114: 279-286.

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Ohmori Y., Kawaguchi T. and Watanabe T. 1994. Location of efferent and afferent neurons innervating the ovary in the hen. Journal of the Autonomic Nervous System 47(1-2): 1-5. Perry, M. M., Gilbert, A. B. and Evans, A. J. 1978. Electron microscope observations on the ovarian follicle of the domestic fowl during the rapid growth phase. Journal of Anatomy 125: 481-497. Robinson, F. E., Etches, R. J., Anderson-Langmuir, C. E., Burke, W. H., Cheng, K. W., Cunningham, F. J., Ishii, S., Sharp, P. J. and Talbot, R. T. 1988. Steroidogenic relationships of gonadotrophin hormones in the ovary of the hen (Gallus gallus). General and Comparative Endocrinology 69: 455-466. Rothchild, I. 2003. The yolkless egg and the evolution of eutherian viviparity. Biology of Reproduction 68: 337-357. Saino, N., Ambrosini, R., Martinelli, R. and Moller, A. P. 2002. Mate fidelity, senescence in breeding performance and reproductive trade-offs in the barn swallow. Journal of Animal Ecology 71: 309-319. Salvante, K. G. and Williams, T. D. 2002. Vitellogenin dynamics during egg-laying: daily variation, repeatability and relationship with egg size. Journal of Avian Biology 33: 391-398. Sato, T., Cheng, Y. N., Wu, X. C., Zelenitsky, D. K. and Hsiao Y. F. 2005. A pair of shelled eggs inside a female dinosaur. Science 308(5720): 375. Schneider, W. J. 1996. Vitellogenin receptors: oocyte-specific members of the lowdensity lipoprotein receptor supergene family. International Reviews in Cytology 166: 103-137. Schuster, M. K., Schmierer, B., Shkumatava, A. and Kucher, K. 2004. Activin A and follicle-stimulating hormone control tight junctions in avian granulosa cells by regulating occluding expression. Biology of Reproduction 70: 1493-1499. Shen, X., Steyer, E., Retzek, H., Sanders, E. J. and Schneider, W. J. 1993. Chicken oocyte growth: receptor-mediated yolk deposition. Cell and Tissue Research 272: 459-471. Skinner, M. K. 2005. Regulation of primordial follicle assembly and development. Human Reproduction Update 11: 461-471. Smith, C. A. and Sinclair, A. H. 2004. Sex determination: Insights from the chicken. Bioessays 26: 120-132. Sossinka, R. 1980. Ovarian development in an opportunistic breeder, the Zebra Finch Poephila guttata castanotis. The Journal of Experimental Zoology 211: 225-230. Stebler, J., Spieler, D., Slanchev, K., Molyneaux, K. A., Richter, U., Cojocaru, V., Tarabykin, V., Wylie, C., Kessel, M. and Raz, E. 2004. Primordial germ cell migration in the chick and mouse embryo: the role of the chemokine SDF-1/ CXCL12. Developmental Biology 272: 351-361. Subedi, K. and Yoshimura, Y. 2005. Expression of MHC class I and II in growing ovarian follicles of young and old laying hens, Gallus domesticus. The Journal of Poultry Science 42: 101-109. Tilly, J. L., Kowalski, K. I., Johnson, A. L. and Hsueh, A. J. 1991. Involvement of apoptosis in ovarian follicular atresia and postovulatory regression. Endocrinology 129: 2799–2801. Tilly, J. L., Kowalski, K. I., Li, Z., Levorse, J. M. and Johnson, A. L. 1992. Plasminogen activator activity and thymidine incorporation in avian granulosa cells during follicular development and the periovulatory period. Biology of Reproduction 46: 195-200.

%$ Reproductive Biology and Phylogeny of Birds Tisckkau, S. A. and Bahr, J. M. 1996. Avian germinal disc region secretes factors that stimulate proliferation and inhibit progesterone production by granulosa cells. Biology of Reproduction 54: 865-870. Tischkau, S. A., Neitzel, L. R., Walsh, J. A. and Bahr, J. M. 1997. Characterization of the growth center of the avian preovulatory follicle. Biology of Reproduction 56: 469-474. Tsunekawa, N. Naito, M., Sakai, T. and Noce, T. 2000. Isolation of chicken vasa homolog and tracing the origin of primordial germ cells. Development 127: 27412750. Valencia, J. and Leyton, V. 1992. Gonadal cycles of Pygoscelis penguins of the South Shetland Islands. Pp 198-207. In W.C. Hamlett (ed.), Reproductive Biology of South American Vertebrates. Springer-Verlag, New York. Waddington, D., Perry, M. M., Gilbert, A. B. and Hardie, M. A. 1985. Follicular growth and atresia in the ovaries of hens (Gallus domesticus) with diminished egg production rates. Journal of Reproduction and Fertility 74: 399-405. Williams, T. D., Dawson, A., Nicholls, T. J. and Goldsmith, A. R. 1987. Short days induce premature reproductive maturation in juvenile starlings, Sturnis vulgaris. Journal of Reproduction and Fertility 80: 327-333. Wilson, C. M. and McNabb, F. M. A. 1997. Maternal thyroid hormones in Japanese quail eggs and their influence on embryonic development. General and Comparative Endocrinology 107: 153-165. Wingfield, J. C. and Farner, D. S. 1993. Endocrinology of reproduction in wild species. Pp. 163-327. In D. S. Farner, J. R. King and K. C. Parkes (eds.), Avian Biology, Vol. 9. Academic Press, London. Witty, J. P., Bridgham, J. T. and Johnson, A. L. 1996. Induction of apoptotic cell death in hen granulosa cells by ceramide. Endocrinology 137: 5269-5277. Woods, D. C., Haugen, M. J. and Johnson, A. L. 2005. Opposing actions of TGFb and MAP kinase signaling in undifferentiated hen granulosa cells. Biochemical and Biophysical Research Communications 336: 450-457. Woods, D. C. and Johnson, A. L. 2005. Regulation of follicle-stimulating hormonereceptor messenger RNA in hen granulosa cells relative to follicle selection. Biology of Reproduction 72: 643-650. Xie, M., Zhang, C., Zeng, W. and Mi, Y. 2004. Effects of follicle-stimulating hormone and 17b-estradiol on proliferation of chick embryonic germ cells in culture. Comparative Biochemistry and Physiology, Part A 139: 521-527. Yao, H. H. and Bahr, J. M. 2001. Germinal disc-derived epidermal growth factor: a paracrine factor to stimulate proliferation of granulosa cells. Biology of Reproduction 64: 390-395. Yao, H. H., Volentine, K. K. and Bahr, J. M. 1998. Destruction of the germinal disc region of an immature preovulatory chicken follicle induces atresia and apoptosis. Biology of Reproduction 59: 516-521. Yasuo, S., Watanabe, M., Okabayashi, N. Ebihara, S. and Yoshimura, T. 2003. Circadian clock genes and photoperiodism: comprehensive analysis of clock gene expression in the mediobasal hypothalamus, the suprachiasmatic nucleus, and the pineal gland of Japanese quail under various light schedules. Endocrinology 144: 3742-3748. You, S., Bridgham, J. T., Foster, D. N. and Johnson, A. L. 1996. Characterization of the chicken follicle-stimulating hormone receptor (cFSH-R) complementary deoxyribonucleic acid, and expression of cFSH-R messenger ribonucleic acid in the ovary. Biology of Reproduction 55: 1055-1062.

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Yoshimura, Y. and Bahr, J. M. 1995. Atretic changes of follicular wall caused by destruction of the germinal disc region of an immature preovulatory follicle in the chicken: an electron microscope study. Journal of Reproduction and Fertility 105: 147-151. Yoshimura, Y. and Fujita, M. 2005. Endocrine disruption in avian reproduction: the histological analysis. Avian and Poultry Biology Reviews 16: 29-40. Yoshimura, Y. and Nishikori, M. 2004. Identification of apoptotic oocytes in the developing ovary of embryonic and post-hatch chicks of Japanese quail (Coturnix japonica). The Journal of Poultry Science 41: 140-146. Yoshimura, Y., Okamoto, T. and Tamura, T. 1993. Electron microscope observations on LH-induced oocyte maturation in Japanese quail (Coturnix coturnix japonica). Journal of Reproduction and Fertility 98: 401-407.

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CHAPTER

7

Spermatogenesis and Testicular Cycles Tom A. Aire

7.1

INTRODUCTION

The seminiferous epithelium in sexually mature and active testis is made up of germ cells at varying levels of development, and Sertoli cells, which are the only somatic, non-dividing cells of the epithelium. Sertoli cells have been described in some detail in Chapter 2. Sertoli cells have multiple functions, including, but not limited to, contributing significantly to the formation of the blood-testis barrier by means of the so-called Sertoli-Sertoli junctional complexes (Dym and Fawcett 1970; Fawcett et al. 1970; Dym 1973), providing anchorage and nutrition for, as well as regulation of, germ cells during development (Morris et al. 1987; Jégou 1991; Skinner et al. 1991; Vogl et al. 1991). The germ cells usually develop, and grow older as they move from the basement membrane of the seminiferous epithelium toward the tubular lumen. Thus, the most primitive or immature germ cells lie on the basement membrane and the mature germ cells, the spermatozoa, line the lumen of the seminiferous tubule. Germ cells develop in close association with one another because as they divide they maintain close linkage through intercellular bridges which are the result of incomplete cytoplasmic divisions (Fawcett 1961).

7.2 7.2.1

SPERMATOCYTOGENESIS Spermatogonia

Spermatogonia are the most immature or primitive germ cells that divide to produce the cells which eventually differentiate into spermatozoa in the seminiferous epithelium of physiologically active testes. Spermatogonia divide Department of Anatomy and Physiology, Faculty of Veterinary Science, University of Pretoria, Onderstepoort, Republic of South Africa. E-mail: [email protected]

& Reproductive Biology and Phylogeny of Birds repeatedly by mitosis to produce spermatocytes, as well as preserving (by selfreplication) a store of stem cells that continue the process of spermatogenesis. The replacement of these dividing stem cells has received considerable attention in mammals (Regaud 1901; Roosen-Runge and Giesel 1950; Clermont and Leblond 1953) but to a much lesser extent in birds (Marchand 1977; Lin and Jones 1992). The existence of several types of spermatogonia in the seminiferous epithelium was first recognized by Regaud (1901) in Rattus norvegicus, the laboratory rat. Several authors subsequently reported varying numbers of generations of these cells in different mammalian species. Ortavant (1959) describes three successive generations of spermatogonia in mammals: type A whose nucleus is dust-like, as first described by Regaud (1901); type B whose nucleus exhibits crust-like chromatin attachments to the inner part of the nuclear membrane. The third generation of spermatogonia, the intermediate spermatogonium, has also been described in several mammalian species (see Courot et al. 1970). Several successive generations of type A spermatogonia have been described subsequently in a number of species of mammals (Courot et al. 1970; Clermont and Bustos-Obregon 1968; Huckins 1971a,b,c; Oakberg 1971a,b; Lok et al. 1982; Hadley and Dym 1983). Thus, Clermont and BustosObregon (1968) have recognized, in R. norvegicus, five successive generations of type A spermatogonia (A 0, A1, A2, A3 and A4) and Ekstedt et al. (1986) three types of A (A1, A2, A3) in the bull, Bos taurus. This large array of spermatogonia notwithstanding, the type A0, regarded as the stem cell (Clermont and BustosObregon 1968), but disputed by Courot et al. (1970), is further subdivided into types Asingle, Apaired, and Aaligned in Rattus norvegicus (Huckins 1971a,b,c). Considerable difficulty has been encountered in defining spermatogonial morphology and pattern of division in Homo sapiens, other primates and in birds, possibly due to the heterogeneous cellular associations that are characteristic of the seminiferous epithelia of these species of animals. Unlike in the laboratory and farm animals, three generations of spermatogonia, viz., type A spermatogonia, made up of two divisions, dark type A (Ad) and pale type A (Ap), and type B spermatogonia, have been described by several investigators in primates (Clermont 1963; Clermont and Antar 1973; Choudhury and Steinberger 1976; Shulze 1979; Hadley and Dym 1983). The nature of spermatogonia and their renewal is even more problematic and controversial in birds, not only because of the heterogeneous cellular associations in the seminiferous epithelium of these animals, but also because the earlier studies were performed in paraffin-embedded tissues. Zlotnik (1947), Kumaran and Turner (1949) and Sharma et al. (1956) in Gallus gallus domesticus (the domestic fowl—henceforth referred to as the rooster), Clermont (1958) in the Mallard drake (Anas platyrhynchos) and Marchand (1977) in Muscovy duck (Cairina moschata) have identified a single type of spermatogonium, while Lake (1956) describes types 1 and 2 spermatogonia in the rooster. Gupta (1955) describes two types of spermatogonia (primary and secondary) in an unspecified species of duck, and Aire et al. (1980) describe

Spermatogenesis and Testicular Cycles

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two types, A and B, in Guineafowl (Numida meleagris), while Yamamoto et al. (1967) identify three types (A, Intermediate and B) in Japanese quail (Coturnix japonica). By using plastic/resin sections and electron microscopy, as well as radiolabeling and autoradiography, Lin and Jones (1992) have shown that there are four types of spermatogonia (dark type A [Ad], 2 pale type A [Ap1 and Ap2] and type B) in Japanese quail. However, according to Ortavant (1959) “in spite of superficial differences the length of life of the various spermatogonial generations is in constant relation to the cycle of the seminiferous epithelium.”

7.2.2

Spermatocytes

Primary spermatocytes are the product of the last mitotic division of type B spermatogonia, and are involved in meiotic division. As in mammals, there are several generations of primary spermatocytes in the seminiferous epithelium of birds because meiosis is an extended and prolonged process.

7.2.2.1

Primary spermatocytes

In mammals, six phases of the meiotic prophase of the primary spermatocyte are recognized: a premeiotic interphase, leptotene, zygotene, pachytene, diplotene and, as the last phase of the meiotic prophase, diakinesis (Ortavant 1959; Courot et al. 1970; Ekstedt et al. 1986). In birds, similar phases of the meiotic prophase have been recognized, generally (Clermont 1958; Marchand 1977; Lin et al. 1990). However, Lin and Jones (1990), in a recent study, describe eight different phases (preleptotene, leptotene, zygotene, pachytene, diplotene, diakinesis, metaphase and anaphase) of primary spermatocytes during the process of spermatogenesis, in Japanese quail. The metaphase, anaphase and telophase are, however, known to occur rapidly (Ortavant 1959).

7.2.2.2

Secondary spermatocytes

Secondary spermatocytes are the two products of the first meiotic division of a primary spermatocyte. The secondary spermatocyte is rarely seen in the seminiferous epithelium because it has a very short life span. The nuclei of these cells are not as large as those of mid- to late primary spermatocytes, but are larger than those of round spermatids which they produce by the second meiotic division. The secondary spermatocyte has been described in members of the Anatidae (Clermont 1958; Marchand 1977), in the rooster (Zlotnik 1947), Guineafowl (Aire et al. 1980) and Japanese quail (Yamamoto 1967; Lin et al. 1990). The nucleus of this cell exhibits a number of chromatin aggregations and a distinct nuclear membrane. Each secondary spermatocyte divides, in the last meiotic division, to produce two haploid round spermatids.

7.2.2.3

Round spermatids

These cells, containing a haploid complement of chromosomes, represent the phase of differentiation in which relatively small cells containing round

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Reproductive Biology and Phylogeny of Birds

nuclei evolve, in a complex and delicate structural transformation, into highly elongated, motile, itinerant cells, the spermatozoa. This unique process in cytodifferentiation is called spermiogenesis or spermateliosis, the former term being more commonly used.

7.3 7.3.1

SPERMIOGENESIS Spermiogenesis in Non-passerine Birds

The complex structural evolution of the spermatozoon from a round cell, with the loss of certain organelles, even as new ones are formed, has intrigued investigators. During this process, and relative to the young, round spermatid, the mature, highly elongated spermatid loses over one hundred times its volume (McIntosh and Porter 1967) or 97% (Sprando and Russel 1988), while the volume of the nucleus is reduced from 110 cubic micrometres to 2 cubic micrometres (McIntosh and Porter 1967) or by 96% (Sprando and Russel 1988). The spermatid also radically changes its shape and evolves a number of morphologically elaborate organelles, including the acrosome, the midpiece and the flagellum (Lin et al. 1997). Biochemical changes that are controlled by genes which are active only in spermatids and in the process of spermiogenesis (Oko 1995), including the elaboration of new and unique structural elements of the spermatozoon, also take place, and are, indeed, mostly responsible for the visible morphological alterations expressed variably in the spermatid. Spermiogenesis is an integral and important part of spermatogenesis in animals, the study and understanding of which phenomenon is essential for a critical evaluation of the form and function of the seminiferous epithelium, in health and disease. Spermiogenesis has been studied and reported extensively in mammals (Leblond and Clermont 1952a,b; Clermont and Leblond 1955; Courot et al. 1970; Fawcett et al. 1971; Setchell 1978; Ekstedt et al. 1986; Plöen and Courtens 1986), and to some extent, also in birds, although most of the reports are fragmentary (Zlotnik 1947; Leblond and Clermont 1952a,b; Clermont and Leblond 1955; Sotelo and Trujillo-Cenóz 1958; Nagano 1962; McIntosh and Porter 1967; Yamamoto et al. 1967; Courot et al. 1970; Fawcett et al. 1971; Mattei et al. 1972; Tingari 1973; Humphreys 1975; Okamura and Nishiyama 1976; Gunawardana and Scott 1977; Marchand et al. 1977; Yasuzumi and Yamaguchi 1977; Ekstedt et al. 1986; Plöen and Courtens 1986; Aire et al. 1980; Baccetti et al. 1980; Kondo et al. 1988; Sprando and Russel 1988; Soley 1992; Góes and Dolder 2002; Aire 2003; Jamieson and Tripepi 2006). Most of these studies are, understandably, on the rooster (e.g. Zlotnik 1947; Nagano 1962; McIntosh and Porter 1967; Tingari 1973; Okamura and Nishiyama 1976; Gunawardana and Scott 1977), and a few on other species of birds, including passerines and the Paleognathae (e.g. Sotelo and Trujillo-Cenóz 1958; Aire et al. 1980; Kondo et al. 1988; Lin and Jones 1993; Góes and Dolder 2002; Aire 2003). The advent of electron microscopy, using both thick and ultra-thin plastic sections, has greatly facilitated the

Spermatogenesis and Testicular Cycles

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study of spermiogenesis in birds whose acrosome is quite small and thin, and therefore only discernible with difficulty in paraffin sections, PAS-stained or otherwise. The study of spermatid development is more profitably based on one or both of the classical systems enunciated by Leblond and Clermont (1952a) or by Roosen-Runge and Giesel (1950) and Ortavant (1954). Leblond and Clermont’s system is based on acrosome development, sub-divided into four phases: the Golgi phase, cap phase, acrosome phase and maturation phase, while that of Roosen-Runge and Giesel (1950) rests on evolving nuclear changes during spermiogenesis. There are few reports in which the full process of spermiogenesis has been detailed in birds, rather than fragmentary, segmental studies, e.g. of the acrosome or nucleus. In this regard complete accounts of spermiogenesis at the ultrastructural level have been reported in only the rooster (Nagano 1962; Gunawardana and Scott 1977), Rhea (Phillips and Asa 1989), Japanese quail (Yamamoto et al. 1967; Lin and Jones 1993), the turkey (Meleagris gallopavo) (Aire 2003) and House sparrow (Passer domesticus) (Góes and Dolder 2002). Several reports dealing with only segments of the developing spermatid have shed considerable light on spermiogenesis in some species of birds. These will be referred to where appropriate in this review. In those studies in which the step-wise system was used, 12 steps of spermiogenesis have been reported in Japanese quail (Lin and Jones 1993) and Turkey (Aire 2003), and 6 steps in House sparrow (Góes and Dolder 2002). In this review, both systems (involving acrosomal as well as nuclear morphogenetic processes, according to Clermont and Perey 1957, and RoosenRunge and Giesel 1950, respectively) will be combined in the so-called “stepwise” changes in spermatid morphogenesis, using spermiogenesis in Turkey (Aire 2003), as a model. Specific features in spermiogenesis of oscine passerine birds will also be highlighted separately. Only relevant, important and contentious features of spermiogenesis will be described and discussed, as necessary and convenient in each step, on account of space limitation. Step 1 spermatid. The youngest spermatids that have just emerged from the second meiotic division, and along with the step 11 spermatids, line the subluminal and luminal border of the epithelium, respectively. The oval nuclei of the round spermatids contain scattered chromatin aggregations in the karyoplasm or adhering to the nuclear membrane (Fig. 7.1A). A few proacrosomal granules may be seen in the large Golgi complex during the late phase of this step. The diplosome, comprising the proximal and distal centrioles articulated at right angle to each other, lies free in the cell cytoplasm. Step 2 spermatid. The nuclear chromatin begins to de-condense and appear uniformly distributed in the nucleoplasm. A large, single acrosomal granule appears in the Golgi complex (Fig. 7.1B). The distal centriole of the diplosome makes contact with the cell cytoplasm, at which junction, an ill-defined annulus occurs. The flagellum grows from this junction.

&" Reproductive Biology and Phylogeny of Birds

Fig. 7.1 Meleagris gallopavo. A. Step 1 spermatid, along with step 11 spermatid. S, Sertoli cell; clumps of chromatin (arrowheads) in the nucleus of step 1 spermatid; Fig. 7.1 Contd. ...

Spermatogenesis and Testicular Cycles

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Step 3 spermatid. The nucleus remains spherical in shape, but chromatin decondensation has advanced into a finely granular matrix, with only a few, small clumps of chromatin present (Fig. 7.2A). The Golgi complex becomes inconspicuous, and the acrosomal granule lies very close to, or just makes contact with the nucleus, and thickening of the nuclear membrane commences at the site of contact. The diplosome nearly makes contact with the nucleus, close to the developing acrosomal and Golgi complex. Step 4 spermatid. The chromatin of the nearly spherical nucleus continues to de-condense and becomes uniformly granulofilamentous (Fig. 7.2B). The homogeneously dense acrosomal granule becomes slightly elongated and forms the acrosome which invaginates into the nucleus; the nuclear membrane becomes thickened at the contact site. The nucleus becomes eccentric within the cytoplasm, and the acrosome abuts on the adjacent Sertoli cell (Fig. 7.2 inset). Sections of microtubules, in small groups, lie close to the nuclear membrane, especially in the rostral portion of the nucleus. The diplosome attaches obliquely at an indentation of the nucleus, by its proximal centriole. The long axis of the distal centriole remains perpendicular to that of the proximal centriole. Unlike in mammals in which the acrosomal granule/vesicle contains a concentrated granule (Burgos and Fawcett 1955; Clermont and Leblond 1955; Plöen and Courtens 1986) or a clear vesicle/vacuole in Tammar wallaby (Macropus eugenii) (Lin et al. 1997), the acrosome of birds arises from the Golgi complex as a homogeneously dense membrane-bound granule (Nagano 1962; de Reviers 1971; Okamura and Nishiyama 1976; Gunawardana and Scott 1977; Lin and Jones 1993; Soley 1996; Aire 2003). Step 5 spermatid. The nucleus, now pear-shaped, contains uniform, finely granular nucleoplasm (Fig. 7.3A). The acrosome elongates further and the central part of the thickened nuclear membrane, at the contact site with the acrosome, invaginates into the nucleoplasm (Fig. 7.3, inset). There is an increased amount of smooth endoplasmic reticulum (SER), sparsely granular endoplasmic reticulum (SGER), as well as lysosomes in the cytoplasm. The invaginated precursor of the endonuclear canal has been reported in the rooster, the Budgerigar (Melopsittacus undulatus), Turkey (Melaeagris gallopavo), Japanese quail and Ostrich (Struthio camelus) (Nagano 1962; Fig. 7.1 Contd. ...

a large Golgi complex (G) and mitochondrial aggregates (M) around it; numerous profiles of SER and a few small lysosomes (L) occur in the cell. Inset: the centriolar complex (C) in the cytoplasm. B. Step 2 spermatid nuclear chromatin is decondensing. A large proacrosomal granule (A) is leaving the Golgi complex (G). Some microtubules (arrowheads) are not arranged in any specific manner in the cell. Inset: the diplosome (D) and the fibrous sheath (F) of the developing tail. Bar: A = 2 µm, B = 1 µm. All figures on spermiogenesis (Figs. 7.1 to 7.6) are those of the turkey, Meleagris gallopavo, and are taken from Aire, T. A. 2003 British Poultry Science 44: 674-682, with the kind permission of British Poultry Science Ltd.

&$ Reproductive Biology and Phylogeny of Birds

Fig. 7.2 Meleagris gallopavo. A. Step 3 spermatid nucleus (N) de-condenses further, and the diplosome (D) associated with lamellae of the Golgi complex (G) lies close to the nuclear membrane. Step 12 spermatid (arrowhead) occurs along Fig. 7.2 Contd. ...

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Humphreys 1975a; Gunawardana and Scott 1977; Baccetti et al. 1980; Saita et al. 1980; Soley 1996; Aire 2003). An osmiophilic content of this invagination is the precursor of the perforatorium. However, Lin and Jones (1993) have observed that the precursor of the perforatorium, in Japanese quail, is an intranuclear granule. Step 6 spermatid. The spermatid nucleus is finely granular, elongated and slightly wavy in profile (Fig. 7.3B). The elongating, dense, acrosome occupies only the central one-third of the rostral surface of the nucleus, and its rostral tip abuts on an adjacent Sertoli cell. Cross-sections of profiles of microtubules of the circular manchette (CM) appear sporadically along the length of the nucleus, in no regular pattern. A CM is absent in some orders (see Chapter 8). Mitochondria migrate to the caudal part of the cytoplasm, and are still round or oval in shape, and not organized in any special fashion. The endonuclear canal is well formed and contains the developing perforatorium (Fig. 7.3B inset). Although the step 6 spermatid nucleus is elongated and slightly wavy in profile in the turkey (Aire 2003) as in the rooster (Gunawardana and Scott 1977) and Japanese quail (Lin and Jones 1993), that of Guineafowl (Aire et al. 1980), Ostrich (Soley 1997) and Crested tinamou (Eudromia elegans elegans), Tinamou (Asa et al. 1986) appears ‘spiral’ or irregular in shape due to differential subnucleolemmal chromatin condensation, and concomitant constriction of the nucleus. Light microscopical observations in the rooster (Zlotnik 1947) and in certain members of the Anseridae (Gupta 1955; Sharma et al. 1956) indicate that the spermatid nucleus, during the elongating phase, is coiled within the cell cytoplasm, apparently to ensure that the elongating nucleus is accommodated in the fixed volume of cytoplasm. This developmental feature needs clarification. The development of the circular manchette (CM) seems to follow a similar pattern in all non-passerine birds investigated (McIntosh and Porter 1967; Okamura and Nishiyama 1976; Gunawardana and Scott 1977; Xia et al. 1986; in the rooster; Humphreys 1975a, in Budgerigar, Fawcett et al. 1971 in a Columba sp.; Phillips and Asa 1989 in the rhea (Rhea americana albisceus); Lin and Jones 1993 in Japanese quail (Coturnix japonica) and Aire 2003 in the turkey), but it is not developed to the same degree in all non-passerine birds.

Fig. 7.2 Contd. ...

with step 3. Inset: The acrosomal granule (A) makes contact with the nuclear envelope which is thickened at the contact site (open arrow). B. Step 4 spermatid. The chromatin has become finely granulofilamentous in appearance. The acrosomal granule (A) invaginates further into the nucleus (N) at a thickened part of the nuclear envelope (arrow). The diplosome (D) attaches to the nucleus at the implantation fossa of the nucleus. Inset: The acrosomal granule lies close to the cell membrane. S, Sertoli cell; arrowhead, microtubules running circularly around the nucleus. Bars: A = 1 µm, Inset = 2 µm; B = 1 µm, Inset = 2 µm.

&& Reproductive Biology and Phylogeny of Birds

Fig. 7.3 Meleagris gallopavo. A. Step 5 spermatid is pear-shaped. Nuclear chromatin is uniformly and finely granulofilamentous. Acrosomal granule (A) Fig. 7.3 Contd. ...

Spermatogenesis and Testicular Cycles

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Whereas the CM is substantially well developed in Ostrich (Soley 1997), it is poorly developed in Columba sp. (Fawcett et al. 1971; Mattei et al. 1972) and the Cuckoo (Crotophaga ani) (Saita et al. 1982) but does not develop in the Swift (Apus apus) and the Nightjar (Caprimulgus europaeus) which have only a longitudinal manchette (Tripepi et al. 1991; Jamieson and Tripepi 2006; see Chapter 8). The function or role of the CM in spermiogenesis is controversial and not clearly understood. McIntosh and Porter (1967), Okamura and Nishiyama (1976), Gunawardana and Scott (1977), Lin and Jones (1993) and Soley (1997) are of the opinion that the manchette has an important function in nuclear shaping, but Fawcett et al. (1971), Asa and Phillips (1988), Phillips (1970, 1974, 1976) are emphatic that the CM plays no role in nuclear shaping in the pigeon spermatid. Myles and Hepler (1982) exhort that the role of the manchette in nuclear morphogenesis, acting singly or in combination with other cell components, must await precise determination. Step 7 spermatid. The elongating acrosome is as wide as the spermatid nucleus (Fig. 7.4A). The nucleus is slimmer than in step 6, slightly curved, and its tapering rostral end projects into the subacrosomal concavity. The fine, granular, nuclear chromatin condenses to become granulofilamentous, and deeply-stained. The cell cytoplasm extends caudally from the distal border of the acrosome, which now projects into a deep crypt of an adjacent Sertoli cell cytoplasm. The CM is well established as a layer of microtubules that aligns closely with the external surface of the elongated nucleus, extending distally from the caudal border of the acrosome to the region of the rostral end of the distal centriole. The spermatid tail is well formed (Fig. 7.4B). Mitochondria begin to elongate, and display longitudinal cristae, as they continue to migrate into the cytoplasm, much of which is displaced caudally. The development of the flagellum of the spermatozoon is generally similar in mammals (Fawcett and Phillips 1969; Yasuzumi et al. 1972; Irons and Clermont 1982a,b; Clermont et al. 1990) and non-passerine birds (Nagano 1962; McIntosh and Porter 1967; Mattei et al. 1972; Tingari 1973; Okamura and Nishiyama 1976; Maretta 1977; Gunawardana and Scott 1977; Phillips and Asa 1989; Soley 1994; Aire 2003). In all the birds, the centriolar pair, lying at right angle to each other, is closely associated with the Golgi apparatus, and, initially, lies mid-way between the cell membrane and the nucleus. The Fig. 7.3 Contd. ...

elongates and is laterally compressed. Arrowhead, obliquely sectioned endonuclear cavity containing the perforatorium; Inset: Arrowheads, thickened, granular nuclear membrane invaginates at its centre; arrow, dense granule in the developing perforatorium. A, acrosome. B. Step 6 spermatid has an elongated nucleus (N); arrowheads, microtubules of developing circular manchette; A, acrosome lies rostral to most of cell cytoplasm; Arrows, ‘shoulder’ of the nucleus. Inset: the endonuclear canal contains the developing perforatorium. Bars: A = 1 µm, Inset: 2 µm; B (including inset) = 1 µm.

' Reproductive Biology and Phylogeny of Birds

Fig. 7.4 Meleagris gallopavo. A. Step 7 spermatid shows granular nuclear chromatin beginning to condense and become more electron-lucent than in step 6. The circular manchette (arrowhead) is established, extending from the nuclear Fig. 7.4 Contd. ...

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diplosome thereafter migrates gradually towards the nucleus, to which it eventually attaches, usually obliquely, at the portion of the nucleus destined to become the caudal pole. The avian proximal and distal centrioles differ in length, with the distal one being slightly to much longer than the proximal centriole (see Chapter 8), unlike in mammals in which both centrioles, if persistent, are of similar length. Also, the distal centriole persists without much modification in birds, but disintegrates during flagellar formation in mammals (Fawcett and Phillips 1969; Phillips 1974). Thus, in non-passerine birds, the distal centriole forms the foundation upon which the midpiece of the spermatozoon is built. It produces the central pair of the axonemal microtubules at the base of this centriole, determines the length of the midpiece and, according to Phillips and Asa (1989), allows the spermatozoon to form a midpiece without moving the annulus, relative to the distal centriole, as occurs in mammals (Phillips 1974). The axonemal microtubules, generally, extend from the proximal centriole caudad, into the flagellum, in mammals (Fawcett and Phillips 1969; Gordon 1972; Fawcett 1975). Although in all birds studied, the proximal and distal centrioles lie perpendicular to each other, in both the spermatid (e.g. Nagano 1962; Marchand 1977; Phillips and Asa 1989; Lin and Jones 1993; Soley 1994; Aire 2003) and mature spermatozoon (Tingari 1973; Phillips and Asa 1989; Thurston et al. 1982; Thurston and Hess 1987), those of Guineafowl initially lie at right angle to each other, but the angle between them gradually becomes obtuse as the centriolar complex inserts in a deep vault in the nucleus of the spermatid (Aire and Soley 2003). Both centrioles subsequently become in-line aligned so that the junction between them is hardly discernible in longitudinal sections. Guineafowl therefore does not lack the proximal centriole, as conjectured by Thurston et al. (1982). However, this type of centriolar alignment is uncommon, and found mainly in invertebrate organisms (Afzelius 1979). The attachment of the tail to the nucleus is simpler in birds than in mammals. In the latter, an electron-dense basal plate or capitulum inserts between the proximal centriole and the implantation fossa. In birds, there is Fig. 7.4 Contd. ...

‘hump’ (arrows) of the cell cytoplasm to the centriolar region, caudally. A, the acrosome is as wide as the nucleus. B. Lysosomes (L) are numerous; P, proximal centriole; D, distal centriole; F, fibrous sheath; arrow, annulus. C. Step 8 spermatid. Inset: the acrosome elongates and is pointed rostrally; C, collar of SER of the Sertoli cell surrounds the acrosome. In the main micrograph, the white arrow shows the dense coarse, round or rod-shaped granules; thick arrow, chromatoid body. D. Circular (arrowheads) and longitudinal (arrows) manchette occur concurrently at a later stage of this step of spermiogenesis; The central part of the nucleus is largely devoid of these dense granules. Bars: A, B = 1 µm, C, D = 2 µm, Inset of C = 1 µm.

'

Reproductive Biology and Phylogeny of Birds

no capitulum, although in the ostrich a thin layer of dense material, in place of the capitulum, occurs (Soley 1994). The cross-striated connecting piece found in mammals is also absent in birds, although a structure akin to it develops in early spermatids; this merges to form the non-segmented columns found in mature spermatozoa in Ostrich (Soley 1994). Step 8 spermatid. The diameter of the nucleus further reduces. The nuclear chromatin begins to condense into coarse, round or rod-shaped granules that are immersed in the granulofilamentous matrix of the karyoplasm (Fig. 7.4C). A chromatoid body is seen in the proximal region of the cell. Other dense, amorphous aggregations, of unknown function, may be seen along the entire length of the centriolar complex. A significant, rarely observed feature of the spermatid during this step is the concurrent occurrence of both the CM and longitudinal manchette (LM), with profiles of the latter being lateral to the former, which appears to be patchy in distribution, at this stage (Fig. 7.4D). The conversion of the CM to the LM is very transient. The chromatoid body has been described in male germ cells in several mammals (Sud 1961; Fawcett et al. 1970; Susi and Clermont 1970; Fawcett 1971; Bawa 1975; Söderstrom 1978; Thorne-Tjömsland et al. 1988), but there is still some controversy concerning its origin, precise structure and function. This structure “is an irregularly shaped, dense mass of fine fibrillar material generally found near the acrosomal vesicle and Golgi complex of early spermatids” (Fawcett 1971). It is typically reticular in structure in sections. The origin and structure of the chromatoid body has been studied in mammals (Sud 1961; Eddy 1970; Fawcett 1971; Bawa 1975). It is apparently formed by an aggregation of filamentous material that is abundant in spermatocytes. There are very few reports, indeed, on the chromatoid body in birds. Gupta (1955) has described the presence of this structure in unfixed seminiferous tubules of the drake. The chromatoid body has been demonstrated in spermatids of Rhea by Phillips and Asa (1989), while Soley (1994) and Aire (2003) have described it in the spermatids of the ostrich and the turkey, respectively. Guineafowl spermatids also possess chromatoid bodies (Soley and Aire unpublished). According to Fawcett (1971) the chromatoid body migrates caudally, disperses in the process, and ultimately disappears, but its fate in the avian spermatid is not known. The role of this structure in spermiogenesis is not clearly understood. It is considered to play roles in the development of the connecting piece in mammals (Fawcett 1971) and in the transportation of ribonucleoproteins to the structures in the neck region of the developing spermatid (Paniagua et al. 1986) or maturation of the nuclear chromatin of the spermatid (Sud 1961), but the latter view has been discounted by Eddy (1970) who failed to demonstrate RNA presence in the chromatoid body, nor its involvement in the elaboration of the connecting piece, in various laboratory animals. The presence of a dual set of manchette microtubules has now been established in mammals and non-passerine birds. The CM appears to be

Spermatogenesis and Testicular Cycles

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poorly developed in a pigeon (Columba sp.) (Fawcett et al. 1971; Mattei et al. 1972) and cuckoo (Crotophaga ani) (Saita et al. 1982), but very well developed in the ostrich, in which it consists of a double set of tubules that are linked together (Soley 1997). The transition to the LM in the elongated spermatids of animals has also provoked some controversy. There appears to be a transition, of quite brief duration, between the disappearance or reorganization of the CM and the establishment of the LM. Both sets of manchette microtubules occur concurrently, for apparently a fleeting period only, in the African collared dove (Streptopelia roseogrisea) (Mattei et al. 1972), Rhea (Phillips and Asa 1989), Ostrich (Soley 1997) and Turkey (Aire 2003), during which period the microtubules of the CM are probably rearranged to become the LM (Phillips and Asa 1989). Although Okamura and Nishiyama (1976) consider the transition to be abrupt in the rooster, an illustration by Gunawardana and Scott (1977), in the same species of bird, indicates a concurrent presence of both sets of manchette microtubules, as has been observed in Rhea (Phillips and Asa 1989) and Turkey (Aire 2003) (see further examples in Chapter 8). Step 9 spermatid. The LM is fully established and extends, beyond the midpiece and annulus, into the trailing cell cytoplasm, caudally (Fig. 7.5A). Step 10 spermatid. The nucleus elongates further in a gentle curve, and with a reduced diameter compared to the preceding spermatids (Fig. 7.5B,C,D). The acrosome is well formed and houses the perforatorium in its subacrosomal space. The coarse nuclear chromatin granules become more electron-dense and more compactly packed together than in spermatid step 9. Mitochondria continue to elongate and increase the density of their matrix. Step 11 spermatid. The nuclear chromatin granules become large, highly electron-dense and compact (Fig. 7.5E,F,G). The spermatid is cylindrical, and maintains the gentle curvature. The acrosome is lanceolate and accommodates a perforatorium that extends from the depth of the endonuclear canal to close to the apex of the acrosome. The tapering end of the nucleus remains protuberant into the subacrosomal space. The LM is still present and is surrounded distally by elongated mitochondria in the caudal part of the trailing cytoplasm. During the latter part of this step, the LM begins to break up patchily. Glycogen aggregations may be seen in the cytoplasm (Fig. 7.5G). The acrosome and the perforatorium have, together, been regarded as the acrosome complex (Baccetti 1979). The perforatorium is a fibrous structure consisting of parallel bundles of filaments (Baccetti 1979) composed of actin (Campanela et al. 1979; Baccetti et al. 1980). The structure of this complex in birds has been described by several authors, in the rooster (Lake et al. 1968; Bakst and Howarth 1975; Thurston and Hess 1987), the turkey (Thurston et al. 1982), Guineafowl (Thurston et al. 1982; Thurston and Hess 1987), Mallard drake (Maretta 1975), Rhea (Phillips and Asa 1989), Crested tinamou (Eudromia elegans elegans) (Asa et al. 1986) and Ostrich (Soley 1993; Baccetti et al. 1991) (see also Chapter 8). However, there are only a few reports on the

'" Reproductive Biology and Phylogeny of Birds

Fig. 7.5 Meleagris gallopavo. A. Step 9 spermatids. The LM is fully established, with the disappearance of the CM. Straight arrows, dense amorphous material; A, annulus; F, fibrous sheath. Step 10 spermatids (B, C and D): B. and C, the Fig. 7.5 Contd. ...

Spermatogenesis and Testicular Cycles

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development of the complex in birds, and these are specifically in the rooster (Nagano 1962; Tingari 1973; Okamura and Nishiyama 1976; Gunawardana and Scott 1977; Baccetti et al. 1980), Budgerigar (Humphreys 1975a), Japanese quail (Saita et al. 1980; Lin and Jones 1993), Guineafowl (Aire 2003) and Ostrich (Soley 1996). In all reports, a similar process of development exists in non-passerine birds. From about Step 6 of spermiogenesis (in Turkey, Aire 2003), the crater in the nucleus formed by and lodging the acrosomal granule, flattens out, and the rostral end of the nucleus becomes convex, once again. Concurrently, the caudal part of the elongating acrosome, at the interface with the nucleus, begins to invaginate, thus, beginning the formation of the subacrosomal space. As this space deepens, the developing rostral end of the perforatorium is pulled along or pushes into the space. The perforatorium is thought to assist in elongating and supporting the acrosome, during its development (Baccetti et al. 1980) but it is known to project after disruption of the acrosome vesicle in the acrosome reaction of some vertebrates, e.g. Lamprey and Sturgeon (see Jamieson 1991). The fully developed perforatorium is embedded in the endonuclear canal and projects into the deep and ample subacrosomal space, in most birds, extending to just beneath the rostral end of the acrosome (Nagano 1962; Tingari 1973; Gunawardana and Scott 1977; Baccetti et al. 1980; Soley 1996; Aire 2003). The perforatorium is absent in passerine birds, vide infra, and in some non-passerines (see Chapter 8). Whereas the rostral tip of the nucleus tapers slightly and projects for a short distance into the subacrosomal space, thus forming the intra-acrosomal portion of the nucleus, in most birds (Lake et al. 1968; Bakst and Howarth 1975; Maretta 1975; Thurston and Hess 1987), the intra-acrosomal portion of the nucleus is extremely long and occupies almost all of the subacrosomal space, bearing the equally long perforatorium in its extensive endonuclear canal in Struthio camelus (Soley 1996). It is noteworthy that the rostral tip of the nucleus of Budgerigar (Humphreys 1975; Jamieson et al. 1995), white-naped Crane (Phillips et al. 1987), cockatiel, and peach-faced lovebird (Jamieson et al. 1995) does not project into the subacrosomal space, as in other birds, but makes direct, en face contact with the caudal rim of the acrosome. In these birds, therefore, the sperm nucleus has no intra-acrosomal portion because the Fig. 7.5 Contd. ...

acrosome is highly elongated and compressed laterally, D. the central part of the nucleus is devoid of chromatin granules; arrowheads, LM; arrows, numerous multivesicular bodies; L, lysosomes; M, mitochondria with longitudinal cristae. Step 11 spermatids (E, F and G); E, chromatin granules are dense and compacted in the nucleus, F, the acrosome (A) is lanceolate, P, perforatorium surrounded by fuzzy material in the subacrosomal space; arrows, LM. G, transverse section of spermatid showing the principal piece of the spermatid; M, mitochondria surrounding the LM (arrow); G, glycogen granules. Bars for all figures = 1 µm.

'$ Reproductive Biology and Phylogeny of Birds

Fig. 7.6 Meleagris gallopavo. Step 12 spermatids. A. The mitochondria (M) form a sheath around the midpiece, and the spermatid continues to withdraw from its Fig. 7.6 Contd. ...

Spermatogenesis and Testicular Cycles

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acrosome does not overlap it. It is also interesting to note that the nucleus of the mature avian spermatid and spermatozoon consists of compact chromatin granules, and not of condensed, homogeneous chromatin, found in insect and mammalian spermatozoa (Okamura and Nishiyama 1976; Phillips and Asa 1989; Thurston et al. 1982), though it approaches homogeneity in Apus apus (Jamieson and Tripepi 2006). Step 12 spermatid. The LM disappears, allowing the elongated mitochondria to form a helical sheath closely around the midpiece (Fig. 7.6A). The evolved mature spermatid moves away from most of the redundant cell cytoplasm which still contains glycogen accumulations (Fig. 7.6B), and profiles of the endoplasmic reticulum as well as multivesicular bodies. The mature spermatid attains the luminal surface of the seminiferous tubule, and is held in place by only slips of Sertoli cell cytoplasm. It is ready for spermiation (Fig. 7.6C). The released spermatozoon possesses no cytoplasmic droplet, as formed in mammals. The dissolution or disassembly of the LM during the early phase of Step 12 of spermiogenesis in Turkey (Aire 2003) appears to make way for the elongated, dense mitochondria, that have aggregated to surround, sometimes, at least, helically, the proximal axoneme as the mitochondrial sheath. A similar observation has been made in mammals (Courot et al. 1970; Phillips 1974), Rhea (Phillips and Asa 1989) and Ostrich (Soley 1994). A departure from this developmental process is in Japanese quail, in which the mitochondrial sheath has already begun to form in Step 10 spermatid, even when the CM is still in place, and is complete before the LM disappears in the early phase of Step 12 spermatids (Lin and Jones 1993). The mitochondrial sheath is also in place when the LM is still well developed, in the absence of a CM, in Caprimulgus europaeus (see Chapter 8). It is not known whether the mitochondria move through the curtain of manchette or migrate into this curtain through its distal end, in this species.

7.3.2

Spermiogenesis in Passerine Birds

The passerine birds constitute a large group among birds, and although they are more derived than, for example, members of the Galliformes, Columbiformes, Anseriformes or Struthioniformes and other ratites, the study of spermatogenesis, and its related phenomena, in this group of birds has lagged behind those of non-passerine birds. They have been the most common birds, from the time man started studying this class of animals. There are only Fig. 7.6 Contd. ...

own redundant, electron-dense cytoplasm (C); N, nucleus of spermatid. B. Transverse sections of step 12 spermatids: the LM disappears and mitochondria (M) then align themselves around the midpiece; glycogen granules (Gl) are still present in the cytoplasm. C. Late phase of step 12, showing a well established mitochondrial sheath. Slips of Sertoli cell cytoplasm (arrows) hold on to the spermatid tenuously, as the spermatid is ready for spermiation. Bars: A = 4 µm; B = 2 µm and C = 1 µm.

'& Reproductive Biology and Phylogeny of Birds

Fig. 7.7 Diagram of the 6 steps of spermiogenesis of the domestic sparrow, Passer domesticus. Two views (A: dorso-ventral view and B: side-view), show Fig. 7.7 Contd. ...

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a few reports on the male gamete and its morphogenesis in passerine birds, such as House sparrow (Passer domesticus) (Yasuzumi 1956; Sotello and Trujillo-Cenóz 1958; Góes and Dolder 2002), Lovebird or Bengalese finch (Lonchura striata var. domestica) (Fawcett et al. 1971; Yasuzumi and Sugioka 1971; Kondo et al. 1988) and Zebra finch (Taeniopygia (=Poephila) guttata) (Nicander 1970). For accounts of mature spermatozoa, see Chapter 8. During the early and late acrosome phase in oscine birds, e.g. House sparrow and Lovebird (Bengalese) finch (Lonchura), the acrosomal vesicle, in Step 2 spermatids, contains a matrix of low and medium density and lodges in a nuclear cavity or depression (Fig. 7.7), without forming a cap (Góes and Dolder 2002), as in non-passerines. As the acrosome elongates, the matrix differentiates into an outer low-density zone as well as a central higher density zone, and assumes a zigzag course, with sharp angulations of its membrane projecting alternately on both sides (Fawcett et al. 1971). The perforatorium is absent. In early spermatids, the nuclear chromatin is loosely arranged (Góes and Dolder 2002) or condenses particularly into fine granules (Kondo et al. 1988). Further condensation of the nuclear chromatin forms denser coarse granules. There is no agreement on the appearance of microtubules around the nucleus of Lonchura spermatids (Fawcett et al. 1971; Kondo et al. 1988). However, microtubules appear very early around the spherical nucleus in a cluster, extending from the rostral to the post-nuclear region (Kondo et al. 1988), but they are only evident in intermediate and late stages of differentiation in Step 3 spermatids (Góes and Dolder 2002) when the nucleus has already assumed a helical form (Nicander 1970; Fawcett et al. 1971; Yasuzumi and Sugioka 1971; Góes and Dolder 2002). The bundle of microtubules describes a helical course on the outer surface of the gyres of the nucleus and the developing flagellum (Fig. 7.7). The mitochondria, located predominantly in the postnuclear region, begin to fuse together, forming a long strand alongside the axoneme. The helical microtubular bundle around the nucleus does not extend rostrally beyond the basal turn (in the region of the base of the acrosome) of the helix, indicating that the spiral shape of the elongated acrosome may not be influenced by the microtubules (Fawcett et al. 1971) but rather by an intrinsic mechanism yet to be understood. Following the establishment of the microtubular helix, during the mid-maturation phase, the long strand of mitochondria winds round the axoneme helically, such that it lies between the axoneme and the microtubule bundle (Fawcett et al. 1971; Kondo et al. 1988; Góes and Dolder 2002). The Fig. 7.7 Contd. ...

different sectioning planes, have been diagrammed for steps 2, 3 and 4, so as to include all organelles. A, acrosome; ax, axoneme; ca, centriolar adjunct; df, dense fibers; ER, endoplasmic reticulum; G, Golgi complex; ig, pro-acrosomal internal granule; ms, mitochondrial sheath; mt, microtubules; Pa, pro-acrosome; sr, spiraling ridges of the acrosome. From Góes, R. M. and Dolder, H. 2002 Tissue and Cell 34: 273-282. Reproduced with permission of Elsevier Science Ltd.

! Reproductive Biology and Phylogeny of Birds bundle of microtubules, the homologue of the manchette in mammals and non-passerine birds (Fawcett et al. 1971), makes shallow indentations on the outer surfaces or ridges of the helical nucleus. During the late maturation phase of the spermatid, acrosomal condensation and elongation is completed, and the cell contains 2.5 gyres in Lonchura striata (Kondo et al. 1988). The bundles of microtubules are arranged in rows that alternate with cisternae of smooth endoplasmic reticulum (Fawcett et al. 1971; Kondo et al. 1988; see also Chapter 8). The role of the latter is unknown. It is generally agreed that the microtubular bundles are transient, being lost before spermiation of mature spermatids in Lovebird (Bengalese) finch (L. striata var. domestica) (Kondo et al. 1988; Fawcett et al. 1971) or shed as the spermatozoa pass through the ductus deferens in Zebra finch (Nicander 1970) and House sparrow (Passer domesticus) (Góes and Dolder 2002), although Yasuzumi and Sugioka (1971) regard them to be a permanent feature involved in motility in the spermatozoa of L. striata. The development of the flagellum of the passerine bird has received even less attention than acrosomal and nuclear morphogenesis. Centriolar complex development in passerine birds is generally similar to that in mammals and non-passerines (Sotelo and Trujillo-Cenóz 1956), but whereas Góes and Dolder (2002) consider that the proximal centriole lodges in an implantation fossa of the nucleus while the distal centriole extends, caudally, to the cell membrane and forms an annulus at the contact junction, Sotelo and TrujilloCenóz (1956) state that one of the centriolar pair in House sparrow disappears, unaccounted for. Nicander (1970) also states that passerine birds possess only one modified centriole, and that both a segmented pericentriolar material as well as the nine identical coarse peripheral tail fibers anchor the neck to the base of the head. A similar striated structure, akin to the mammalian connecting piece, has been shown in micrographs of Lonchura striata spermatozoa, published by Fawcett et al. (1971). For further details of mature sperm, see Chapter 8.

7.4 SPERMIATION Spermiation has been reported in only two birds, the rooster (Sprando and Russel 1988) and Japanese quail (Lin and Jones 1993). The process is similar in both species of birds, belonging to the Galliformes. During the process of spermiation, the spermatid cytoplasm tends to condense and become more electron-dense relative to the cytoplasm of other germ cells, as well as the Sertoli cells (Sprando and Russel 1988). Residual bodies, light-staining in birds, but highly condensed in mammals, form late in spermiogenesis, near the time of sperm release (Sprando and Russel 1988). They are phagocytised by Sertoli cells (Sprando and Russel 1988; Lin and Jones 1993). The tubulobulbar complex has not been observed in birds. Further studies on spermiation, involving other orders of birds are necessary for a complete picture of this phenomenon in this large class of animals. Unlike in mammals

Spermatogenesis and Testicular Cycles

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(Fawcett and Phillips 1969), there are no reports of cytoplasmic droplets in newly released spermatozoa of birds (Lake 1981; Sprando and Russel 1988) except in those of Ostrich (Aire and Soley 2000). This indicates that the loss of the excess cytoplasm in the region of the head of the spermatid in the ostrich is probably similar to that described for the Galliformes (Sprando and Russel 1980; Lin and Jones 1993), but unlike in mammals where it is from the junction between the head and neck of the spermatid. The study of the formation of the cytoplasmic droplet in the spermatozoon, and its loss in the post-testicular spermatozoon of Ostrich is in progress in our laboratory.

7.5

KINETICS OF SPERMATOGENESIS IN BIRDS

In most mammals, cross-sections of seminiferous tubules contain varying generations of germ cells arranged in a definite, successive, repeatable order, from the basement membrane to the lumen of the tubule (von Ebner 1871; Regaud 1901). It has been established that each generation of germ cells, resulting from stem cell divisions, are all linked by cytoplasm bridges (Dym and Fawcett 1971) to form a mass of syncytial cells that develop in synchrony. According to Perey et al. (1961) each generation of germ cells is at exactly the same step of development, and normally associated with other generations in a manner that is predictable and constant for the species. Thus, “groups of spermatids at a given step of development are always associated with the groups of spermatocytes and spermatogonia” (Perey et al. 1961), as well as the same group of Sertoli cells which provide support, nutrition and regulatory mechanisms for their development (Dym and Fawcett 1970; Morris et al. 1987a; Jégou 1991; Skinner et al. 1991). This constant grouping of the same generations of cells, arising from the same spermatogonium in a segment of the seminiferous epithelium is known as a cellular association. Leblond and Clermont (1952a,b) identified 14 consistent cellular associations in the seminiferous epithelium of Rattus norvegicus. The identification or classification of cellular associations has been facilitated by the use of specific morphological changes in developing spermatids. Cellular associations occur in an orderly synchronized, successive series in any given area of the seminiferous tubule, such that, in Rattus norvegicus, the 14 associations, arranged in a succession, numbered I to XIV, constitute a cycle of the seminiferous epithelium. Each cellular association occurs in a specific period in the cycle, and is known as a stage of the cycle. In most mammals, each crosssection of the seminiferous tubule consists of a single stage of the cycle of the seminiferous epithelium. But in birds, as in primates (Roosen-Runge 1952; Clermont 1963; Chowdhury and Marshall 1980; Dietrich et al. 1986; Johnson et al. 1981), cross-sections of the seminiferous tubule display several different (heterogeneous) cellular associations or stages of the seminiferous epithelium, e.g. there are up to 6 cellular associations in the rooster (Courot et al. 1970), 10 in Japanese quail (Lin and Jones 1990) and 3 in primates (Schulze 1982; Schulze and Rehder 1984; Dietrich et al. 1986; Schulze et al. 1986).

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Reproductive Biology and Phylogeny of Birds

Although Gunawardana (1976) and Gunawardana and Scott (1977) used the term “stage” in their studies of the the rooster, they were, in fact, referring to steps of spermiogenesis. Certain constraints, such as heterogeneous, as well as atypical cellular associations occupying small areas of the seminiferous tubule (Table 7.1), unlike in non-primate mammals (Clermont 1963; Dietrich et al. 1986; Aire et al. 1980), the lack of sensitivity of the PAS method for classifying steps of spermiogenesis in birds and the use of paraffin-embedded material, have confounded the study of various aspects of spermatogenesis in birds (Clermont 1958; Yamamoto et al. 1967; de Reviers 1971; Aire et al. 1980). In paraffin-embedded tissues, eight stages of the cycle of the seminiferous epithelium were identified in each of Japanese quail (Yamamoto et al. 1967), Mallard (Clermont 1958) and Guineafowl (Aire et al. 1980). However, the introduction and use of electron microscopic facilities and processes have made it possible for the identification of subtle aspects of avian spermatogenesis, thus significantly enriching our knowledge (Lin and Jones 1990; Lin et al. 1990; Lin and Jones 1992). It is noteworthy that these studies are the most complete and up-to-date evaluations of the seminiferous epithelium in any avian species. This review will therefore draw heavily on these reports. Lin et al. (1990) have been able to classify the seminiferous epithelium of Coturnix into 10 stages (I to X), and 12 steps of spermiogenesis, using acrosomal development (Leblond and Clermont 1952b) and nuclear morphogenesis of the spermatids (Roosen-Runge and Giesel 1950; Ortavant 1954) as criteria, in plastic sections. Subsequently, and by means of autoradiography and radiolabeling, Lin and Jones (1992) have also determined the process of stem cell renewal, and spermatogonial proliferation and differentiation into spermatocytes (Fig. 7.8). Figure 7.9 shows the cycle of the seminiferous epithelium of Japanese quail, which includes spermatogonial types, as well as other generations of germ cells and their occurrences in the various, successive stages of the cycle of the seminiferous epithelium. Furthermore, both the frequency and duration of the 10 stages of Table 7.1 Area of the wall of a seminiferous tubule occupied by a cellular association of the seminiferous epithelium and by a Sertoli cell, and the number of Sertoli cells in a cellular association Cellular associations Quail a

No. measured

Sertoli cells 2

Area (mm )

No. measured

Area (mm 2)

No. per association

208 37 12760 ± 978 23 1625 ± 732 7.9 ± 1.9 24 21291 ± 1812 21 1405 ± 455 15.2 ± 4.1 269a 31 19655 ± 1737 24 1133 ± 88 17.3 ± 2.9 278a Meanb 17902 ± 2614 1388 ± 142 13.5 ± 2.8 Values are mean ± s.e.m. for 3 birds, with the s.e.m. calculated from the variance between astages within a seminiferous tubule and banimals. From Lin, M. and Jones, R.C. 1990. Journal of Reproduction and Fertility 90: 361-367, Table 1. © Society for Reproduction and Fertility (1990). Reproduced by permission.

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the cycle of the seminiferous epithelium (Tables 7.2 and 7.3) have been determined for Japanese quail (Lin and Jones 1990). The duration of one cycle of the seminiferous epithelium in Japanese quail is estimated to be 2.69 ± 0.08 days. Compared to mammals, spermatogenesis in birds, which are generally very promiscuous, is a very rapid process because they invest heavily in rapid sperm production, since, unlike mammals, they do not store them for any appreciable length of time. For example, whereas the testis weight relative to body weight in Japanese quail is between 2.26 and 3.3% (Clulow and Jones, 1982; Aire 2005), that in R. norvegicus is 0.67% (Clulow and Jones 1982). It is generally agreed that seminiferous tubular diameter, epithelial height, testicular weight and spermatogenesis are positively related. Table 7.2 Relative frequency (mean ± s.d.) and duration of the stages of the cycle of the seminiferous epithelium in the Japanese quail Stage I

II

III

IV

V

VI

VII

VIII

IX

X

Frequency (%)

11.9 14.8 24.1 10.3 8.2 6.4 9.4 5.5 3.8 5.4 ±3.1 ±5.1 ±3.9 ±3.4 ±1.4 ±0.5 ±5.5 ±2.4 ±1.5 ±0.8 Duration (h)* 7.7 9.5 15.5 6.6 5.3 4.1 6.1 3.6 2.5 3.5 *Based on the estimate of 64.4 h for one cycle. From Lin, M., Jones, R. C. and Blackshaw, A. W. 1990. Journal of Reproduction and Fertility 88: 481-490., Table 1. © Society for Reproduction and Fertility (1990). Reproduced by permission.

Table 7.3

Estimates of the duration of the seminiferous epithelium in the Japanese quail Most advanced labeled cell

Time (h) after injection of [ 3H] thymidine

Cycle stage*

No. of cycles traversed

Cell type 2 26 50 74 98

Leptotene primary spermatocyte Pachytene primary spermatocyte Pachytene primary spermatocyte Pachytene primary spermatocyte Secondary spermatocyte

Duration of one cycle Days

Hours

-

-

2, IV

-

2, VII

0.345

2.90

69.6

3, II

0.759

2.63

63.1

3, V

1.187

2.53

60.7

3, X 1.493 2.68 64.3 Mean± s.e. 2.69±0.08 64.4±1.88 *No difference found among the animals within each group. From Lin, M., Jones, R. C. and Blackshaw, A. W. 1990. Journal of Reproduction and Fertility 88: 481-490, Table 2. © Society for Reproduction and Fertility (1990). Reproduced by permission.

!" Reproductive Biology and Phylogeny of Birds

Fig. 7.8 Diagram showing the mode of stem cell renewal and proliferation of spermatogonia in the quail. Numeric superscripts show the number of germ cells Fig. 7.8 Contd. ...

Spermatogenesis and Testicular Cycles

7.5.1

!#

Wave of the Seminiferous Epithelium

The completion of a full series of cellular associations, between two successive appearances of the same stage, along the length of the seminiferous tubule, is known as the wave of the seminiferous epithelium. Thus, for Rattus norvegicus, the wave of the seminiferous epithelium is “a series of adjacent segments, each of which includes the 14 possible types, in addition to any segment which is involved in modulation” (Perey et al. 1961). ‘Modulations’ are irregularities caused by temporary and spatially limited reversion or inversion of the numerical order in successive occurrences of stages in the cycle of the seminiferous epithelium (Perey et al. 1961; Dietrich et al. 1986). In R. norvegicus, the numbering of the stages in the wave decreases from the rete testis, into which the loop of the seminiferous tubule opens, at both ends (Perey et al. 1961). Modulation appears to be a frequent occurrence in the waves, because only 20% of the waves are unaffected, while up to 17% have more than three modulations in R. norvegicus (Perey et al. 1961). The phenomenon of modulation has not been reported in birds, apparently because of the anastomotic nature of the seminiferous tubules in this class of animals (Lin and Jones, 1990). The occurrence of heterogeneous cellular associations in the cross-section of the seminiferous tubule of birds and primates has impeded the study of the kinetics of the seminiferous epithelium in these animals. That is why only a few studies on the nature of the wave of the seminiferous epithelium in birds and primates have been reported, in Japanese quail (Lin and Jones 1990), and Homo sapiens (Schulze 1982; Schulze and Rheder 1984). Perey et al. (1961) show that in most mammals, waves are not arranged spirally in the seminiferous tubule. However, Lin and Jones (1990) have demonstrated a spiraled helical arrangement of the waves of the seminiferous epithelium in Japanese quail, as has been described for some primates (Schulze 1982; Schulze and Rehder 1984; Dietrich et al. 1986; Schulze et al. 1986). This arrangement in primates and the quail, as perhaps also in other birds, may partly explain why there are heterogeneous cellular associations in the epithelium of cross-sections of the seminiferous tubules in these animals, but not in other mammals that lack this helical pattern of wave arrangement. The crab-eating macaque, one of the primates exhibiting heterogeneous cellular associations, displays frequent modulations in the wave of the seminiferous epithelium (Dietrich et al. 1986). The branching pattern of seminiferous tubules in birds has not made it possible to determine the presence and pattern of modulations. In adult animals, there are no further migrations of primordial Fig. 7.8 Contd. ...

in each generation. See Fig. 7.9 for time of occurrence of each cell type and division during the cycle of seminiferous epithelium. From Lin, M. and Jones, R. C. 1992 Cell and Tissue Research 267: 591-601, Figure 12. Reproduced with the kind permission of Springer Science and Business Media.

!$ Reproductive Biology and Phylogeny of Birds germ cells or gonocytes (which mature to become spermatogonia) into the seminiferous tubules, neither do Sertoli cells undergo further mitotic divisions; the numbers of Sertoli cells are therefore constant in the seminiferous epithelium. The process of spermatogenesis commences with the mitotic divisions of spermatogonia in the active testis. Spermatogenesis is a continuous process in the fully stimulated and functionally active testis. The continuation and sustenance of spermatogenesis is therefore contingent upon the availability of stimulated spermatogonia whose successive mitotic divisions will produce spermatocytes. The replacement of these dividing spermatogonia has engaged the attention of reproductive biologists over the years. According to Roosen-Runge (1962) the process of spermatogenesis in mammals is cyclic, such that in any given area of the seminiferous tubule, a new generation of germ cells begins to differentiate before the last generation has completed development. These new generations initiate their development at definite and periodic intervals, in fixed relation to preceding generations. The nature and process of spermatogonial replacement and/or conversion to primary spermatocytes has been examined in several mammals, although reports are not without discrepancies, contradictions and controversy (Roosen-Runge 1951; Leblond and Clermont 1952b; Oakberg 1956a; Clermont 1958; Clermont and Leblond 1959; Hochereau 1967; Clermont 1972; Clermont and Antar 1973; Clermont and Hermo 1975; Roosen-Runge 1977; Huckins 1978; Setchell 1982). The methods of investigation have been part of the problem, but it appears certain that, in all animals, “Cycle after cycle, the primitive cells repeat the same behavior” (Roosen-Runge 1962), with regard to the pattern and nature of spermatogonial differentiation and renewal. The heterogeneous cellular associations, as well as the small areas occupied by stages of the cycle of the seminiferous epithelium might have impeded studies involving spermatogonia and their divisions in birds. Thus, there are only two main reported studies on spermatogonial divisions, differentiations and renewal in birds (Clermont 1958; Lin and Jones 1992). Clermont (1958) employed colchicine as a method of evaluation of the spermatogenic epithelium in Mallard, but the limitations of this method include the absence of adequate knowledge of the structural features of all possible types of spermatogonia. In addition, colchicine arrests mitotic division at metaphase, and cells in such a state contributed to the counts. The studies reported by Lin and Jones (1992), in order to overcome these limitations, used autoradiography and radiolabeling to investigate the process of spermatogonial differentiation and renewal, in Japanese quail. In an 8-stage cycle of the seminiferous epithelium in Mallard, Clermont (1958) observed and described three types of spermatogonia: type A (stem) spermatogonium which divides during Stage V of the cycle of seminiferous epithelium. The products of this division are two, non-identical, variablytracked cells: a new, resting type A (stem cell) spermatogonium and a differentiating type B spermatogonium. The latter undergoes mitotic division, during Stage VIII of the seminiferous epithelial cycle, to produce two type C

Spermatogenesis and Testicular Cycles

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spermatogonia, which, on division during Stage III of the cycle, continue with germ cell differentiation, into spermatids. In their study on Japanese quail, Lin and Jones (1992) have described four types of spermatogonia in the seminiferous tubules, as follows: a dark type A (Ad), 2 variants of pale type A (Ap1 and Ap2), and a type B (Figs. 7.8 and 7.9). By means of electron microscopy, the morphological features of each type of spermatogonium have been described in detail, for the first time, in an avian species (Lin and Jones 1992). The following brief review is taken, mainly, from this work. Dark type A (Ad) spermatogonia lie on the basal lamina, are elliptical in shape and stain densely (Fig. 7.10A). The eccentric nuclei are small, ovoid and dark-staining, and the nucleoplasm contains uniformly dispersed heterochromatin aggregations. The cytoplasmic organelles appear sparse, and include a moderate abundance of mitochondria located basolateral to the nucleus, and a few strands of RER. Type Ad spermatogonia are regarded as the undifferentiated stem cells in Japanese quail, as (a) since they are present in all stages of the cycle of the seminiferous epithelium (Lin and Jones 1990), (b) they occupy and make greater contact with the inner surface of the basal lamina than other spermatogonial types, (c) they form discrete, solitary cells, and (d) they reflect radiolabeling less frequently than the other spermatogonial types. The appropriateness and adequacy of this method of identification of the types and profiles of spermatogonial division is underscored by the observed number of spermatids (32), in each bundle embedded in Sertoli cells of the quail. The pale type Ap1 spermatogonia have the largest ovoid, spermatogonial nuclei, which are light-staining, although they contain a few (3-4) large and several smaller aggregations of chromatin (Fig. 7.10B). The cytoplasm is also light-staining, and slightly endowed with organelles. However, type Ap2 abounds in numerous, basally aggregated, mitochondria and associated RER, ribosomes, and an active Golgi complex (Fig. 7.11A). Cytoplasmic bridges commonly link adjacent Ap2 spermatogonia. Type Ap1 occurs in Stages X, I and III, while type Ap2 occurs during Stages III, IV, V and VI of the seminiferous epithelium (Fig. 7.9). The type B spermatogonia, obviously the last in the series to divide mitotically, are often connected to their fellows by cytoplasmic bridges (Fig. 7.11B). Both their cytoplasm and nuclei are densely stained with toluidine blue, in plastic sections. The nuclei are ovoid and display variably-shaped clumps of chromatin attached to the inner surface of the nuclear membrane. Multiple nucleoli (2-4) are scattered within the nucleoplasm. A prominent Golgi complex is basally situated. Mitochondria lie on either side of the nuclear poles. The cell lies on the basal lamina. The type Ad spermatogonia are considered by Lin and Jones (1992) to be the stem cells because they occur in all stages of the cycle of the seminiferous epithelium, as well as being solitary cells and not showing radiolabeling as frequently as the other types of spermatogonia. As a stem cell, the type Ad spermatogonium duplicates itself, during Stage IX of the cycle, by producing,

Fig. 7.9 The cycle of the seminiferous epithelium in the Japanese quail showing associations of germ cells in the 10 stages of the cycle. Ad, Dark type A spermatogonia; Ap1, pale type A1 spermatogonia; Ap2, pale type A2 spermatogonia; B, type B spermatogonia; L, leptotene primary spermatocytes; Z, young primary spermatocytes in zygotene; P, pachytene primary spermatocytes; Dp, diplotene primary spermatocytes; An, anaphase primary spermatocytes; II, secondary spermatocytes; 1-12, Step 1 to Step 12 spermatids. From Lin, M. and Jones, R. C. 1992 Cell and Tissue Research 267: 591-601, Figure 13. Reproduced with the kind permission of Springer Science and Business Media.

!& Reproductive Biology and Phylogeny of Birds

Spermatogenesis and Testicular Cycles

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Fig. 7.10 Coturnix japonica. Spermatogonia. A. Spermatogonium type Ad lies on the basal lamina, has a relatively small nucleus displaying small heterochromatin aggregations; mitochondria (M) are sparse and admixed with short profiles of RER (arrowheads). B. The spermatogonium type Ap1 also lies on the basal lamina, and displays a relatively large, less heterochromatic nucleus than Ad type. At least, two centrally located nucleoli occur in the nucleoplasm. Only sparse organelles occur in the cytoplasm. Bars: 2 µm for both figures. Original.

! Reproductive Biology and Phylogeny of Birds

Fig. 7.11 Coturnix japonica. A. Spermatogonium type Ap2 nucleus has clumps of heterochromatin lying in the nucleoplasm or attached to the nuclear membrane. A relative abundance of organelles, preponderantly mitochondria, occur in the cytoplasm. B. Spermatogonium type B has a nucleus that is relatively euchromatic and displays only a few clumps of heterochromatin as well as two to four nucleoli. The organelle content is moderate. Arrowheads, a cytoplasmic bridge between adjacent type B spermatogonium. Bars: 2 µm for both figures. Original.

Spermatogenesis and Testicular Cycles

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mitotically, a replicate (type Ad) stem cell, and a type Ap1 spermatogonium that differentiates to continue the proliferation and differentiational phases of the process of spermatogenesis. Types Ap1 spermatogonia occur during Stages X, I and II of the spermatogenetic cycle. Each type Ap1 spermatogonium divides and produces two type Ap2 spermatogonia during Stage II of the cycle, while the Ap2 spermatogonia, together, produce four type B spermatogonia in Stage VI of the cycle. The last mitotic division by the spermatogonial series occurs during Stage III of the cycle, when the type B spermatogonia, together, produce eight primary spermatocytes. The meiotic segment of spermatogenesis begins with primary spermatocytes that, eventually and ideally, produce thirty-two spermatids. The small number of spermatogonial divisions (three) in Japanese quail, compared, for example, with 6 to 7 in R. norvegicus (Hilscher and Makoski 1968; Clermont and Bustos-Obregon 1968; Huckins 1971a,b; Oakberg 1971a,b; Clermont and Hermo 1975) and 6 in Bos taurus (Hochereau 1967), is in accord with the small area occupied by stages of the cycle of the seminiferous epithelium, and, also, with the number of Sertoli cells in each stage, in the quail (Table 7.1). Theoretically, 32 spermatids arise from one stem cell (Fig. 7.8), and this number was actually counted in a bundle of spermatids embedded in Sertoli cells in the Japanese quail (Lin and Jones 1992). Germ cell degeneration occurs frequently in mammalian seminiferous tubules (RoosenRunge 1962). Germ cell loss, of about 22%, has been reported in the rat (Roosen-Runge 1958 cited by Roosen-Runge 1962) and in Mus musculus, the mouse (Oakberg 1956). Whether all 32 spermatids are present all the time, without loss of germ cells remains to be determined in birds. Numerous multinucleated giant cell balls are not uncommonly seen in the seminiferous epithelium of sexually mature and active ostriches (personal observations), as well as in cycling sexually immature Ostrich (Madekurozwa et al. 2002). The nature of these balls of cells has not been determined, but more studies on the kinetics of spermatogenesis in birds, in general, need to be undertaken in order that the process of spermatogenesis in this large class of animals may be better understood. Jones and Lin (1993) have calculated that Rattus norvegicus produces more spermatids per stem cell than Japanese quail by about 128-fold, and that the area occupied by a stage of the cycle of the seminiferous epithelium is 55 times more than that of Japanese quail. This very low value of spermatids (32) per stem cell in Japanese quail may be a consequence of the small area occupied by a stage in the cycle, compared to R. norvegicus.

7.5.2

Duration of Spermatogenesis

The proper evaluation of the function of the testis in health and disease is predicated upon a thorough understanding of the entire process of spermatogenesis, and the factors controlling it. An important aspect of this process is the life span of not only the individual cells but also their relationships to other cell types in the process. Spermatogenesis begins with the first spermatogonial division that initiates the spermatogenic series (Ortavant 1959). Discrepancies in the estimation of the duration of

!

Reproductive Biology and Phylogeny of Birds

spermatogenesis arise, partly, from certain methods used in investigating this process. The duration of spermatogenesis has been studied in various mammalian species, and according to Ortavant (1959) all germ cells that proceed in their development do so at the same speed, and hence the spermatogenic cycle seems to be a biological constant. The estimation of the duration of spermatogenesis in birds has, as in other aspects of male reproductive biology, also received scant attention. The employment of radiolabeling and autoradiography has, however, considerably improved the determination of this process. By means of radiolabeling and daily ejaculation methods, de Reviers (1975) has estimated the duration of spermatogenesis in the rooster to be between 12 and 13 d, which is the period elapsing between injection of tritiated thymidine and the release of spermatozoa from the seminiferous epithelium, and the presence of labeled spermatozoa in semen 13-14 d following radiolabeling. Labeled spermatozoa are, similarly, present in semen of the Barbary drake 12 d after [3H]thymidine injection (Marchand et al. 1977; Marchand 1979). Lin et al. (1990) estimate that the duration of one cycle of the seminiferous epithelium in Japanese quail is 2.69 ± 0.08 d. Therefore, Lin and Jones (1992) estimate (Fig. 7.9), that the process of spermatogenesis in the same species of bird lasts through 4.747 cycles or 12.77 d (i.e. 2.69 d ¥ 4.747 cycles), from the first spermatogonial (type Ad) division during Stage IX of the cycle to the release of spermatozoa from the seminiferous epithelium during Stage V. The duration of spermatogenesis, therefore, is similar in all birds studied thus far, and is found to be a much faster process than in mammals (25 days in Boar, Sus scrofa: Swierstra, 1967; 29 day in Sheep, Ovis aries: Ortavant 1959; 32d in Rabbit (Oryctolagus cuniculus): Amman et al. 1965). Also worthy of note is that the duration of spermatogenesis is constant, and independent of season, although transit time for spermatozoa through the excurrent ducts may vary with season (de Reviers 1975).

7.6

RESPONSE OF BIRDS TO REPRODUCTIVE DEMANDS ON THE TESTIS

Birds are generally socially monogamous, but a high proportion engage in extra-pair copulation during the short breeding season, especially in the temperate zone region (see, among others, Volume 6B, Chapters 6 to 9). Gallus domesticus has been known to mate up to 30 times a day (Penquite et al. 1930). If more spermatozoa originate from one division of a stem cell in the rat, and also in other mammals, than in birds, e.g. the quail, how do birds make up for this deficiency in number of germ cells per stem cell, in their reproductive process involving frequent copulations during the breeding season? It is known that reproduction in most species of birds is seasonal or discontinuous, an adaptation to a period environment (Wilson and Donham 1988). Most birds therefore have a short period of breeding, especially in the higher latitude (Wingfield et al. 1997a), and many are polygynous. For birds, reproduction is a very energy-demanding and resourced process. It therefore

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needs precise timing to coincide with good environmental conditions (Lack 1968; Perrins 1970; Price et al. 1988; van Noordwijk et al. 1995; Wikelski et al. 2000). Several biological strategies have been adopted by birds to accomplish their reproductive goals. Birds, generally, have both large absolute and, in particular, relative testis weights, as compared to mammals (Lake 1981; Clulow and Jones 1982), and sperm production has a high positive correlation with testes mass (Møller 1989). Similarly, there is a strong positive relationship between relative testicular size and relative sperm production, as well as between relative testes size and relative size of sperm reserve in mammals (Møller 1989), a situation that may be similar in birds (Møller 1991; Pitcher et al. 2005). It is therefore expected, and it has been shown, that several birds, e.g. the quail, have high absolute and relative testis weights, and hence a relatively large sperm content in ejaculates (Clulow and Jones 1982). However, figures need to be obtained for birds that are more volant than these relatively terrestrial species. Several species of birds are polygynous and/or are engaged in multiple pair and extra-pair copulations. Accordingly, relatively large testes ensure that such birds can engage in frequent pair, and, in particular, extrapair copulations, in order to avoid being cuckolded (Birkhead 1998). Most species of feral birds and several domestic birds are polygynous, and therefore engage in intense sperm competition. In passerine birds, the development of the seminal glomus, as a sperm-storage structure, is necessary in birds that can not afford to bear the burden of very large testes sizes, and accordingly, have low daily sperm production rates [e.g. 1.885 ¥ 106 day–1, or 35 ¥ 106 d–1 g–1 testis tissue for Zebra finch (Taeniopygia guttata)] (Birkhead et al. 1995) compared with a non-passerine bird [92.5 ¥ 106 d–1 g–1 testis tissue for Japanese quail] (Clulow and Jones 1982). The mass of this structure is positively correlated with body mass and with the relative testis mass (Birkhead 1998), and has been shown to attain maximum size in polyandrous or promiscuous passerine birds that are engaged in intense sperm competition (Birkhead et al. 1991). Sperm transport from the testis to the seminal glomus in the Zebra finch occurs primarily at night (Birkhead, T. personal communication), and the organ is considered to be a temporary overnight storage depot for spermatozoa (Birkhead et al. 1994) at temperatures that are lower than core body temperature (Wolfson 1954). What is lost in the small number of spermatogonial divisions and therefore sperm numbers per group of Sertoli cells (Lin and Jones 1993) is compensated for by the relatively large testes size/mass in birds (see Japanese quail and Rattus norvegicus, above (Clulow and Jones 1982). Thus, the daily sperm production rate (106/g testis) is 92.5 for Coturnix and 23.7 for R. norvegicus (Clulow and Jones 1982). Avian testes have high fluid content (Lake 1957; Aire 1979a; Clulow and Jones 1988) that flushes this large number of mostly immotile spermatozoa rapidly through the seminiferous tubular lumina into the excurrent duct system. The fluid absorptive properties and the microenvironments of the various excurrent ducts of the testis are indispensable for the maturation, viability and fertilizing ability of the post-

!" Reproductive Biology and Phylogeny of Birds testicular spermatozoa that traverse them (Turner 1991; Hinton and Palladino 1995; Hess et al. 1997). The efferent ducts of birds are extremely well developed, constituting a large volume proportion of between 35 and 62% (Aire 1979b) of the epididymis, and absorb about 86% (Clulow and Jones 1988) of the copious flow of testicular fluid into the epididymis. Referring to the reproductive organs of man, as probably also applicable to most mammals, de Graaf (1668, cited by Joclyn and Setchell 1972), long ago, observed as follows: “During the time the ingredients of the semen are propelled through the very long ducts of the testicle, the semen is elaborated in their cavities in such a way that what was watery and ash-like in the testicles becomes milky and thick in the epididymides. It is clear that the ducts of our tubules, epididymides and vasa deferentia are very long and were shaped by a most discerning Nature primarily so that the seminal matter might be better elaborated by long delay in transit”. It is also interesting and noteworthy that, according to Orgebin-Crist (1998), irrespective of the species, the length of the epididymis, or the rate of sperm production, the passage of spermatozoa through the epididymis of different mammals lasts approximately 10 days. The ‘long delay’ necessary for sperm maturation is absent in birds, in which regard, Nature seems to have adopted new strategies. It is established that whereas it takes spermatozoa of Japanese quail only one day to traverse the entire length of the excurrent duct system, it takes over 8 days, in the rat, to do the same (Clulow and Jones 1982). The rapid transit time of spermatozoa through the excurrent ducts of the testis, in birds, seems to be correlated with several structural and physiological modifications in both the spermatozoa and the ducts themselves. These modifications or peculiarities serve birds very well in sperm competition, which requires the availability of a large number of ejaculates of viable spermatozoa per day. In mammals, “there are dynamic interactions between the epididymal epithelium, the microenvironment and the spermatozoa, interactions which ensure optimal conditions for sperm maturation” (Hinton and Palladino 1995). On the other hand, spermatozoal motility in birds does not appear to require the involvement of secretions or luminal microenvironment of the ducts. Spermatozoa, obtained from the testis of the rooster have been shown to exhibit motility, and are able to fertilize eggs (Munro 1938). Motility of avian spermatozoa, however, increases cranio-caudally, as they traverse the excurrent ducts in non-passerine birds (Munro 1938; Bedford 1979) and Ostrich (Aire and Ozegbe, unpublished observations), although they are only motile in the seminal glomera in passerine birds (Bedford 1979). Avian spermatozoa are transported rapidly, and, unlike in mammals, are not normally stored in the excurrent ducts for any appreciable length of time, a requirement for sperm competition by male birds. Therefore avian spermatozoa do not require androgen-dependent prolongation of their viability in non-passerine birds, as is required in mammals (Munro 1938; Bedford 1979). Passerine birds appear to be different in this regard, because their spermatozoa may require androgen for their viability since the existence and perhaps maintenance of the seminal glomus is androgen-dependent (Bailey 1953).

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Birds, generally, have adopted a relatively simple form of spermatozoal maturation or modification along the reproductive tract. In mammals exposure of the itinerant spermatozoa, as they pass through the various microenvironments of the epididymis of mammals confers upon these cells their motility capability and fertilizing ability (Bedford 1967; Turner 1991). Further adaptations adopted by birds for rapid transport and maturity of spermatozoa through the excurrent ducts include the relative non-reliance of spermatozoa for proteins secreted by the duct epithelia for further maturational processes, as well as not requiring capacitation in order to fertilize ova (Howarth 1970, 1995). Although spermatozoa acquire about four Wolffian duct proteins as they pass through the epididymal duct unit of the rooster, Guineafowl and Japanese quail (Esponda and Bedford 1985; Morris et al. 1987b), and only one androgen-dependent protein of 17kDa in Japanese quail (Kidd 1982, cited by Jones 1998), the functions of these proteins, which are order-specific in birds (Esponda and Bedford 1985; Morris et al. 1987), is unknown. Although these and other proteins in the duct lumina may not be necessary for promoting the fertilizing ability of avian testicular spermatozoa (Howarth 1970, 1983), they do not have decapacitation ability on rooster spermatozoa (Dukelow et al. 1967), and it has not been determined if the proteins are necessary for sperm viability and/or fertility in the female reproductive tract. The motility, viability, fertility and durability of avian spermatozoa seem to be intrinsic, largely, rather than generated or sustained by the series of microenvironments of the various duct lumina through which they pass, rather rapidly. An intriguing phenomenon is the ability of the male fowl to allocate spermatozoa both strategically and differentially to a number of females in varying circumstances, such as female promiscuity, novelty, and reproductive quality (Birkhead et al. 2003).

7.7

TESTICULAR CYCLES

Reproduction in birds is a cyclical phenomenon (Lofts and Murton 1973), and this is an adaptation to a periodic environment, especially at middle and high altitudes (Wilson and Donham 1988). The purpose of seasonal breeding is to ensure that the mechanisms that govern gametogenesis are synchronized by environmental stimuli so that the young are produced during the most appropriate and favorable time of the year for their survival (Lofts and Murton 1973). In addition, in certain seasonally unpredictable environments, organisms cannot anticipate and prepare for a regular yearly reproductive period, and have therefore adopted strategies, within the reproductive cycle, to produce, opportunistically, their young at any time of the year (Hau et al. 2004). Seasonal breeding also permits the sexually resting or inactive bird to allocate vital resources to other organismal functions, such as build up of flight muscles in migrating birds (Bairlein and Gwinner 1994; Gwinner 1996). In all of these birds, therefore, testicular, and hence general reproductive activity, undergo morphological and functional changes that are imposed on them by hormonal changes which, in turn, are a result of the response of the neuroendocrine system to environmental signals or cues.

!$ Reproductive Biology and Phylogeny of Birds With the approach of or during unfavorable environmental circumstances, the reproductive activity is slowed down and/or halted. There is a carefully integrated and balanced relationship between the bird and its environment (Lofts and Murton 1966; Lofts et al. 1970), such that changes in the environment are monitored by the neuroendocrine system of the animal. Thus, environmental cues or stimuli exert influence on the neuroendocrine system, with a view to causing appropriate response with regard to stimulating or causing the cessation of gonadotropic release from the pituitary gland. According to Wingfield et al. (1997b) the endocrine system is well conserved in all vertebrates, and it is likely that any deviation from the general vertebrate model may represent adaptations rather than phylogenetic constraints. The degree of neuroendocrine activities is directly related to gonadal size and function. Natural selection has favored the establishment of annual breeding cycles, and an endogenous rhythm seems to fine-tune responses to environmental cues (Lofts and Murton 1973; Marshall and Serventy 1958).

7.8

ENVIRONMENTAL EFFECTS ON REPRODUCTION IN MALE BIRDS

It is now generally agreed that the degree of environmental predictability has considerable and far-reaching implications on the reproductive physiology of the organism (Wikelski et al. 2000) and, not the least, in birds. Thus, birds are able to transduce seasonal or environmental cues into appropriate physiological signals, via the neuroendocrine system to produce the various phases of the reproductive cycle. There are several environmental and other parameters that impact upon the reproductive cycle of birds, including photoperiodism, temperature, food availability, nesting sites, environmental perturbations, etc. According to Gwinner (1996) circannual rhythms are synchronized with and modified by environmental factors in a complex way but the endogenous mechanisms usually respond to environmental cues such that an optimal adjustment to season and latitude is guaranteed. The effect of each of these cues on the reproductive activity of the male bird will, hereunder, be briefly examined:

7.8.1

Photoperiodism

There has been almost a deluge of reports on studies involving the influence of photoperiodism in reproductive activities in birds since Rowan’s (1925) first publication on this subject. This review will, in the circumstances, be rigorously selective in presenting an up-to-date picture of this environmental index as it affects male reproduction in birds. Photoperiod is now widely known as a primary proximate factor that controls annual rhythms in the biology of a considerable variety of middle and high-latitude avian species (Wilson and Donham 1988; Gwinner 1996) and a major environmental stimulus that synchronizes the breeding season in some species of birds (Wolfson 1959; Marshall and Serventy 1956; Engels 1961). Long days stimulate LH secretion in intact males merely by reducing

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the efficacy of testosterone negative feedback on the hypothalamohypophyseal axis (Wilson 1985). Thus, the reduced sensitivity of the hypothalamo-hypophyseal axis to testosterone enhances LH secretion, and hence, its concomitant effect on spermatogenesis. Major increases in the levels of LH and FSH are pre-requisites for testicular development as well as recrudescence in resting organs. Periodic photoperiodism is, however, supplemented, modified and fine-tuned by a variety of other factors (Immelman 1971, 1973; Murton and Westwood 1977; Farner and Follet 1979; Farner and Gwinner 1980; Wingfield and Farner 1980; Wingfield and Kenagy 1991; Wingfield et al. 1992; Gwinner 1996), which, together, bring about a coherent organismal response. Although day-length is still maximal and above the gonad-stimulating threshold, during summer, the chronically stimulated gonads are no longer responsive to it, and, indeed, normally commence regression, followed by reproductive collapse (Wilson and Donham 1988). By mechanisms, yet unknown, the hypothalamo-hypophyseal axis becomes insensitive to a normally stimulating photoperiodic threshold. Consequently, the testes regress in structure and function (Robinson and Follet 1982). This phase of adequate, but progressively ‘ineffective’ and non-stimulating, day-length on testicular function is known as “photorefractoriness” (Robinson and Follet 1982). Refractoriness, according to Robinson and Follet (1982), does not seem to depend upon the activity of the hypothalamo-hypophyseal axis as it develops in castrated or androgen-implanted birds. It is a necessary universal phenomenon in birds because it prevents the animal from assuming a dangerous physiological condition at an inappropriate season, e.g. for rearing the young during food scarcity (Marshall 1961; Lofts 1962; Lofts and Coombs 1965; Lofts and Murton 1973), and providing a resting and regenerative period in the reproductive cycle during which spermatogenesis, along with other reproductive activities, is terminated so that the ‘exhausted’ gonad can be rehabilitated (Marshall 1961) and rejuvenated. In temperate zone birds, annual reproductive cycles have evolved in response to proximate environmental information (especially photoperiod, food situation, and nesting sites) that regulates breeding seasonally (Wingfield 1980; Wingfield et al. 1992; Ball 1993; Nager and van Noordwijk 1995; Morton 2002). Even location and latitude (elevation) can influence reproductive development (Perfito et al. 2004). In temperate zone birds, testicular recrudescence begins in spring and reproduction terminates by early summer, whereupon the testes rapidly regress and remain small throughout the winter months (Lofts and Murton 1973). However, it was thought that in the tropical zone, the changes in photoperiod were too small to be used as seasonal cues by birds. Recent reports have begun to show that most tropical birds exhibit a distinct seasonal pattern in life history parameters such as breeding activity (Snow and Snow 1964; Fogden 1972; Stiles 1980; Bell 1982; Dittami and Gwinner 1990; Tye 1991; Hau et al. 1998; Wikelski et al. 2000). Hau et al. (1998) and Wikelski et al. (2000) have demonstrated that tropical zone birds are extremely sensitive to variation in

!& Reproductive Biology and Phylogeny of Birds photoperiod of as little as 17 minutes, and use it as an environmental cue, and as an important characteristic of their natural tropical habitat. The ability of these birds to transduce the photoperiodic changes into physiological signals, fine-tuned by supplementary cues, is considered to be similar to the situation in temperate zone birds (Wingfield 1980; Wingfield and Moore 1987). An adequate level of environmental signal causes an elevation of LH levels (Wikelski et al. 2000) in birds. This occurs as an adjustment of their reproductive activity to differences in the availability, as well as quality, of food (Hau et al. 2000). Ground doves (Columba squamata), in seasonal savannas in Venezuela, while breeding, mostly, during the dry season, retain fully functional testes independent of rainfall or longer photoperiod. It is significant and note-worthy that a member of the Ratitae, the emu (Dromaius novaehollandiae), differs from many, if not most, birds, in being a short-day breeder in southwestern Australia (Malecki et al. 1998). This bird breeds between May and August, during the winter period. The control of the reproductive cycle of the emu appears to have an endogenous component that uses seasonal change in photoperiod and rainfall as the critical environmental factor in order to ensure breeding at the appropriate time of the year (Malecki et al. 1998; Sharp et al. 2005). It appears that rainfall and food availability have an overriding influence on increasing day-length in this species. It is of interest that testes sizes in tropical wild birds exhibit relatively small seasonal fluctuations (Bosque et al. 2004), and that the relative testes sizes are much smaller (by about 10-fold) than those in temperate zone birds (Wikelski et al. 2000). Some species are, understandably, opportunistic breeders, and may maintain an activated reproductive system all year round (Hau et al. 2004). Polygynous species, nevertheless, have larger testes than monogamous species (Wikelski et al. 2000), and, as in temperate zone birds, plasma testosterone levels are positively correlated with the total number of spermatozoa produced and proportion of normal sperm morphology (Wikelski et al. 2000; Garamszegi et al. 2005). Wikelski et al. (2003) consider that the pace of life is slower in birds in the more sedentary tropical environment than in the northern temperate migratory individuals of the same species. That is probably why the tropical species, living in ‘stable’ saturated environments tend to experience less extra-pair copulation and tend to breed less synchronously throughout the year than do the generally synchronous temperate or high-altitude species (Stutchbury et al. 1998). Temperate or high altitude species have a short breeding season, and an elevated plasma T level throughout the season (Levin and Wingfield 1992; Moore et al. 2002; Wingfield et al. 1997a).

7.8.1.1

Supplementary environmental signals

7.8.1.1.1 Food availability Breeding in birds usually coincides with suitable feeding conditions (Lofts and Murton 1973; Wingfield 1988; Hau et al. 1998; Wikelski et al. 2000). Food

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availability is regarded as an ultimate factor or as supplemental information in seasonal breeding activities of tropical birds (Fogden 1972; Snow 1976; Sinclair 1978; Poulin et al. 1992; Young 1994). Some birds are able to withstand the deleterious effects of low temperature if they have free access to food (Wingfield 1984; Wingfield et al. 1982). The availability of nesting sites and the process of nest-building has been shown, also, at times, to exert proximate control over the reproductive schedule in the Mountain white-crowned sparrow (Zonotrichia leucophrys oriantha) (Morton 2002).

7.8.1.1.2

Ambient temperature

The influence of ambient temperature on reproductive activities in birds has received conflicting appraisal (Farner and Mewaldt 1952; Lewis and Farner 1973; Wingfield et al. 1997b). Low ambient temperature probably stimulates the hypothalamo-hypophyseal-thyroid axis to cause an increase in blood T3 and/or T4 (Assenmacher 1973; Smith 1982), but T4 levels have been found to be lower in Mountain white-crowned sparrows (Zonotrichia leucophrys oriantha) subjected to low temperature (Wingfield et al. 1997b). Earlier studies showed that low ambient temperature delayed photoperiod-induced gonadal growth (Marshall 1949; Lofts and Murton 1966; Storey and Nicholls 1982), but Lewis and Farner (1973) and Wingfield et al. (1997b) have come to the conclusion that various temperature regimens have marginal or no effect on testes sizes and growth, as well as cloacal protuberance in birds that are also exposed to long days. However, low temperature retards gonadal regression, while high temperature has different effects on gonadal recrudescence, onset of breeding and termination of breeding, in ways that are not clearly understood (Wingfield et al. 1997b). The interaction of both proximate and supplementary environmental cues and stimuli appears to be part of this lack of understanding. Disturbances during any phase, especially during the active breeding phase of the reproductive cycle, may trigger off responses that may compromise gonadal structure and function in birds (Wingfield 1988).

7.8.1.1.3

Overcrowding

Overcrowding has been known to cause a reduction in body weight, testis dysfunction (as evidenced by a fall in seminiferous tubular diameter and epithelial height), spermatogenic inhibition, Leydig cell function and therefore androgen production (De Tapas et al. 1974).

7.9

CYCLICAL MORPHOLOGICAL CHANGES IN THE TESTIS

Most wild birds and a few domestic birds are seasonal breeders. In such birds, there is a sequence of events, mediated by environmental cues, that occurs in a cyclical manner in the reproductive organs. During the breeding or sexually active phase of the reproductive cycle, such birds possess reproductive organs that are maximally differentiated and hormonally stimulated, and thus have maximally functioning epithelial cells. During this phase, the reproductive organ and tract produce and modify the gametes for maximum viability and

!  Reproductive Biology and Phylogeny of Birds fertility. This phase is followed by an involutionary phase, a period of declining, and ultimately total cessation of, function in the testis and its excurrent ducts. This phase includes, or is followed by (depending on the authors), a period of recrudescence during which phase of the reproductive cycle both the gonad and tract are prepared, morphologically and functionally, for resumption of full reproductive activity. Marshall (1961) and Lofts and Murton (1973) have described three main phases in testicular function, in the annual reproductive cycle of the male bird, viz. Regeneration phase, Acceleration phase and Culmination phase. On the other hand, Mehrotra (1962) and Traciuc (1969) have divided the reproductive cycle of birds into four phases, in their reports on seasonal effects on the excurrent ducts of birds, as follows: Mehrotra (birds in India) (1) Presecretion phase (October to December) (2) Reproductive phase (January to March) (3) Regression phase (April to July) (4) Refractory phase (July to September)

Traciuc (birds in Central Europe) Progressive phase (March) Active secretory phase (April) Reconstruction phase (May to June) Resting phase (June to February).

It is clear that the latter authors have split the regeneration phase (Marshall 1961; Lofts and Murton 1973) into their respective regression and refractory phases of the reproductive cycle. These are minor non-significant differences, which should, however, be noted.

7.9.1 Basic Distinguishing Structural Features of the Testis in Juvenile and Reproductively Resting Sexually Mature Birds It is essential that certain morpho-physiological features of the testis in the three main phases of reproductive development or activity (juvenile, sexually mature and active, and sexually mature but inactive or resting) be established so that by evaluating the structure of any testis, the reproductive history of the bird can be appropriately determined. The following table (Table 7.5), sketches the main differences in the testes of (a) sexually immature/juvenile birds, (b) sexually mature and active birds and (c) sexually mature but resting birds, represented histologically in Fig. 7.12.

7.9.2

Structural Changes in the Testes of Sexually Mature Birds

At the end of the active period of reproduction certain profound changes occur in the structure of the testis during the regeneration phase of the reproductive cycle. According to Humphreys (1975b), cyclic changes in the testis seem to commence with the Leydig cells, and subsequently, the tubular and interstitial tissue in each phase of the reproductive cycle.

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Table 7.5 Main morphological features of testes in three different phases of development and/or activity Sexually immature a. Small testis, firm to touch. b. Cut surface relatively ‘dry’ and does not bulge out. c. Small seminiferous tubular diameter. d. Intertubular tissue is moderately voluminous and uniformly granular. e. Seminiferous epithelium contains numerous basal cells, mainly supporting (future Sertoli) cells and spermatogonia or gonocytes in a homogeneous matrix. There are no evident lipid droplets or dense bodies in the cytoplasm of the supporting cells in plastic sections. Tubular lumen is absent or only small spaces occur centrally, when fluid secretion commences.

Sexually mature and active Sexually mature but resting Large testis, soft to touch. Cut surface very fluid, and bulges out. Large seminiferous tubular diameter. Intertubular tissue is sparse and compact. Full germ cell complement in seminiferous epithelium. Sertoli cells contain a few dense bodies and residual bodies in the apical cytoplasm in toluidine bluestained plastic sections. Tubular lumen is large and fluid-filled

Small testis, firm to touch. Cut surface relatively ‘dry’ and does not bulge out. Small seminiferous tubular diameter. Intertubular tissue is abundant, with foamy islets. Seminiferous epithelium is distinctly heterogeneous, contains abundant, dense bodies, lipoidal and lipofuchsin granules in Sertoli cell cytoplasm; spermatogonia and only a few, degenerating spermatocytes are present. Tubular lumen is usually obliterated.

(a) During the regenerative or preparatory phase, in spite of its name, involutionary or regressive changes occur throughout the testis, from the testicular capsule to the core of the seminiferous epithelium. This phase is usually dependent upon environmental cues, such as shortening day-length and/or declining food availability. The testis is in the refractory phase, a period of declining, followed by arrest of, reproductive activity during maximal day-length (Robinson and Follet 1982). This phase is a normal feature of the reproductive cycle of birds, except for some domestic species, such as the domestic fowl, that have been bred for continuous reproductive activity. The testis involutes and its size is greatly reduced. The testicular capsule sloughs off and is replaced by a new one generated from below by fibroblast proliferation (Marshall and Serventy 1957). In the early stage, the interstitial tissue appears foamy, following accumulation of lipid droplets in Leydig cells and certain peritubular myofibroblasts (Fig. 7.13A). Macrophages and other mononuclear cells invade the interstitium and remove degenerating cells. A new set of Leydig cells seems to develop from myofibroblasts (Nicholls and Graham 1972; Aire 1997). The length of time required for the rehabilitation of the interstitial cells may vary from species to species (Lofts and Murton 1973). The Sertoli cells in the seminiferous tubules are filled with lipid materials including lipofuschin materials, and, along with spermatogonia, are the main cells left in the seminiferous epithelium. At the height of this phase, most of the worn and degenerating cells within the seminiferous tubule and interstitial tissue are removed by macrophage

!

Reproductive Biology and Phylogeny of Birds

Fig. 7.12 Struthio camelus (A); Anas platyrhynchos (B, C). A. seminiferous cords of juvenile testis surrounded by an obvious cellular intertubular tissue. The cords are devoid of lumina and exhibit mainly supporting cells and gonocytes. B. The seminiferous epithelium epithelium (E) of a sexually mature and active testis displays a full complement of germ cells and a wide lumen (L). C. Involuted seminiferous tubules in a regressed testis. The intertubular tissue is bulky and, once again, quite cellular, while the seminiferous epithelium contains mainly Sertoli cells that are laden with lipid droplets, and lipofuschin and dense granules. The tubular lumen is obliterated. Bars: A = 200 µm, B = 10 µm, C = 200 µm. Original.

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activity, and the testicular capsule sloughs off (Lofts and Murton 1973; Breucker 1978). The testicular framework is ready for regeneration by gonadotropins and the attendant hormonal action, following stimuli from appropriate environmental and endogenous cues, such as increasing daylength, rainfall and abundant food sources. The seminiferous tubules of sexually mature but gonadally inactive birds are physiologically atrophic, and contain both Sertoli and germ cells, comprising spermatogonia and a few primary spermatocytes. Sertoli cells are relatively preponderant, fill the highly reduced or obliterated tubular lumen, and are greatly laden with numerous, dense inclusions, as well as numerous, large, lipid droplets and lipofuschin pigments resulting from phagocytosis of degenerating germ cells (Fig. 7.13B). Macrophages in the seminiferous tubules of regressing testes in the Mute swan (Cygnus olor), aid Sertoli cells, with which they seem to have a preferential contact, in the regressing testes, to dispose of large amounts of cell debris, by phagocytosis (Breucker 1978). The Sertoli-Sertoli occluding junctions are intact structurally and functionally in the seminiferous epithelium in regressed testes of Mallard, although this junction appears to be apically displaced, due to atrophy of Sertoli cell cytoplasm (Pellietier 1990). The basal lamina of the seminiferous tubule is irregular in outline, thickened, relatively electron-dense, and often invaginates into the germinal epithelium in the form of finger-like plicae or folds (Fig. 7.14A), whose function probably facilitates the flow of raw materials into or out of the seminiferous tubule (Chakraborty et al. 1976; Aire 1997). These folds contain numerous electron-lucent globules whose nature and function are unknown (Aire 1997). The interstitium becomes relatively bulky and foamy in appearance, as a result of development of lipid droplets in certain myofibroblasts and particularly in extant Leydig cells (Lofts and Murton 1973; Aire 1997). Peritubular tissue remains intact in involuted testes, but the increased content of rough endoplasmic reticulum (RER), accumulation of lipid droplets and the change from lamellar to tubular cristae in the mitochondria (Fig. 7.14) of some of the myofibroblasts lend support to the hypothesis that certain of these cells undergo morphological and functional cytodifferentiation, preparatory to transforming into Leydig cells in the recrudescent testis (Aire 1997). Leydig cells in involuted testes contain an unusually large number of dense granules that are probably lysosomes, an abundance of unextracted lipid droplets, swollen and vesiculated mitochondria, and highly heterochromatic nuclei (Fig. 7.15A,B). The SER is profoundly atrophic. Two types of Leydig cells are present in the interstitium (Fig. 7.15B). The one, apparently normal, contains electron-dense ground substance, numerous lipid droplets and well formed polymorphous mitochondria bearing tubular cristae within an electron-dense matrix, while the other displays a rarified or reticular, electron-lucent cytoplasm, partially extracted lipid droplets, dense bodies, which are probably lysosomes, a heterochromatic nucleus, and vesiculated, disorganized mitochondria. The latter cell appears to undergo

! " Reproductive Biology and Phylogeny of Birds

Fig. 7.13 Anas platyrhynchos. A. Early changes in the intertubular tissue of a regressing testis. Leydig cells and myofibroblasts display lipid droplets. B. A TEM micrograph of the seminiferous epithelium of a fully regressed testis. The epithelium contains Sertoli cells predominantly. Dense bodies (straight arrows) abound in the Sertoli cell cytoplasm and the basal lamina (curved arrow) is thickened. S, Sertoli cell nucleus. Bars: A = 10 µm, B = 20 µm. Original.

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Fig. 7.14 Anas platyrhynchos. A. The basal lamina (arrows) of the seminiferous tubule (E) of a regressed testis is highly thickened, electron-dense, irregular in outline and contains numerous electron-lucent globules. Folds of the basal lamina (arrows) project into the Sertoli cell cytoplasm (S). Peritubular myofibroblasts (M) contain large lipid droplets (L) and irregularly-shaped nuclei (N). Electron-lucent globules also occur in the intercellular boundaries of the myofibroblasts. B. A myofibroblast contains a large lipid droplet (asterisk) and two, large mitochondria with tubular cristae (arrowheads). Bars: both figures at 1 µm. From Aire, T. A. 1997 Onderstepoort Journal of Veterinary Research 64: 291-299, Figures 11 and 12 (inset). Reproduced with permission of the Editor.

! $ Reproductive Biology and Phylogeny of Birds

Fig. 7.15 Anas platyrhynchos. A. The interstitial tissue between longitudinal sections of two seminiferous tubules increases in volume and shows two Leydig cells (Le) in a state of inactivity. D, an unusual number of dense bodies. B. Two Leydig cells (A and B) of varying electron-density and mitochondrial structure are present in the interstitium of a regressed testis. Type A cell is electron-dense and contains intact mitochondria (M) with tubular cristae, dense bodies (D), and numerous partially extracted lipid droplets. Type B Leydig cell is electron-lucent and contains vesiculated mitochondria (arrowheads), numerous lipid droplets and a heterochromatic nucleus. Bar: A = 20 µm, B = 1 µm. Figure A, original. Figure B is from Aire, T. A. 1997 Onderstepoort Journal of Veterinary Research 64: 191-199. Reproduced with permission of the Editor.

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degeneration. According to Lofts and Bern (1972), Leydig cells, in the nonbreeding season, disintegrate, are removed by macrophages, and are replaced by a new generation of Leydig cells. (b) The accelerative phase: The refractive period prepares the reproductive organs for receptivity and amenability to internal physiological stimulation. This process is normally mediated by gonadotropins secreted by the adenohypophysis, following appropriate environmental cues and appropriate stimuli from the brain that impinge on this endocrine gland. Blood levels of FSH and LH undergo increasing elevation (Haase 1983). The activities engendered in the reproductive organs, as a result of positive stimulation by gonadotropins are usually quite rapid, and bring about a flurry of growth activities in the gonads, which phenomenon is known or referred to as recrudescence. The period of recrudescence is usually short, if inhibiting factors such as low temperature (Lofts and Murton 1966) or lack of rainfall (Marshall 1970), are excluded or overridden. The Sertoli cells lose their lipoidal accumulations, spermatogenesis resumes, and the bird is ready to commence reproductive activity. (c) The culmination phase is when the male bird is morphologically and functionally prepared to be reproductively active, usually before the female bird is. Blood FSH, LH and T (testosterone) levels are at their peak. The Leydig cells lose most of their lipid droplets and both the smooth endoplasmic reticulum and mitochondria are restored to normal structural and functional states. The appropriate environmental and endogenous cues are present, and spermatogenesis progresses satisfactorily. Testosterone and semen production peak, and LH is correlated with testosterone level (Penfold et al. 2000). The absolute and relative weights of the testis attain maximum levels, and both the seminiferous tubular diameter and epithelial height attain full dimensions and functioning. The normal structure of the mature and active gonads and excurrent ducts are restored.

7.10 CYCLICAL MORPHOLOGICAL CHANGES IN THE EXCURRENT DUCTS OF THE TESTIS The seasonal effects on the structure of the excurrent ducts during the reproductive cycle of animals have received very scant attention. Even in mammals, there are very few reports of seasonally-induced changes in the excurrent ducts (Suzuki and Racey 1976; Pudney and Fawcett 1984). Aire (2002a,b) describes changes in various duct units in the epididymis of domestic, farm birds in three main phases of the reproductive cycle (prepubertal, sexually mature and active, sexually mature but inactive or resting) so that discrepancies may be prevented or minimized in the interpretation of normal structure in birds, during the sexually active phase, as distinct from the other two phases as well as their intermediate stages. The following account of changes in the various excurrent ducts comes, in the

! & Reproductive Biology and Phylogeny of Birds main, from the reports on fringillid (passerine) birds (Bailey 1953), in the Knob-billed goose (Sarkidiornis (=Anser) melanotus) (Mehrotra 1962), Jackdaw (Corvus monedula) (Traciuc 1969), Starling (Sturnus vulgaris) (Barker and Kendall (1984), Quelea (Quelea quelea) and Wild puffin (Fratercula artica) (Bhat and Maiti 2000), rooster, Mallard and Guineafowl (Aire 2002a,b). Only salient features will be emphasized in this account.

7.10.1

The Reproductive (Active Secretory/Culmination) Phase

The normal structure of the various segments of the excurrent duct system of birds has been described and discussed in considerable detail in Chapter 2. The rete testis lacunae are lined by a squamous to cuboidal epithelium comprising non-ciliated cells. Organelles are normal in structure, number and distribution. Small subapical vacuoles and multivesicular bodies, few small lipid droplets as well as a large, deeply indented, relatively euchromatic nucleus, occur in the cell. The efferent ducts: the lumen of the proximal efferent duct (PED) is large, irregular in outline and contains a few germ cells, while that of the distal efferent duct (DED) is regular in outline and contains a concentrated accumulation of sperm. The non-ciliated (NC) types I and II cells are maximally stimulated, structurally and functionally, and contain euchromatic nuclei which are round or oval in shape and situated in the basal half of the cells. The subapical endocytic or tubulovacuolar system is remarkably abundant and fully functional in the NC type I, but only scarcely noticeable in the NC type II, cell. Dense bodies are numerous and mixed with lysosomes in varying phases of activity in the type I, but are absent in type II, cells. The width of the mitochondria of both types I and II cells is much greater, by over 70%, than those of ciliated cells (Aire 2002a). The disappearance of lipid droplets seems to be a little protracted because large lipid droplets occur basally in the NC type I cells although they appear structurally and functionally primed, as evidenced by the full restoration of the subapical endocytic apparatus and dense bodies, that are normally atrophic or absent, respectively, in the regressed phase of the reproductive cycle. Lofts (1964) made similar observations in the testes of quelea birds. In the epididymal duct unit, the ductus epididymidis appears distinct and full of sperm. It runs distally into an easily recognised, wavy, turgid, and whitish ductus deferens, also full of spermatozoa. The receptaculum ductus deferentis is full of sperm. The papilla ductus deferentis is easily seen, projecting into the dorsolateral aspect of the urodeum. The apical microvilli of the NC type III cell are numerous, short and evenly distributed. The basally situated nuclei are oval/oblong in shape and relatively euchromatic. The Golgi complex is well developed and extensive. Profiles of smooth and sparsely granulated endoplasmic reticulum (SER and SGER, respectively) are abundant and moderately distended. The lateral plasmalemma is highly and intricately folded.

Spermatogenesis and Testicular Cycles

7.10.2

! '

The Regression (Reconstructive/Regeneration) Phase

The rete testis spaces contain accumulations of desquamated testicular germ cells at varying stages of degeneration, and are phagocytised by a large number of infiltrated macrophages and other mononuclear cells. Certain ducts are obliterated in Jackdaw, according to Traciuc (1969), but this has not been observed in the rooster, Mallard or Guineafowl, even in the most involuted organs (Aire 2000a,b). The proximal efferent duct (PED), as in the rete channels, contains an accumulation of degenerating desquamated testicular germ cells and mononuclear cell series that are actively spermiophagic. The duct diameters are, subsequently, greatly reduced, but the epithelium remains cuboidal to columnar. The epididymal duct unit contains degenerating spermatozoa and early germ cell series in an amorphous matrix that is eosinophilic and stretches the now cuboidal epithelium.

7.10.3

Refractory (Resting) Phase

In the rete testis, a highly irregular nucleus occupies most of the cytoplasm, is more heterochromatic than in the active phase, and may be indented by numerous, fairly large, lipid droplets and dense heterogeneous bodies that abound in the cytoplasm, during this phase (see Fig. 7.16). Several ciliated (C) cells appear in the epithelium, and are interspersed between the nonciliated (NC) cells that normally compose this epithelium (Fig. 7.16B). Androgen withdrawal seems to provoke ciliogenesis in the excurrent ducts of the testis in mice (Schleicher et al. 1984) and the middle segment of the epididymis of mice treated with the estrogen-like compound, 2, 6cis-diphenylhexamethylcyclotetrasiloxane (Aire, pers. obs.). Luminal macrophages and degenerating germ cells are few, having removed the degenerating luminal germ cell debris. The efferent duct unit contains granulofilamentous and some membranebound material in the PED lumen (Fig. 7.17), but the DED lumen is greatly reduced and empty. The columnar epithelial height of the PED is lower, but increases by over 30% in the DED than in the active phase of the reproductive cycle (Aire 2002b). In NC type I cells, the microvilli are considerably shortened, with highly irregular nuclei, especially in Mallard, occupying most of the shrunken cytoplasm. The subapical tubulovacuolar system atrophies in both duct segments, leaving only their silhouettes. Dense bodies which are typical of the NC type I cell are much fewer in number and much larger in size in Mallard, but are smaller, more numerous and contain lipofuschin-like granules in Guineafowl, during this phase (Aire 2002b). Mitochondria are smaller in NC type I cells, and are now similar in size to those of ciliated cells. Very large lipid droplets, some of which are as large as the nucleus (Fig. 7.18), occur in the supra- and infra-nuclear regions of the cell (Aire 2002b).

!! Reproductive Biology and Phylogeny of Birds

Fig. 7.16 Numida meleagris (A) and Anas platyrhynchos (B). A. The regressed rete testis epithelial cells accumulate a large number of lipid droplets (arrowheads) and dense, heterogeneous bodies, and the nuclei are highly irregular in shape and heterochromatic. Other organelles are inconspicuous. B. Ciliated cells (c) are yet to disappear from the rete testis of birds already in the active state, immediately following recrudescence. Bars: 20 µm for both figures. Figure A adapted from Aire, T. A. 2002. Anatomy Histology Embryology 31: 113-118, Figure 4. Reproduced with permission of Blackwell Publishing Ltd. Figure B, original.

The epididymal duct unit has an atrophic epididymal duct that is relatively straight, and contains caseous, amorphous eosinophilic coagulum. The epithelium of the duct undergoes profound ultrastructural changes (Aire 2002a). Microvilli are small and inconspicuous, the lateral plasmalemma is less folded, and highly elongated, heterochromatic nuclei occupy most of the rarified cytoplasm, SER and SGER are inconspicuous, while short, narrow profiles of RER appear more numerous (Aire 2002a). The seminal glomus shrinks, and its epithelial cells appear pyknotic in fringillid birds (Bailey 1953).

Spermatogenesis and Testicular Cycles

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Fig. 7.17 Anas platyrhynchos. The PED lumen is empty, save for granulofilamentous material (asterisk). The nuclei (N) of the non-ciliated type I cells are elongated, irregular in outline and heterochromatic. Large extracted lipid droplets (L) occur in the cell. Dense, polymorphic bodies are present in the supranuclear region. Non-ciliated cells undergo profound structural changes and tend to be crowded-out, over-shadowed by ciliated cells (C) that undergo minimal structural changes. Bar: 20 µm. From Aire, T. A. 2002. Journal of Morphology 253: 64-75. Figure 6. Reproduced with permission of Wiley-Liss, Inc.

!!

Reproductive Biology and Phylogeny of Birds

Fig. 7.18 Anas platyrhynchos. A. Lipid droplets (L) in the non-ciliated cells of an involuted PED may be nearly as large as the heterochromatic nucleus (Nu). Supranuclear dense bodies (D) are few but large. Mitochondria (M) of ciliated cells (C) are as large as those of non-ciliated cells (N). B. A light microscope section, and C. A TEM section of the PED, soon after gonadal recrudescence, exhibiting very large lipid droplets in the infranuclear region of the non-ciliated type I cells. Bars: A = 2.5 µm, B = 20 µm, C = 2 µm. Figure A is from Aire, T. A. 2002. Journal of Morphology 253: 64-75, Figure 7a. Reproduced with permission of Wiley-Liss, Inc. Figures B and C, original.

Spermatogenesis and Testicular Cycles

!!!

In general, the PED epithelium undergoes more profound structural changes than that of all other excurrent ducts of the testis during the regression and refractory phases of the reproductive cycle. Similarly, the NC type I cell of the PED is more greatly affected by androgen withdrawal than the NC types II and III as well as ciliated cells, with the latter showing less profound effects than any of the three types of non-ciliated cells (Aire 2002a).

7.10.4

Presecretion (Progressive/Acceleration) Phase

Rete testis tubules enlarge, their epithelium flattens into low cuboidal or squamous cells, with slightly flattened nuclei (Bailey 1953). The numerous, fairly large lipid droplets in the RT cells during the previous reproductive phase appear to coalesce and form large ‘globules’, occupying most of the cell cytoplasm. At the end of this phase, some ciliated cells are still present in the epithelium (Fig. 7.16). Diameters of the efferent duct unit increase and the epithelia undergo hypertrophy. Cellular division of both ciliated and non-ciliated cells occurs in the ducts (Mehrotra 1962), and organelle content of the cells becomes conspicuous, again. It has been noted that this phase is extremely short and not commonly observed in birds. In the drake, large lipid droplets that accumulate in the PED during the previous reproductive phase are still observed in the epithelium (Fig. 7.18), well beyond the full resumption of spermatogenesis and sperm accumulation in the excurrent ducts of the testis. It appears that some of the lipid droplets are extruded from the cells into the duct lumen (Aire, T. A., personal observations). The epithelial cells in the epididymal duct unit divide rapidly as the duct diameters increase. Some of the daughter cells are pushed inward and obscure the duct lumen (Bailey 1953). The diameter of the ductus deferens increases, and the epithelial cells of the seminal glomus proliferate rapidly, changing into low columnar or cuboidal type with round nuclei. Some of the new epithelial cells are sloughed into the lumen of the seminal glomus of various fringillid birds (Bailey 1953). Based on the review, above, on cyclical changes in the testis and its excurrent ducts, it is clear that more comprehensive, phase-controlled studies in both passerine and non-passerine birds are absolutely necessary. Available reports are few and disjointed, and some of them employ confusing and erroneous nomenclature for the ducts, in the light of more recent findings.

7.11

ACKNOWLEDGMENTS

I wish to acknowledge University of Pretoria for providing some resources that aided the writing of this review. The technical assistance of the Electron Microscope Unit of the Faculty of Veterinary Science of the University of Pretoria is also highly appreciated. Professor J. T. Soley kindly made several useful suggestions during the course of the compilation of the review. The helpful and insightful comments of Professor Tim Birkhead are gratefully acknowledged. Drs. Peter Ozegbe and Wahab Kimaro provided invaluable

!!" Reproductive Biology and Phylogeny of Birds technical assistance in the composition of the text and figures. Mrs. Wilma Olivier made the excellent line diagrams.

7.12

LITERATURE CITED

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Dietrich, T., Schulze, W. and Riemer, M. 1986. Untersuchung zur gliederung des keimepithels beim javaneraffen (Macaca cynomolgus) mittels digitaler bildverarbeitung. Urologe, Ausgabe A. Zeitschrift für Klimische und Praktische Urologie 25: 179-186. Dittami, J. P. and Gwinner, E. 1990. Endocrine correlates of seasonal reproduction and territorial behavior in some tropical passerines. Pp. 225-233. In M. Wada (ed.), Endocrinology of Birds: Molecular to Behavioral. Japanese Scientific Society Press, Japan, Tokyo. Dukelow, W. R., Chernoff, H. N. and Williams, W. L. 1967. Properties of decapacitation factor and presence in various species. Journal of Reproduction and Fertility 14: 393-399. Dym, M. 1973. The fine structure of the monkey (Macaca) Sertoli cell and its role in establishing the blood-testis barrier. Anatomical Record 175: 639-656. Dym, M. and Fawcett, D. W. 1970. The blood-testis barrier in the rat and the physiological compartmentation of the seminiferous epithelium. Biology of Reproduction 3: 308-326. Dym, M. and Fawcett, D. W. 1971. Further observations on the number of spermatogonia, spermatocytes and spermatids connected by intercellular bridges in the mammalian testis. Biology of Reproduction 4: 195-215. Eddy, E. M. 1970. Cytochemical observations on the chromatoid body of the male germ cell. Biology of Reproduction 2: 114-120. Ekstedt, E., Söderquist, L. and Plöen, L. 1986. Fine structure of spermatogenesis and Sertoli cells (Epitheliocytus sustentans) in the bull. Anatomy Histology Embryology 15: 23-48. Engels, W. L. 1961. Photoperiodism and the annual testicular cycle of the bobolink (Dolichonyx oryzivorus), a transequatorial migrant as compared with two temperate zone migrants. Biological Bulletin (Woods Hole) 120: 140-147. Esponda, P. and Bedford, J. M. 1985. Surface of the rooster spermatozoon changes in passing through the Wolffian duct. Journal of Experimental Zoology 234: 441-449. Farner, D. S. and Follet, B. K. 1979. Reproductive periodicity in birds. Pp. 829-872. In E. J. W. Barrington (ed.), Hormones and Evolution. Academic Press, New York. Farner, D. S. and Gwinner, E. 1980. Photoperiodicity, circannual and reproductive cycles. Pp. 331-366. In A. Epple and M. H. Stetson (eds.), Avian Endocrinology. Academic Press, New York. Farner, D. S. and Mewaldt, L. R. 1952. The relative roles of diurnal periods of activity and diurnal photoperiods in gonadal activation in male Zonotrichia leucophrys gambelii (Nuttal). Experientia 9: 219-221. Fawcett, D. W. 1961. Intercellular bridges. Experimental Cell Research, Supplement 8: 174-178. Fawcett, D. W. 1971. Observations on cell differentiation and organelle continuity in spermatogenesis. Proceedings of the International Symposium on the Genetics of the Spermatozoon, August 1971, 37-68. R. A. Beatty and S. Gluecksohn-Waelsch (eds.). Fawcett, D. W. 1975. The mammalian spermatozoon. Developmental Biology 44: 394-436. Fawcett, D. W. and Phillips, D. M. 1969. The fine structure and development of the mammalian spermatozoon. Anatomical Record 165: 153-184. Fawcett, D. W., Anderson, W. A. and Phillips, D. M. 1971. Morphogenetic factors influencing the shape of the sperm head. Developmental Biology 26: 220-251.

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Spermatogenesis and Testicular Cycles

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Wingfield, J. C. and Kenagy, G. J. 1991. Natural control of reproduction. Pp. 181-241. In P. K. T. Pang and M. P. Schreibman (eds.). Handbook of Comparative Endocrinology. Academic Press, New York. Wingfield, J. C. and Moore, M. C. 1987. Hormonal, social, and environmental factors in the reproductive biology of free-living male birds. Pp. 149-175. In D. Crews (ed.), Psychobiology of Reproductive Behaviour: An Evolutionary Perspective. Prentice Hall, Englewood Cliffs, USA, New Jersey. Wingfield, J. C., Hahn, T. P., Levin, R. and Honey, P. 1992. Environmental predictability and control of gonadal cycles in birds. Journal of Experimental Zoology 261: 214-231. Wingfield, J. C., Hahn, T. P., Wada, M. and Schoech, S. J. 1997b. Effects of day length and temperature on gonadal development, body mass, and fat depots in Whitecrowned sparrows, Zonotrichia leucophrys pugetensis. General and Comparative Endocrinology 107: 44-62. Wingfield, J. C., Jacobs, J. and Hillgarth, N. 1997a. Ecological constraints and the evolution of hormone-behavior interrelationships. Pp. 22-41. In C. S. Carter, I.I. Laderhandler and B. Kirkpatrick (eds.), The Integrative Neurobiology of Affiliation, Vol. 807. Annals of the New York Academy of Sciences, New York. Wingfield, J. C., Smith, J. P. and Farner, D. S. 1982. Endocrine responses of Whitecrowned Sparrows to environmental stress. Condor 84: 399-409. Wingfield, J. C. 1980. Fine temporal adjustment of reproductive functions. Pp. 367389. In A. Epple and M. H. Stetson (eds.), Avian Endocrinology. Academic Press, New York. Wolfson, A. 1954. Sperm storage at lower than body temperature outside the body cavity in some passerine birds. Science 120: 68-71. Wolfson, A. 1959. The role of light and darkness in regulation of refractory period in gonadal and fat cycles of migratory birds. Physiological Zoology 32: 160-176. Xia, L., Clermont, Y., Lalli, M. and Buckland, R. B. 1986. Evolution of the endoplasmic reticulum during spermiogenesis of the rooster: an electron microscopic study. American Journal of Anatomy 177: 301-312. Yamamoto, S., Tamate, H. and Itikawa, O. 1967. Morphological studies on the sexual maturation in the male Japanese quail (Coturnix coturnix japonica). II. The germ cell types and cellular associations during spermatogenesis. Tohoku Journal of Agricultural Research 18: 27-37. Yasuzumi, G. 1956. Electron microscopy of the developing sperm-head in the sparrow testis. Experimental Cell Research 11: 240-243. Yasuzumi, F. and Yamaguchi, S. 1977. Some aspects of spermiogenesis in the domestic pigeon. Okijamas Folia Anatomica Japonica 54: 139-174. Yasuzumi, G. and Sugioka, T. 1971. Spermatogenesis in animals as revealed by electron microscopy. XXI. Microkaryosomes and microtubules appearing during spermiogenesis of the lovebird Uroloncha striata var. domestica Flower. Zeitschrift Zellforschung 114: 451-459. Young, B. E. 1994. The effects of food, nest predation and weather on the timing of breeding in Tropical House Wrens. Condor 96: 341-353. Zlotnik, I. 1947. The cytoplasmic components of germ-cells during spermatogenesis in the domestic fowl. Quarterly Journal of Morphological Science 88: 353-365.

n n

CHAPTER

8

Avian Spermatozoa: Structure and Phylogeny Barrie G.M. Jamieson

8.1

INTRODUCTION

It was intended that this chapter would be confined to a review of ultrastructural works on bird spermatozoa and phylogenetic implications but it soon became apparent that the work, nearly a century old, of Gustaf Retzius (1909, 1911, 1912) and to a lesser extent the earlier publications of Emil Ballowitz (1886, 1888, 1913) still comprised a large proportion of our knowledge of avian sperm morphology. I have therefore included their light microscopical observations, and pertinent drawings of Retzius, with the later ultrastructural works in this chapter. Retzius’ drawings are comparable with those of Ernst Haekel (1862) for the extraordinary visual acuity, the excellence of the optical systems, and the dedication of these authors, on which their production depended. The light microscopical investigations of McFarlane (1963) are also of great value. The illustrations by Ballowitz are drawn to a smaller scale than those of Retzius and, though providing significant information, are not reproduced in the present work. Avian species examined for sperm morphology by Ballowitz, Retzius and McFarlane are tablulated in Tables 8.1, 8.2 and 8.3 respectively. This chapter is largely restricted to sperm morphology and ultrastructure and a phylogenetic analysis of these. For a consideration of spermatogenesis see Aire, Chapter 7 of this volume. For sperm biology see such works as Birkhead and Møller (1992), Briskie et al. (1997), Froman et al. (2002), and references therein, and, in this volume, Briskie and Montgomerie, Chapter 9, who give an extensive bibliography, including the important works of Birkhead and colleagues; and Stepinska and Bakst, Chapter 10. The School of Integrative Biology, University of Queensland, Brisbane, Queensland 4072, Australia

!# Reproductive Biology and Phylogeny of Birds To place the structure of the avian spermatozoon in an evolutionary perspective it is useful to briefly consider the general characteristics of amniote spermatozoa and particularly of crocodile sperm as crocodiles are widely held to be the extant sister group of birds.

8.2

AMNIOTE SPERMATOZOA, BASIC FEATURES

From detailed comparative and cladistic considerations, the following characteristics of a hypothetical plesiomorphic amniote spermatozoon (Fig. 8.1) may be recognized. This model is virtually identical with that of the lowest extant amniotes, the Chelonia, Crocodylia and Sphenodontida. Amniote sperm are seen to have few basal synapomorphies relative to the tetrapod ground plan which is deduced from common features of the amniote and lissamphibian sperm (Jamieson 1995, 1999, and references therein) and reference to sperm of sarcopgterygian fish (Jamieson 1991).

8.2.1 Amniote Sperm Plesiomorphies Plesiomorphic features of the generalized amniote spermatozoon, retained from their tetrapod ancestry (see also sarcopterygian fish in Jamieson 1991), and still seen in Chelonia, Sphenodontida, Crocodylia and to varying degrees in other amniotes, are as follows. The spermatozoon (Fig. 8.1) is elongate and filiform, with an anterior hollow conical acrosome vesicle overlying a simple subacrosomal cone. The base of the acrosome invests the tapered anterior tip (rostrum) of the nucleus and rests on pronounced nuclear ‘shoulders’. The subacrosomal space within the acrosome contains two or three axial rods (putative perforatoria) or, less likely, only one rod. These penetrate the nucleus deeply, almost to its base, in endonuclear canals. The nucleus is plesiomorphically elongate and cylindrical in amniotes from Chelonia through Sphenodon, crocodiles, squamates, birds, monotremes and, in therian mammals, the pangolin alone (Leung and Cummins 1988), as in lissamphibians (Scheltinga and Jamieson 2003a, b; Scheltinga et al. 2003). At the base of the nucleus there is a compact fossa (implantation fossa) with which are associated two triplet centrioles of which the distal forms the basal body of the flagellar axoneme. Whether the presence of an annulus in amniotes is plesiomorphic or an apomorphic reversal is debatable. The terminal portion of the 9+2 axoneme forms a short endpiece distinguished from the principal piece by the absence of the fibrous sheath.

8.2.2 Amniote Spermatozoal Synapomorphies The Chelonia and Sphenodontida are considered the most basal extant amniotes and have virtually identical spermatozoa (Healy and Jamieson 1992; Jamieson 1995; Jamieson 1999). The characteristics of these include features considered synapomorphies of the Amniota which are simultaneously symplesiomorphies for Chelonia and Sphenodontida and for the remaining amniotes, including birds. The amniote synapomorphies include:

Avian Spermatozoa: Structure and Phylogeny

!#

Fig. 8.1 Hypothetical plesiomorphic amniote sperm. From Jamieson, B. G. M. 1999. Pp. 303-331. In C. Gagnon (ed). Spermatozoal Phylogeny of the Vertebrata. The Male Gamete. From Basic Science to Clinical Applications, Cache River Press, Vienna, USA, Fig. 10.

Elongation of the distal centriole. The distal centriole is extremely elongate and extends the entire length of the long midpiece (the latter defined by its mitochondria) in turtles, the tuatara, crocodiles (Fig. 8.2), and paleognaths (Figs. 8.4-8.9), an apparent basal synapomorphy of amniotes. These elongate centrioles differ from most metazoan basal bodies in being penetrated by two central singlets from the axoneme. Thus in spermatids of the ratite Rhea, the distal centriole elongates and, late in spermiogenesis, becomes penetrated by

!#

Reproductive Biology and Phylogeny of Birds

a central pair of tubules from the developing axoneme (Phillips and Asa 1989). The shorter, though still elongate distal centriole in the rooster and the somewhat shorter centriole in Guineafowl (0.6 µm) and Geopelia striata (0.5 µm) (Jamieson 1995), the short centriole in squamates, and the vestigial centriole in monotremes possibly represent secondary reduction in length of the distal centriole (Healy and Jamieson 1992), culminating in almost total reduction in therian mammals (Jamieson 1999). Mitochondria, with concentric cristae. In turtles, tuatara (Healy and Jamieson 1992; 1994; Jamieson and Healy 1992; Jamieson 1995, 1999), Caiman crocodylus and Crocodylus johnstoni (Jamieson 1995; Jamieson et al. 1997; Jamieson 1999) (Fig. 8.2), the mitochondria have concentric cristae, known elsewhere in amniotes only in the sperm of some marsupials, notably the Woolly opossum, Caluromys philander (see Fawcett 1970; Phillips 1970) and the Virginia opossum, Didelphis virginiana (Temple-Smith and Bedford 1980) and also in the macropod Lagorchestes hirsutus (Jamieson 1999; Johnston et al. 2004). The mitochondrial cristae in the three ‘reptilian’ taxa (Chelonia, Sphenodontida, Crocodylia) usually surround a large central dense body. In all other amniotes studied, the cristae have a “conventional” appearance, being linear or curved, as in Lissamphibia, but never concentric, and do not surround a dense body. In spermatids of Sphenodon (Healy and Jamieson 1992; Jamieson and Healy 1992), the cristae have the linear appearance usual for metazoan sperm and the concentric arrangement is a late development. Phylogenetic “reversion” of concentric cristae to the linear condition seen in other amniotes would need only suppression of this final transformation (Jamieson and Healy 1992). Concentric cristae also occur in the spermatozoa of Gymnophiona (Scheltinga et al. 2003) and Urodela (Scheltinga and Jamieson 2003a). Although noting that multiple, homoplastic origin of concentric cristae would not be dismissed with certainty, Scheltinga et al. (2003) proposed that concentric cristae are an autapomorphy of tetrapods and not of amniotes as had previously been suggested by Jamieson (1999). They would therefore be symplesiomorphic for amniotes. The concentric arrangement appears to have been lost in all birds. The annulus. A dense ring, the annulus, at the posterior end of the midpiece is a feature of many metazoan sperm. It is clearly plesiomorphic for amniotes, occurring in all classes (Jamieson and Healy 1992), including paleognaths and several non-passerine orders, but absence in Dipnoi possibly indicates apomorphic re-acquisition in tetrapods. Irrespective of such reversal it is clearly a symplesiomorphy for Aves. Fibrous sheath. A dense fibrous sheath (Fig. 8.1) must, clearly, have developed, as an annulated structure, in the earliest amniotes as it is present in all amniote classes. With the exception of squamates, in which it penetrates the midpiece, it commences immediately behind the midpiece, as in turtles, Sphenodon (Healy and Jamieson 1992; Jamieson and Healy 1992) and crocodiles (Jamieson 1995, 1999; Jamieson et al. 1997); in ratites (Figs. 8.4 - 8.8),

Avian Spermatozoa: Structure and Phylogeny

!#!

Fig. 8.2 Diagrammatic longitudinal section of the spermatozoon of Crocodylus johnstoni. The spermatozoon of the Palaeognathae is closely similar to the crocodile spermatozoon. From Jamieson, B. G. M., Scheltinga, D. M. and Tucker, A. D. 1997. Journal of Submicroscopic Cytology and Pathology 29: 265-274, Fig. 1.

galliforms (Fig. 8.11, 8.14, 8.18), anseriforms (Fig. 8.19, 8.20), gruiforms (reduced, Fig. 8.32), charadriiforms (Fig. 8.34) and in mammals (Jamieson 1995, 1999). Nine peripheral axonemal fibers. Nine longitudinal dense fibers (coarse fibers) peripheral to the nine axonemal doublets, or to the distal centriole also where this is elongated as in chelonians, Sphenodon, crocodiles and paleognath birds,

!#" Reproductive Biology and Phylogeny of Birds are a fundamental feature of amniote sperm (Fig. 8.1), being found in all classes (Healy and Jamieson 1992; Jamieson and Scheltinga 1993; Jamieson 1995, 1999). As nine peripheral fibers are seen in lampreys and the fish Pantodon (references in Jamieson 1991) but also in heterobranch and cephalopod molluscs (references in Jamieson 1999) it might be considered that nine is the basic sarcopterygian, rather than merely amniote, number and that amphibians have lost all but those represented by the fibers at doublets 3 and 8. However, there is no evidence in extant Lissamphibia for such a reduction and the presence of only two lateral elements in dipnoans and Latimeria suggests that nine fibers were an amniote synapomorphy, albeit homoplastic with the other, non-amniote taxa. Fibers at 3 and 8. It is possible that a further basal amniote apomorphy is enlargement and lateral displacement of two fibers, at doublets 3 and 8, and that all fibers in the centriolar region intruded into the inter-triplet radii, as in ‘lower’ amniotes (Chelonians, Sphenodon and crocodiles) (Jamieson 1995, 1999). This displacement is not retained in birds. Longitudinal columns. In the principal piece, two longitudinal keel-like outward projections or thickenings (longitudinal columns) of the fibrous sheath opposite doublets 3 and 8, may be present, with or without inward projections to these doublets, as shown for mammalian spermatozoa (Fawcett 1975; Jamieson 1999) and Ostrich sperm (Baccetti et al. 1991; Soley 1993). Retronuclear body transformation. The striated columns of mammalian sperm are possibly derivatives of the tetrapod retronuclear body, present in dipnoans and (as the neck structure) in urodeles (Jamieson 1999). No definite conclusion can be made as to whether structures in bird sperm are homologues, viz. the non-segmented columns in Ostrich sperm and the three projections (stated to be probably equivalent to striated columns) in Crested pigeon (Ocyphaps lophotes) (Fig. 8.24C).

8.3

SPERMATOZOA OF CROCODYLIA

As the Crocodylia are traditionally considered to be the extant sister group of the birds a consideration of their sperm ultrastructure provides insights into the evolution of bird spermatozoa (we presumably will never know the sperm of theropods). The ground plan for the Crocodylia, as exemplified by Crocodylus johnstoni (Jamieson 1995, 1999; Jamieson et al. 1997) (Fig. 8.1), is very similar to that of the Chelonia and Sphenodon. All three have two or three endonuclear canals and, though requiring further confirmation for crocodiles, concentric cristae with intramitochondrial bodies. In Crocodylus johnstoni the mitochondria are subspheroidal to slightly elongate and possess few septate to (more externally) concentric cristae; a central dense mitochondrial body reported for Caiman crocodylus (Saita et al. 1987) is questionably present. Caiman crocodylus, resembles ratites in having only one perforatorium but this may be a reversion from multiple perforatoria homoplastic with the single condition in ratites.

Avian Spermatozoa: Structure and Phylogeny

!##

Synapomorphic conditions in the Crocodylia have little relevance to consideration of avian evolution. However, it may be noted that one condition, a dense sheath investing the two central singlets within the elongate distal centriole, previously considered an apomorphy of crocodile sperm (Jamieson 1999) is also seen in Ostrich sperm. It could therefore have been an apomorphic state of a common (theropod?) + crocodile + avian stock. Restriction of the endonuclear canal in caiman sperm to the anterior region of the nucleus indicated by Saita et al. (1987) appears to be apomorphic relative to the longer condition in Crocodylus johnstoni but requires confirmation. A similar trend to shortening in ratites and, progressively, in non-passerines presumably occurred in parallel.

8.4

SPERMATOZOA OF AVES—INTRODUCTION

As mentioned in the introduction, the light microscopical works of Ballowitz (1886, 1888, 1913) and of Retzius (1909, 1911, 1912) retain their significance in current avian spermatology. Species which they investigated are tabulated here (Tables 8.1, 8.2). Table 8.1 Bird species examined by Ballowitz for spermatozoal morphology by light microscopy Date

Figure

1888 1888

135-147 128-134

1888

121-127

1888

98-109

1888

110

1888 1888 1888

111-113 115 116-120

1888

114

1888

85-90

1888

91-97

1888 1888 1888

84 63-75 76-83

Species Galliformes Haushahns Truthahns Anseriformes Tadorna vulpanser Piciformes Picus major Cuculiformes Cuculus canorus Charadriiformes Larus ridibundus Larus canus Vanellus cristatus Falconiformes Milvus ater Caprimulgiformes Caprimulgus europaeus Columbiformes Haustaube Passeriformes Corvida Corvus frugilegus Oriolus galbula Lanius collurio

Valid name if different

Common name

Gallus domesticus Meleagris gallopavo

Rooster Turkey

Tadorna tadorna

Sheld-drake Great spotted woodpecker

Dendrocopos major

Cuckoo

Vanellus vanellus

Black-headed gull Common gull Lapwing Black Kite Nightjar

Columba livia

Domestic pigeon

Oriolus oriolus

Rook Golden oriole Red-backed shrike Table 8.1 Contd. ...

!#$ Reproductive Biology and Phylogeny of Birds Table 8.1 Contd. ...

Date

1888 1888 1888 1888 1888 1888 1888 1888 1888 1888 1888 1888 1888 1888 1888 1888 1888 1913

Figure

48-49 4, 11-15, 25, 38-39 19, 26-28 5 8, 20-22 36 37 51-52 16-17 18, 50 6-7 29-34, 4044 35 24 1-3, 23, 54-62 9-10 45-47, 53

Species

Valid name if different

Common name

Passeriformes Passerida Passer domesticus Muscicapa grisola

Muscicapa striata

House sparrow Spotted flycatcher

Hirundo rustica Chelidon urbica Sylvia nisoria Sylvia atricapilla Sylvia cinerea Sylvia hortensis Rubicilla phoenicura Motacilla flava Phyllopneuste hypolais Phyllopneuste sibilatrix

Hippolais icterina? Phylloscopus sibilatrix

Swallow House martin Barred warbler Blackcap Whitethroat Garden warbler Great Rosefinch Yellow wagtail Icterine Warbler Wood Warbler

Sitta europaea Fringilla canabina Fringilla caelebs

Carduelis canabina Fringilla coelebs

Nuthatch Linnet Chaffinch

Ligurinus chloris Emberiza citrinella Uria lomvia

Carpodacus rubicilla?

Cardeulis chloris

Greenfinch Yellowhammer Thick-billed murre (Brünnich’s guillemot)

Table 8.2 Bird species examined by Retzius for spermatozoal morphology by light microscopy. (Modified from Afzelius 1995) Volume (date) 16 (1911) 14 (1909) 14 14 14 14 14 14

Species Struthioniformes Struthio molybdophanes Galliformes Gallus gallus Anseriformes Anas boschas domestica Fuligula fuligula Gruiformes Fulica atra Rallidae Crex crex Charadriiformes Uria Troile Larus fuscus

Valid name if different

Common name

Struthio camelus

Somalian ostrich Domestic rooster

Anas platyrhynchos Aythya fuligula

Domestic duck Tufted duck Common coot Corncrake

Uria aalge

Common guillemot Lesser black-backed gull Table 8.2 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

!#%

Table 8.2 Contd. ...

Volume (date)

Species

14 14 14 14 14 14, 16

Vanellus vanellus Tringa alpina Totanus ochropus Scolopax rusticola Pavoncella pugnax CoIumbiformes Columba livia domestica Psittaciformes Psittacus sp. Strigiformes Syrnium aluco Apodiformes Cypselus apus Piciformes Dendrocopos major Passeriformes Corvida Corvus cornix Corvus frugilegus Perisoreus infaustus Coloeus monedula Pica pica Lanius collurio Passerida Sturnus vulgaris Turdus musicus Aedon luscinia Muscicapa atricapilla Phylloscopus sibilator Alauda arvensis Anthus obscurus Fringilla coelebs Chrysomitris spinus Chioris chioris Passer domesticus Emberiza citrinella

14 14 16 14 14

17 (1912) 17 16 14 16 14 14 14 14 14 14 14 14 14 14 14 14

Valid name if different

Calidris alpina Tringa ochropus Philomachus pugnax

Common name Lapwing Alpine dunlin Green sandpiper Woodcock Ruff Domestic pigeon A parrot species

Strix aluco

Tawny owl European swift Great spotted woodpecker

Corvus corone

Corvus monedula

Turdus philomelos Luscinia luscinia Ficedula hypoleuca Phylloscopus sibilatrix Anthus spinoletta Carduelis spinus Carduelis chloris

Hooded crow Rook Siberian jay Jackdaw Magpie Red-backed shrike Starling Song thrush Thrush nightingale Pied flycatcher Wood warbler Skylark Rock pipit Chaffinch Siskin Greenfinch House sparrow Yellowhammer

Some of the species of birds for which sperm are described in this chapter are illustrated in Fig. 8.3.

8.5

NEORNITHES

According to Gauthier and de Queiroz (2001) the name “Aves” refers to the crown clade stemming from the most recent common of Ratitae (Struthio camelus Linnaeus 1758), Tinamidae (Tetrao [Tinamus] major Gmelin 1789) and

!#& Reproductive Biology and Phylogeny of Birds Table 8.3 (1963)

Orders and families with number of genera and species examined by McFarlane

Non-passerines

Passeriformes

Procellarilformes Hydrobatidae (1 gen., 1 sp.) Ciconiiformes Ardeidae (2 gen., 2 spp.) Anseriformes Anatidae (1 gen., 1 sp.) Falconiformes Accipitridae (1 gen., 1 sp.) Galliformes Tetraonidae (1 gen., 1 sp) Phasianidae (1 gen., 1 sp.) Gruiformes Rallidae (1 gen., 1 sp.) Charadriiformes Charadriidae (1 gen., 1 sp.) Scolopacidae (2 gen., 2 spp.) Recurvirostridae (1 gen., 1 sp.)

Suborder Tyranni (Suboscines) Dendrocolaptidae (2 gen., 3 spp.) Furnariidae (1 gen., 1 sp.) Formicariidae (2 gen., 2 spp.) Cotingidae (4 gen., 4 spp.) Pipridae (2 gen., 3 spp.) Tyrannidae (12 gen., 14 spp.) Suborder Passeres Corvida Corvidae (2 gen., 2 spp.) Laniidae (1 gen., 1 sp.) Vireonidae (2 gen., 4 spp.) Passerida (Oscines) Alaudidae (1 gen., 1 sp.) Hirundinidae (4 gen., 4 spp.) Paridae (1 gen., 4 spp.) Sittidae (1 gen., 2 spp.) Troglodytidae (2 gen., 2 spp.) Mimidae (3 gen., 3 spp.) Muscicapidae (=Turdidae) (3 gen., 7 spp.) Sylviidae (2 gen., 2 spp.) Bombycillidae (1 gen., 1 sp.) Sturnidae (1 gen., 1 sp.) Coerebidae (1 gen., 1 sp.) Parulidae (13 gen., 27 spp.) Ploceidae (1 gen., 1 sp.) Icteridae (8 gen., 8 spp.) Thraupidae (7 gen., 8 spp.) Fringillidae (17 gen., 23 spp.)

Laridae (3 gen., 7 spp.) Rynchopidae (1 gen., 1 sp.) Alcidae (1 gen., 1 sp.) Columbiformes Columbidae (5 gen., 6 spp.) Cuculiformes Cuculidae (1 gen., 1 sp.) Strigiformes Strigidae (1 gen., 1 sp.) Caprimulgiformes Caprimulgidae (1 gen., 1 sp.) Apodiformes Apodidae (1 gen, 1 sp.) Trochilidae (5 gen., 5 spp.) Trogoniformes Trogonidae (1 gen., 1 sp.) Piciformes Picidae (1 gen., 1 sp.) Ramphastidae (1 gen., 1 sp.)

Neognathae (Vultur gryphus Linnaeus 1758). It seems preferable, however, to use the term Neornithes for this assemblage.

8.6

PALAEOGNATHAE

This is the Parvclass Ratitae of Sibley and Ahlquist (1990). According to Gauthier and de Queiroz (2001), “Palaeognathae” refers to the crown clade

CMYK !#'

CMYK

CMYK

Avian Spermatozoa: Structure and Phylogeny

Fig. 8.3 Some bird species examined for spermatozoal structure. A. Mallard (Anas platyrhynchos) Anseriformes. B. Coot (Fulica atra) Gruiformes. C. Crested pigeon (Ocyphaps lophotes) Columbiformes. D. Crimson Rosella (Platycercus elegans) Psittaciformes. E. Magpie lark (Grallina cyanoleuca) Corvida, Passeriformes. F. Southern ant-eater chat (Myrmecocichla formicivora) Passerida, Passeriformes. Photos © A, B, C, E, Barrie Jamieson. D, Christopher Tudge. F, Eric Hermann.

CMYK

!$ Reproductive Biology and Phylogeny of Birds stemming from the most recent common ancestor of Tinamidae (Tetrao [Tinamus] major Gmelin 1789) and Ratitae (Struthio camelus Linnaeus 1758).

8.6.1

Order Struthioniformes

The Order Struthioniformes contains the families Apterygidae, Casuariidae, Rheidae and Struthionidae. The Struthionidae is monotypic for the genus Struthio; Casuariidae contains the genera Casuarius and Dromaius; Rheidae contains the genera Pteroicnemia and Rhea.

8.6.1.1

Taxa examined

The following Struthioniformes have been examined for spermatozoal ultrastructure (see also Table 8.4): Struthionidae, Struthio camelus, Ostrich (Soley 1989; Baccetti et al. 1991; Soley 1993, 1994; Soley and Roberts 1994; Soley 1994b, 1996; 1999); Rheidae, Rhea americana albisceus, Rhea (Asa et al. 1986; Phillips and Asa 1989); Casuariidae, Dromaius novaehollandiae, Emu (Baccetti et al. 1991).

8.6.1.2

Overview of struthioniform spermatozoa

This diagnostic resumé is drawn chiefly from the accounts for Ostrich (Struthio camelus) sperm (Baccetti et al. 1991; Soley 1993, 1999; Soley and Roberts 1994). Lesser detail has been obtained from the accounts for Rhea (Rhea americana albisceus) (Phillips and Asa 1989) and Emu (Dromaius novaehollandiae) (Baccetti et al. 1991). The nucleus is an elongate cylinder and extends as a tapering nuclear rostrum far apically into the subacrosomal space (amniote symplesiomorphies). The acrosome vesicle is conical, shorter than the nucleus (amniote symplesiomorphy), about one tenth (Dromaius) to one fifth (Struthio) its length. The midpiece is shorter than the head (amniote symplesiomorphy), only about a quarter to a third of its length. An annulus is present at the junction of midpiece and principal piece (amniote synapomorphy by reversal?). The principal piece, several times the length of the nucleus, is defined by the presence of an annulated fibrous sheath (amniote synapomorphy) ensheathing the axoneme. There is a short endpiece consisting of axoneme (and plasma membrane) only. A subacrosomal space contains homogeneous material but no subacrosomal cone is present (synapomorphy of Aves). The nucleus is penetrated axially by an endonuclear canal (amniote symplesiomorphy) which contains the putative perforatorium. The perforatorium does not extend apically beyond the tip of the nucleus (Struthioniform + Timamiform synapomorphy), except, at most, for a little filamentous material. The endonuclear canal and enclosed perforatorium are absent in Dromaius (autapomorphy of this genus). There is a small basal nuclear fossa behind which is the proximal centriole orientated at right angles to the long axis of the spermatozoon (amniote symplesiomorphy). This is followed by the distal centriole (basal body) in the long axis and continuous with the axoneme. The distal centriole, also with nine triplets, is extremely elongate (amniote synapomorphy, reducing in non-paleognaths) and extends through the length

Avian Spermatozoa: Structure and Phylogeny

Table 8.4

!$

Ultrastructural studies on bird spermatozoa*

Taxon Struthioniformes Rhea americanus albisceus, Rhea

Struthio camelus, Ostrich Dromaius novaehollandiae, Emu Tinamiformes Eudromia elegans, Crested tinamou Anseriformes Anas platyrhynchos, Mallard Branta sandvicensis, Hawaiian goose Craciformes Galliformes Gallus gallus/domesticus, Rooster

Coturnix japonica, Japanese quail

Coturnix chinensis, Blue-breasted quail Meleagris gallopavo, Turkey

Tragopan caboti, Cabot’s tragopan Numida meleagris, Guineafowl

Turniciformes Piciformes Melanerpes carolinus, Red-bellied woodpecker Galbuliformes Bucerotiformes Upupiformes Coraciiformes Coliiformes Cuculiformes

Reference Asa et al. 19861 Phillips and Asa 1989 1 Baccetti et al. 19911 2; Soley 1989 1, 1993 1. 1994a,b1, 19961, 19991; Soley and Roberts 19942 Baccetti et al. 19911 2 Asa et al. 19861 2; Asa and Phillips 19871 Humphreys 1972 1; Maretta 1975a, b1 Humphreys 19721 None Grigg and Hodge 19491 2 ; Bonadona 19542; Nagano 19601, 19621; Mclntosh and Porter 19671; Nicander and Hillstrom 1967 1; Krustev and Danov 1968 1; Lake et al. 19681; Nicander 1970b 1; Tingari 1973 1; Bakst and Howarth 19751; Gunawardana and Scott 19771; Bakst and Sexton 19791; Bakst 19801; Xia et al. 19851; Xia et al. 19861; Xia et al. 19881; Bae and Kim 19871; Woolley and Brammall 19871; Thurston and Hess 1987 1; Sprando and Russell 1988 1; Jamieson 19991 Saita et al.19801; Maretta et al. 19821; Lin and Jones 19931; Woolley 19951; Tripepi et al. 19911; Vernon and Woolley 19991 This study1 Marquez and Ogasawara 1975 2; Bakst and Sexton 19791; Baccetti et al. 19801; Bakst 19801; Bradley et al. 19861; Thurston and Hess 1987 1 2 Wen et al. 19971 Hess et al. 19861; Thurston et al. 19821 2 ; Thurston and Hess 1987 2; Aire and Soley 20031 None Henley et al. 19781 None None None None None Table 8.4 Contd. ...

!$

Reproductive Biology and Phylogeny of Birds

Table 8.4 Contd. ...

Taxon

Reference

Crotophaga ani, Cuckoo

Saita et al. 1982; Tripepi et al. 1991 1; this study1

Psittaciformes Melopsittacus undulatus, Budgerigar

Agapornis roseicollis, Peach-faced lovebird Platycercus elegans, Crimson Rosella Nymphicus hollandicus, Cockatiel Apodiformes Apus (=Cypselus) apus, Common swift Apus melba, Alpine swift Trochiliformes Musophagiformes Caprimulgiformes Caprimulgus europaeus, European nightjar Columbiformes ‘Pigeon’ Columba livia, Domestic pigeon

Streptopelia roseogrisea, Turtle dove Geopelia striata, Peaceful dove Ocyphaps lophotes, Crested pigeon Gruiformes Grus vipio, White-necked crane The following 7 orders were previously placed in the order Ciconiiformes sensu Sibley and Ahlquist (1990) Charadriiformes Jacana jacana Falconiformes Falco peregrinus, Peregrine falcon Pelecaniformes Procellariformes Podicepediiformes Sphenisciformes Gaviiformes Passeriformes SUBOSCINES Tyrannldae Tyrannus verticalis, Western kingbird Tyrannus tyrannus, Eastern kingbird Contopus virens, Eastern wood peewee

Humphreys 19751; Samour et al. 19861 2; Jamieson et al. 19951. Jamieson et al. 19951 Jamieson 19991 Jamieson et al. 19951 Tripepi et al. 1984 1; this study 1; Jamieson and Tripepi 20051 Tripepi et al. 19911 None None Tripepi et al. 19911; this study1 Fawcett et al. 19711 Yasuzumi and Yamaguchi 19771 (spermiogenesis only); Vernon and Woolley 19991 Mattei et al. 19721 Jamieson et al. 19951; 1999 1; this study1 Jamieson et al. 19951; Jamieson 19991; this study1 Asa and Phillips 19871 2; Phillips et al. 19871 2

Saita et al. 19831; Tripepi et al. 19911 Wagley 19802 None None None None None

McFarlane 19711 3 Feduccia 19791 Feduccia 19791 Table 8.4 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

!$!

Table 8.4 Contd. ...

Taxon

Reference

Myiarchus crinitus/ M. griseisticta, Great crested flycatcher OSCINES Corvida Corvidae Cyanocitta cristata, Blue jay Corvus splendens, Crow Grallinidae Grallina cyanoleuca, Magpie lark Vireonidae Vireo olivaceus, Red-eyed vireo Vireo griseus, White-eyed vireo Passerida MUSCICAPOIDEA Muscicapidae (including Turdidae) Musclcapidae Myrmecocichila formicivora, S. Ant-eater chat Turdus greyi, Clay-colored robin Turdus merula, Blackbird Turdus migratorius, American robin Sturnidae Sturnus vulgaris , Starling Sylvioidea Paridae Parus bicolor, Tufted titmouse Hirundinidae Tachycineta thalassina, Violet-green swallow Certhioidea Troglodytidae Thryothorus ludovicanus, Carolina wren Troglodytes troglodytes, Wren Certhiidae Certhia brachydactyla Sittidae Sitta europaea PASSEROIDEA Parulidae Dendroica pinus, Pine warbler Dendroica dominica, Yellow-throated warbler Protonataria citrea, Prothonotory warbler Fringillidae Ammodramus maritimus (=Ammospiza maritima), Seaside sparrow Fringilla coelebs, Chaffinch

McFarlane 19711 3 ; Asa and Phillips 1987 1

Henley et al. 19781 Bawa et al. 19901 2 Jamieson 19951,19991 McFarlane 19713; Henley et al. 1978 1 Asa and Phillips 1987 1

Jamieson et al., unpublished1; this study1 McFarlane 19631 Furieri 19611 McFarlane 1971 1 3; Henley et al. 19781 Koehler 1995 1 2 ‘ Vernon and Woolley 1999

12

McFarlane 1971 1 3; Henley et al. 19781 McFarlane 19711 3

Asa and Phillips 1987 1 Tripepi and Perrotta 19911 Tripepi and Perrotta 19911 Tripepi and Perrotta 19911

Asa and Phillips 1987 1 Asa and Phillips 1987 1 Asa and Phillips 1987 1

McFarlane 19631 Furieri 19621; Tripepi and Perrotta 19911 Table 8.4 Contd. ...

!$" Reproductive Biology and Phylogeny of Birds Table 8.4 Contd. ...

Taxon

Reference

Pipilo erythrophthalmus, Rufous-sided towhee Piranga rubra, Summer tanager Serinus canaria, Canary Serinus canaria, Canary ¥ Cardeulis cardeulis Goldfinch, hybrid Zonotrichia albicollis, White-throated sparrow Carduelis (=Chloris) chloris Icteridae Agelaius phoeniceus, Redwing Icterus galbula, Northern oriole Molothrus ater, Brown-headed cowbird Quiscalus quiscula, Grackle Emberizldae Cardinalis cardinalis, Cardinal Emberiza cirlus, Cirl bunting Estrildidae Lonchura striata (=Uroloncha striata), Love bird Lonchura castaneothorax ¥ L. puntulata Taeniopygia guttata, Zebra finch

Henley et al. 19781

Passeridae Passer diffusus, Grey-headed sparrow Passer domesticus, House sparrow Passer italiae, italian sparrow Thraupidae Piranga rubra, Summer tananger Pioceidae Philetairus socius, Social weaver Euplectes orix, Red bishop Ploceus capensis, Cape weaver Quelea qualea, Red-billed quelea Prunellidae Prunella colIaris, Alpine accentor

Henley et al. 19781 Humphreys 19721 Swan 19851 Henley et al. 19781 Tripepi and Perrotta 19911 Asa and Phillips 19871; Koehler 19952 Asa and Phillips 19871 Koehler 19951 Koehler 19951 2 Henley et al. 19781; Koehler 1995 1 2 Tripepi and Perrotta 19911; Tripepi et al. 19911 Yasuzumi and Sugioka 19661; 19711. Yasuzumi 19741 2; Kondo et al. 19881 Swan and Christidis 19871 Nicander 1970a1; Fawcett et al. 19711; Asa and Phillips 19871 Vernon and Woolley 19991 Humphreys 19721; Asa and Phillips 19871 2 This study1 Koehler 19951 2 Furieri 19611 McFarlane 19711 3 Jamieson et al., unpublished1; this study1 This study2 This study2 This study2 Chiba and Nakamura 20011 2

1

TEM 2SEM 3fide Koehler (1995) *For a list of additional species examined by Birkhead et al. (2006) see section 8.10.12.17. For additional species examined by Tripepi and Perrotta (1991) see section 8.10.10.1.

of the midpiece. It is penetrated by the two central singlets of the axoneme (amniote synapomorphy) around which is a dense sheath (crocodile-bird synapomorphy, not described for Rhea and Dromaius and lost in nonstruthioniforms). The midpiece consists of approximately ellipsoid (Struthio,

Avian Spermatozoa: Structure and Phylogeny

!$#

Rhea) or spheroidal (Dromaius) mitochondria ensheathing the axoneme (amniote synapomorphy), four (Struthio), four to six (Rhea), or five to six (Dromaius) per transverse section, joined by electron-dense cement, totaling about 20-25 (Struthio), 30 (Rhea) or 40 (Dromaius) in a poorly defined helix, ending posteriorly at the annulus; cristae longitudinal plates, not in the basal amniote concentric arrangement. The fibrous sheath of the principal piece consists of semicircular ribs, enlarging near axonemal doublets 3 and 8 as longitudinal columns (amniote synapomorphies, replaced by amorphous sheath in other nonpasserines). A small dense fiber is attached to each doublet in the principal piece (Struthio, Rhea, shorter in Dromaius) (the fibers are a crocodile-avian symplesiomorphy, though reduced in the latter). At the posterior end of the endpiece, the doublets separate into individual components, and the dense contents of the A microtubules disappear. The resulting disorganized collection of 20 lucent microtubules displays a random decrease in number towards the tip (amniote plesiomorphy?).

8.6.1.3

Struthio camelus

The following account of Ostrich sperm is drawn from (Soley 1984b, 1989, 1993, 1999; Soley and Roberts 1994), with some substitution with the terminology of the present author, and from the earlier, succinct account of Baccetti et al. (1991). Spermatogenesis (Soley 1994, 1996) is discussed in Chapter 7 of this volume. General morphology. The mature spermatozoon of Ostrich is illustrated in Figs. 8.4 and 8.5. Examined by scanning electron microscopy (SEM) (Fig. 8.5QU) it is a 60 m long (70-80 mm in Soley 1989, 1999), filiform cell with an anterior tapering, cylindrical, 16 mm long head (acrosome + nucleus) (13 mm in Soley 1999) which reaches a diameter of 0.8 mm (Baccetti et al. 1991) near its base (greatest width of nucleus 0.5 µm, Soley 1999). A slight circular depression, the posterior ring of Soley (1999), marks the posterior limit of the 2 mm long acrosome. Immediately posterior to the head is the 3 mm long, 0.7 mm wide, midpiece, followed by the 40 mm long, 0.7 mm wide, cylindrical principal piece of the tail, tapering towards its posterior limit and followed in turn by 1 mm long, 0.2 mm wide, endpiece (Baccetti et al. 1991). A raised hooplike ring at the junction of the midpiece and principal piece was identified as the site of the annulus (Soley and Roberts 1994). Soley gives detailed tables of dimensions from a total of 200 sperm from 10 Ostriches, in which mean lengths, in mm, were: acrosome 1.91; nucleus (excluding the portion covered by the acrosome) 10.95; total head 12.86; midpiece 3.16; principal piece 51.18; endpiece 2.39; total tail 56.73; total sperm length 69.59. The mean tail to head ratio was 4.43. Differences in dimensions relative to those given by Baccetti et al. (1991) may be real as Soley and Roberts (1994) recognized two populations, possibly subspecies, in their own data, with one bird intermediate between the two. The head. The head of Ostrich sperm is a slightly curved, cylindrical structure tapering gradually at its most anterior aspect and measures 13 mm in length

!$$ Reproductive Biology and Phylogeny of Birds

Fig. 8.4 Struthio camelus, Ostrich. Schematic representation of a spermatozoon illustrating the various components of the head (acrosome and nucleus) and tail (midpiece, principal piece and endpiece). The three figures on the right show finer details, not drawn to the same scale) of the structures labeled on the whole spermatozoon (left). A. Acrosome. B. Midpiece. C. Principal piece. Relabeled after Soley, J. 1999. Pp. 129-158. In D. C. Deeming (ed). The Ostrich: Biology, Production and Health, CAB International, Fig. 6.5.

Avian Spermatozoa: Structure and Phylogeny

!

%$and 0.5 mm in width at its widest point (dimensions from Soley and Roberts 1994, throughout). The tip of the head is invested by the 2 mm long acrosome vesicle containing fine homogeneous material of moderate electron density (Fig. 8.5A, B) (Baccetti et al. 1991; Soley 1993). A subacrosomal space, 30 nm wide, is present between the inner acrosomal and nuclear membranes but this widens to 150 nm at the tip of the head (Baccetti et al. 1991). Flocculent material of an electron-density similar to the contents of the acrosome is observed in this space, often in close association with the nuclear membrane (Fig. 8.5A, B). Baccetti et al. (1991) state that the apical subacrosomal space contains a bundle of filamentous material which emerges from the endonuclear canal, opening at the nuclear tip; it is correctly regarded as a continuation of the rod (perforatorium) within the endonuclear canal. However, the prenuclear material is minimal and Soley (1993) did not recognize its presence. He regarded the avian perforatorium as merely residual. In contrast, Baccetti et al. (1991). considered that the subacrosomal filaments were probably actinic, as in other birds (Campanella et al. 1979). An area of close contact existed between the plasmalemma and nuclear membrane at the caudal extremity of the acrosome, forming a structure similar to the posterior ring of mammalian sperm (Fig. 8.5A) (Soley 1993). Nucleus. The nucleus forms a cylinder except for the part covered by the acrosome where it tapers sharply, as the nuclear rostrum, to end in a fine point beneath the tip of the acrosome. From the tip of the nucleus an axial endonuclear canal, lined by invaginated nuclear membrane, and containing the rod-like perforatorium, extends through the length of the rostrum for roughly 1/4 of the length of the main body of the nucleus (Soley 1993) or for ca 5 mm and about the apical third of the head, being ca 30 nm wide (Baccetti et al. 1991) (Fig. 8.5A-C). The chromatin is compact and electron-dense (Fig. 8.5C) excepting small light regions indicative of incomplete condensation throughout the nucleus, particularly in the rostrum. The basal, implantation fossa consists of a small central concavity surrounded by a shallow circular moat or series of depressions running around the perimeter of the nuclear base (Soley 1993). This was interpreted as two parallel implantation fossae, each with an undulating basal lamina by Baccetti et al. (1991). The neck and midpiece. Beneath the base of the nucleus, a short (0.3 mm) proximal centriole displays the characteristic nine sets of triplet microtubules (Baccetti et al. 1991; Soley, 1993). These are embedded in a ring of dense amorphous material. The central cavity of the centriole sometimes contains flocculent or granular material similar to that observed between the mitochondria of the midpiece (Fig. 8.5E). Dense material associated with the juxta-nuclear surface of the proximal centriole fills the center of the nuclear fossa, merging with similar peripheral material provided by dense nonsegmented columns emanating from the walls of the proximal and distal centrioles (Fig. 8.5D, E). The distal centriole is perpendicular to the proximal centriole, and occupies the entire length of the midpiece. In transverse section

!$& Reproductive Biology and Phylogeny of Birds

Fig. 8.5 Struthio camelus, Ostrich. A-P. Transmission electron micrographs. A. Longitudinal section (LS) of acrosome and anterior nucleus. B. Transverse section (TS) of acrosome vesicle and the enclosed nuclear rostrum with central Fig. 8.5 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

!$'

it consists of a narrow ring of electron-dense material from which nine evenly spaced dense projections jut into the centriolar cavity. Nine sets of characteristically arranged triplet microtubules are situated between the projections. Within the centriolar cavity are two singlets (central tubules) (Baccetti et al. 1991; Soley 1993, 1999). Around these Soley (1993) describes a rod, sometimes eccentric, of dense material extending posteriorly as far as the annulus and containing a pair of microtubules (Fig. 8.5D, E). The rod is occasionally seen in the form of two closely apposed but separate units, each containing a single microtubule. Posterior to the annulus a typical central pair of microtubules extends throughout the rest of the tail. The 0.3 mm long midpiece is slightly wider (0.65 mm) than the nucleus and contains about 20 (Soley 199) or 24-25 (Baccetti et al. 1991) mitochondria arranged in an helical pattern around the proximal and distal centrioles (Fig. 8.5D, F, S) as a single layer. The mitochondria have flattened, rectangular profiles (Figs. 8.4B, 8.5D), although round or oval forms are sometimes

Fig. 8.5 Contd. ...

endonuclear canal. C. TS nucleus and perforatorium. D. LS midpiece. Note the termination of the inner dense rod of the distal centriole in the vicinity of the annulus (white arrow). The dense walls of the distal centriole, which runs the length of the midpiece are indicated by black arrows. E. LS of the neck region showing the proximal centriole cut transversely. F. TS midpiece and distal centriole. A dense rod of material containing two singlet microtubules is eccentrically situated. Small black arrow heads indicate atypical cristae. G. LS proximal region of principal piece, showing annulus at posterior limit of midpiece, and fibrous sheath. Arrows indicate ribs of this sheath. H. Similar section but showing (arrows) longitudinal columns of the fibrous sheath. I. LS of a more distal region of the principal piece, through longitudinal columns (arrows) of the fibrous sheath. J. TS of the proximal principal piece, showing, above and below, the longitudinal columns. Dense fibers 3 and 8 have been incorporated into the columns. K. TS of two sperm tails posterior to the region with dense fibers but still surrounded by the fibrous sheath. L. LS at junction of midpiece and principal piece, showing the annulus I tangential section. M. LS distal region of principal piece. The ribs of the fibrous sheath are shown in cross section (arrows) and longitudinal profile (squat arrow). N. LS endpiece. Note the staggered termination of the fibrous sheath (arrows). O. TS intermediate region of the principal piece. The coarse fibers have disappeared. Doublets 3 and 8 are connected to the longitudinal columns. The cytoplasmic layer surrounding the axoneme is thinner than further proximally. P. TS endpiece at disruption of 9+2 pattern, resulting in a collection of doublets (arrows) and dense and translucent singlet microtubules. Q-U. Scanning electron micrographs. Q. Entire spermatozoon. R. Junction of principal piece and endpiece. S. Midpiece, with 5 tiers of mitochondria. T. Tapered tip of the nucleus (rostrum), between arrows, revealed after loss of the acrosome vesicle. U. Detail of the head and midpiece. A-P modified after Soley, J. T. 1993. Onderstepoort Journal of Veterinary Research 60: 119-130, Figs. 1-16. Q-U after Soley, J. T. and Roberts, D. C. 1994. Onderstepoort Journal of Veterinary Research 61: 239-246, Figs. 1-5.

!% Reproductive Biology and Phylogeny of Birds observed (Fig. 8.5D, S) and in tangential sections they appear as rectangular or polygonal structures (Fig. 8.5L) (Soley 1993) but Baccetti et al. (1991) found them to be elliptic in both longitudinal and transverse section; four in each cross section, joined by an electron-dense cement. The cristae are longitudinal, in a dense matrix. Atypical cristae containing paracrystalline inclusions occur in some mitochondria (Figs. 8.5F, arrow heads). The inclusions display two forms depending on the plane of section. Those sectioned longitudinally presented the appearance of tight junctions while oblique sections revealed a pattern of parallel fibers with a regular spacing and direction. Sandwiched between the mitochondria are conspicuous accumulations of granular material (Soley 1993) (Fig. 8.5F). Annulus. A well developed annulus (Figs. 8.4B, 8.5D, G, H) marks the boundary between the midpiece and the principal piece (Baccetti et al. 1991; Soley 1993, 1999), although a retro-annular recess is not apparent. The annulus is situated beneath the last row of midpiece mitochondria in close association with the plasmalemma and is composed of homogeneous, electron-dense material. Principal piece. The principal piece forms the longest segment of the tail (ca 50 mm) and consists of the 9+2 axoneme surrounded by a ribbed fibrous sheath (Soley 1993, 1999) (Fig. 8.5D-O). The ribs are semicircular and about 50 nm thick, enlarging near the axonemal doublets 3 and 8 (Baccetti et al. 1991) as the ‘longitudinal columns’ described below by Soley (1993). The A microtubule of each doublet is a circular structure filled with dense material whereas the B microtubule formed an incomplete lucent cylinder. Dynein arms project from each A microtubule towards the neighboring doublet and radial links form a connection with the central microtubules (Figs. 8.5K, O). The principal piece is a tapered structure and gradual decreases in diameter along its length, coupled with changes in the composition of the fibrous sheath, allow three regions to be distinguished: 1. The first region lies immediately posterior to the annulus where the tail abruptly narrows to a diameter of 0.5 µm. The axoneme is surrounded by a loosely arranged fibrous sheath consisting of two dense longitudinal columns connected by circumferential bands of dense material. The longitudinal columns and the interconnecting ribs appeared to be composed of alternating layers of electron-dense and loosely packed material, giving both structures a laminated appearance. The columns lie in line with the two central microtubules in the position occupied by coarse fibers 3 and 8 in mammalian sperm. Peculiar to this region is the presence of nine small (rudimentary, Soley 1999), dense, coarse fibers (the accessory fibers, reniform in cross section, noted by Baccetti et al. 1991) lying between the fibrous sheath and the axonemal doublets, each in close association with an A microtubule. A prominent cytoplasmic layer, maximally 60-80 nm wide (Baccetti et al. 1991) containing fine flocculent material is interposed between the fibrous sheath and the cell membrane (Fig. 8.5G-J, L).

Avian Spermatozoa: Structure and Phylogeny

!%

2. In the second region the diameter of the tail narrows to 0.4 µm, the rudimentary coarse fibers disappear, and the ribs of dense material appear as solid structures. The longitudinal columns, however, retain their laminated appearance while the cytoplasmic layer becomes narrower. Septum-like inward extensions of the longitudinal columns make contact with the adjacent microtubular doublets (Fig. 8.5J). 3. The third region of the principal piece is characterized by a progressive decrease in diameter, from 0.3 µm to approximately 0.2 µm. The longitudinal dense columns became solid, less conspicuous, structures which eventually disappear leaving a thin dense band of material surrounding the axoneme. The plasmalemma is closely applied to the layer of dense material (Fig. 8.5I, K, M) (Soley 1993). Endpiece. The endpiece forms the short, narrow, 2.4 mm long (Soley and Roberts 1994) or 2-3 mm (Soley 1999), termination of the tail and consisted of the axoneme covered only by plasmalemma. Because the transition from principal piece to endpiece is gradual, remnants of the fibrous sheath are sometimes seen around the axoneme (Soley 1993) (Fig. 8.5N). The organized structure of the axoneme is disrupted towards the end of the tail (Baccetti et al. 1991; Soley 1993, 1999). The specific orientation of the axonemal microtubules is lost, the dynein arms and radial spokes linking the microtubules have disappeared, the doublets separate into individual components, and the dense contents of the A microtubules disappear (Fig. 8.5P). The resulting disorganized collection of 20 lucent microtubules displays a random decrease in number at the tip of the endpiece (Soley 1993). The crocodiloid spermatozoon. Ballowitz and Retzius recognized the ‘sauropsid’ features of non-passerine spermatozoa. However, the ratite and lower non-passerine spermatozoon, especially the former, would more appropriately be termed crocodiloid. Features of Ostrich sperm which are similar to those of crocodiles are: the pointed acrosome vesicle; the perforatorium in a long endonuclear canal; the midpiece with several tiers of mitochondria surrounding an extremely long distal centriole and terminating at an annulus; presence of nine dense fibers; and a fibrous sheath consisting of transverse ribs (Figs. 8.4C, 8.5G). All of these features are also seen in Chelonia (Healy and Jamieson1992) and are basic (symplesiomorphic) to amniotes, only the fibrous sheath and the long distal centriole being amniote synapomorphies. The sole spermatozoal synapomorphy of crocodiles and birds is the dense sheath investing the two central singlets within the elongate distal centriole (Jamieson 1999). In birds this sheath is known only in ratites (Ostrich) and the Galloanserae.

8.6.1.4

Rhea americana albisceus

The following account of the sperm of Rhea, Rhea americana albisceus, is drawn from Phillips and Asa (1989). Additional data are added from perusal of their illustrations and current terminology is employed.

!%

Reproductive Biology and Phylogeny of Birds

Head. The head is curved and tapered. A substantial acrosome composed of moderately electron-dense homogeneous material fits over the anterior portion of the nucleus. A narrow cylindrical structure (endonuclear canal) extends from the anterior portion of the acrosome to deep within the nucleus (Fig. 8.6C, D). The center of the cylinder (putative perforatorium) is composed of material that is about the same electron density as the acrosome, which is circumscribed by an electron-lucid region, in turn surrounded by a thin region of moderate electron density (Fig. 8.6C, D). The chromatin is compact but not as condensed as is observed in spermatozoa of insects or mammals (Fig. 8.6AD). A short midpiece lies between the head and principal piece (Fig. 8.6A). The long principal piece comprises most of the length of the cell. Neck region and centrioles. As in many other animals, the neck region of Rhea sperm is characterized by a precisely shaped posterior portion of the nucleus. Electron-dense material associated with the proximal centriole fits into the contour of the sperm nucleus and associated nuclear membrane (Fig. 8.6A, D). The proximal centriole is short, only about 0.4 mm long, and is inlaid with electron-dense material (Fig. 8.6D, E). The distal centriole is much longer. It apparently extends the entire length of the midpiece. Both centrioles are embedded in electron-dense material. Although the distal centriole displays the characteristically disposed triplet subtubules, the central tubules of the flagellum extend into the center of the centriole (Fig. 8.6E). Midpiece. The midpiece contains about 30 mitochondria, four to six in transverse section, and about seven longitudinally, with a very dense matrix typical of spermatozoal mitochondria. In the interior of the mitochondrion distal to the centrioles, there is a complex configurations of mitochondrial membranes (Fig. 8.6A, D, E). Principal piece. Transverse sections through the principal piece reveal a small fibrous sheath. The sheath is larger opposite doublet tubules 3 and 8 and is connected to these two doublets by a thin band of dense material, as in mammals, but enlargement is only slight, less than in the latter (Fig. 8.6F). Dense axonemal fibers. There are very tiny dense fibers present only for a short region of the principal piece (see Fig. 8.6F), given (erroneously?) in the original account as the very anterior portion of the “midpiece”. They are no larger than microtubules and are associated with the doublet subtubules A and B (Fig. 8.6F). Remarks. The spermatozoon of Rhea is closely similar to that of Ostrich, the similarities including the deep extension of the endonuclear canal and perforatorium into the nucleus. Soley (1993) considers Ostrich sperm to differ from that of Rhea, however, in that the central tubules are embedded in a core of dense material which only disappears in the vicinity of the annulus. The account of Phillips and Asa (1989) does not refer to this feature and no sheath of dense material is visible in the relevant micrograph (Fig. 8.6E). In both Ostrich and Rhea, modifications of the mitochondrial membranes have been observed, with those of Rhea (Phillips and Asa 1989) resembling

Avian Spermatozoa: Structure and Phylogeny

!%!

Fig. 8.6 Rhea americana albisceus. A. Longitudinal section (LS) midpiece, penetrated by the long distal centriole, and terminating at the annulus. B. LS basal region of acrosome vesicle enclosing the nuclear rostrum and, at the center of this, the perforatorium. C. Transverse section (TS) of a nucleus and (right) the acrosome vesicle and nuclear rostrum. D. LS neck region showing short proximal centriole perpendicular to the distal centriole. E. TS midpiece; right, through the proximal centriole; left, through the distal centriole, showing two central singlets. F. TS through the principal piece of several spermatozoa. Relabeled after Phillips, D. M. and Asa, C. S. 1989. Anatomical Record 223: 276-282, Fig. 1. Reprinted with permission from Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc.

myelin figures and those of Ostrich adopting the form of atypical cristae containing paracrystalline material. The significance of these structural modifications is unknown. Although organized and arranged in a fashion

!%" Reproductive Biology and Phylogeny of Birds similar to that of mammalian sperm, the ribbed sheath of Ostrich sperm is more flimsy in structure, resembling that described in Rhea (Phillips and Asa 1989) and tinamou (Asa et al. 1986; Soley 1993). The latter author correctly observes that the ribbed form of the fibrous sheath seen in ratites does not occur in other non-passerines. We may add that the transverse ribbing is a basic amniote (Fig. 8.1) and crocodile (Fig. 8.2) feature. This unique crocodiloid feature of ratite sperm endorses the basal position of ratites in bird phylogeny. The proximal segment of the principal piece of Ostrich sperm displays a prominent cytoplasmic layer situated between the axoneme and the plasmalemma. This layer is filled with fine amorphous material and resembles a similar region seen in Rhea sperm (Phillips and Asa 1989). Tinamou sperm are structurally similar in this respect although particulate material which morphologically resembles glycogen is found throughout this region (Asa et al. 1986). A similar region is absent in other non-passerine birds (Soley 1993). As Soley (1993) observes, penetration of the long distal centriole by the central singlets of the axoneme is a feature shared by Rhea and Ostrich but is absent from Tinamou. As noted by the present author, this penetration is a crocodilian (and chelonian) feature and is therefore a symplesiomorphy of Rhea and Ostrich signifying an unchanged basal amniote condition but not necessarily indicating rheid-struthionid monophyly.

8.6.1.5

Dromaius novaehollandiae

General morphology. The mature sperm cell of Emu seen by SEM (8.7A-C) is ca 65 mm long. The cylindrical, tapered head measures ca 12 mm in length and 0.8 mm in maximum diameter. The acrosomal region is 1.5 mm long. The midpiece is 3 mm long and not as deeply marked as in Ostrich. The principal piece is ca 47 mm and the endpiece 3 mm long. Compared to Ostrich, the spermatozoon of Emu has a longer tail and a shorter head, while the midpiece (i.e. the distal centriole) is the same length (Baccetti et al. 1991). TEM sections of the spermatozoon are shown in Fig. 8.7D-I. Fig. 8.7 Dromaius novaehollandiae, Emu, spermatozoa. A-C. Scanning electron micrographs. D-I. Transmission electron micrographs. A Whole spermatozoon. ¥ 2200. B. Enlarged view excepting posterior flagellum. ¥ 4400. The head is shorter, the midpiece equal in length, and the tail longer than in Ostrich. C. Posterior end, showing the endpiece. ¥ 19000. D. Acrosome vesicle (A) on the anterior tapering rostrum of the nucleus (N). ¥ 49600. E. Longitudinal section (LS) of the posterior nucleus and the midpiece. ¥ 63600. F. LS midpiece and anterior principal piece. ¥ 33400. G. Transverse section (TS) of the acrosome vesicle and nuclear rostrum. ¥ 47700. As in A, note the absence of an endonuclear canal and perforatorium. H. TS of the principal piece showing the fibrous sheath. ¥ 44200. I. TS endpiece; note absence of fibrous sheath. ¥ 45000. A, acrosome vesicle; AF, accessory fibers; AX, axoneme; dc, distal centriole; FS, fibrous sheath; M, mitochondria; N, nucleus; Modified after Baccetti, B., Burrini, A. G. and Falchetti, E. 1991. Biology of the Cell (Paris) 71(1-2): 209-216. Figs. 3-5, 7, 12, 13, 9, 21, 20.

Avian Spermatozoa: Structure and Phylogeny

Fig. 8.7

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!%$ Reproductive Biology and Phylogeny of Birds Acrosome. Examined in sections (Fig. 8.7D, G) by transmission electron microscopy (TEM), the acrosomal complex is made up of a truncate-conical, 100 nm thick acrosome vesicle, which holds the apical tapered portion of the nucleus, and of an extremely thin bundle of microfilaments evident in the apical subacrosomal space (Fig. 8.7D). This space is extremely reduced if compared to that of Ostrich, and no endonuclear canal and no rod-like subacrosomal structure are evident. Therefore, the perforatorium, represented only by the sparse extranuclear microfilaments, is almost absent. Nucleus. The nucleus, which is strongly condensed, is cylindrical and similar to that of Ostrich. Centrioles and midpiece. The same is true for the centrioles, while the mitochondria are smaller, spheroidal, 5-6 in a cross-section of the midpiece, numbering 8-10 in longitudinal section (Fig. 8.7E) and reach a total number of 40 or more. They are closely juxtaposed, and the intermitochondrial cement is sparse. This organization explains the reduced demarcation of the midpiece, seen from the exterior by SEM. No penetration of the distal centriole by the two axonemal singlets is apparent in the micrograph (Fig. 8.7E) but absence perhaps requires confirmation. Tail. The axoneme has the same characteristics as in Ostrich, but the region containing the small accessory dense fibers (Fig. 8.7H) is even shorter, and the fibrous sheath (Fig. 8.7H) is progressively thinner toward the end part of the tail and is, by definition, lacking from the endpiece (Fig. 8.7I). The tail is longer than in Ostrich, and contains the basic 9 + 2 axoneme (Baccetti et al. 1991). Remarks. The spermatozoon of Emu resembles that of Ostrich and Rhea in most respects but shows three remarkable departures. Two of these, the absence of an endonuclear canal and contained perforatorium, are clearly correlated. The third is the rounded form of the apex of the acrosome vesicle. Loss of the perforatorium is enigmatic if, as the author hypothesizes, it contributes to the acrosome reaction at fertilization. Soley (1993) remarks that no definite function has yet been ascribed to the avian perforatorium and states that it would appear merely to represent a residual structure. However, Campanella et al. (1979) found that the turkey perforatorium consisted of actin and Baccetti et al. (1980) believed that it supported the conical shape of the acrosome. A role in the acrosome reaction and fertilization has been shown for the perforatorium which lies in an endonuclear canal penetrating most of the nucleus in the lamprey. In Lampetra fluviatilis, the central fiber (putative perforatorium) is capable of extrusion as a 50 µm long ‘head filament’ (Afzelius and Murray 1957; Kille 1960). In the acrosome reaction, the plasma membrane is drawn out into a slender sheath containing the central fiber (Stanley 1967). The putative perforatorium undergoes no observable change on extrusion (Follenius 1965). In L. planeri sperm in the egg coatings show a true acrosome reaction in which the central fiber is extended in an acrosomal tubule which penetrates the egg envelopes to reach the egg surface (Nicander

Avian Spermatozoa: Structure and Phylogeny

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and Sjödén 1968, 1971). In the hagfish Eptatretus an acrosome reaction occurs with formation of an acrosomal process with a filamentous core and is deduced to involve polymerization of actin (Morisawa and Cherr 2002). Analysis of proteins in the rat perforatorium, which does not form an acrosome process, failed to detect actin (Oko et al. 1990). It is thus difficult to accept a merely residual status for the avian perforatorium. On the other hand, even if it is involved in an acrosome reaction in Ostrich and Rhea, which has yet to be demonstrated, it may also have a supportive function for the conical acrosome vesicle as Baccetti et al. (1980) suggested and its loss in Emu could conceivably be related to the rounded form of the vesicle. It is clear that in Emu, as in many other birds, e.g. pigeons and passerines, similarly lacking a perforatorium, an acrosome reaction must nevertheless occur.

8.6.2

Order Tinamiformes

The Tinamiformes contain only the family Tinamidae, the Tinamous, with 9 genera and 48 species.

8.6.2.1

Eudromia elegans

The spermatozoon of Crested tinamou has been described by Asa et al. (1986) and Asa and Phillips (1987). Their accounts are paraphrased and augmented here. The sperm head. The head of Tinamou spermatozoon is cylindrical and slightly curved. The smooth, tapered anterior-most end is characterized by a small bump at the tip (Fig. 8.8A). Acrosome. Although the form of acrosome vesicle is undescribed, it is seen in the illustration to be a hollow cone, the inner and outer membranes of which are almost in contact at the tip and are more widely separate at about midlength than elsewhere. The vesicle contents appear homogeneous and moderately electron-dense. In thin sections, a small space is observed between the cell membrane and the outer acrosomal membrane (Fig. 8.8B). Nucleus. The nucleus (Fig. 8.8A) is a moderately elongate cylinder. In contrast with the acrosomal region, the cell membrane appears to be closely associated with the nuclear membrane in the region immediately posterior to the acrosome. The chromatin rarely appears completely condensed. In oblique or longitudinal sections, at the edge of the nucleus thick strands of chromatin are seen disposed obliquely to the long axis of the nucleus. This arrangement suggests that the chromatin spirals around the endonuclear canal. Endonuclear canal and perforatorium. Transverse and longitudinal sections of the nucleus reveal a tube-like structure (endonuclear canal) which extends from one end of the nucleus to the other and abuts the tip of the acrosome (Fig. 8.8B). Neck Region and Midpiece. The posterior edge of the sperm nucleus is indented as a concave disk. A short proximal centriole is situated near the

!%& Reproductive Biology and Phylogeny of Birds

Fig. 8.8 Eudromia elegans elegans, Crested tinamou. A. Scanning electron micrograph of the anterior portion of a spermatozoon. About 20 mitochondria are Fig. 8.8 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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base of the nucleus (Figs. 8.8C, D, 8.9A). Dense columns of the neck piece surround this centriole and extend into the distal centriole for, apparently, about half of its length. The distal centriole is 3 mm in length, the entire length of the midpiece, and is embedded in dense material (Figs. 8.8C, 8.9A). The mitochondria are roughly spherical. The midpiece contains about 20 mitochondria arranged in seven tiers with about five around the centriole (Figs. 8.8C, D, 8.9A). Flocculent material is observed between the mitochondria (Figs. 8.8C, D, 8.9A) (Asa et al. 1986). Annulus and principal piece. A distinct annulus is situated posterior to the short midpiece (Figs. 8.8C, 8.9), as in mammalian spermatozoa (Fawcett, 1975). Asa et al. (1986) state that no dense fibers were observed in tinamou spermatozoa, either in the midpiece or the principal-piece. However, they later state (Asa and Phillips 1987) that dense fibers are present in the proximal principal piece to which they are restricted. Periodic structures are observed in longitudinal sections of the fibrous sheath (Figs. 8.8C, 8.9C). In transverse section, the fibrous sheath shows the typical ribs opposite dense fibers 3 and 8. Material which appears morphologically similar to glycogen surrounds the fibrous sheath in the anterior region (Fig. 8.9C) (Asa et al. 1986). Tinamou sperm and ratite phylogeny. Spermatozoal characters have been considered equivocal as to whether the Tinamiformes or Struthioniformes are the most primitive (Soley 1993) or, in other words, if they are sister-groups, which is the apomorph sister-group. Asa et al. (1986) observe that Tinamou is exceptional among avian species in the great length of what is here termed the endonuclear canal, extending to the base of the nucleus. They considered it possible that the contents of the canal possess actin which can polymerize to cause the acrosome to be propelled through egg investments during the acrosome reaction as shown for other groups.

Fig. 8.8 Contd. ...

observed in the midpiece. ¥ 6800. B. Longitudinal section of the anterior portion of the sperm head. The contents of the acrosome vesicle and moderately electron dense. A perforatorium lying in an endonuclear canal runs through the center of the nucleus where it abuts the tip of the acrosome vesicle. ¥ 32500. C. Midsagittal section through the midpiece. The short proximal centriole and the long distal centriole are each seen in longitudinal section. The distal centriole, surrounded by spheroidal mitochondria, extends the entire length of the midpiece as is crocodile and other ratite sperm. The posterior end of the nucleus has a concave circular fossa. ¥ 22000. D. Transverse sections through the midpiece. The distal centriole, embedded in electron dense material, is observed throughout the length of the midpiece. Flocculent material is interspersed between some mitochondria. ¥ 53000. Av, acrosome vesicle; ax, axoneme; dc, distal centriole; m, mitochondrion; nr, nuclear rostrum; pc, proximal centriole; Adapted and relabeled after Asa, C., Phillips, D. M. and Stover, J. 1986. Journal of Ultrastructure and Molecular Structure Research. 1986; 94(2): 170-175, Figs. 1, 2, 5-7. With permission from Elsevier.

!& Reproductive Biology and Phylogeny of Birds

Fig. 8.9 A. Longitudinal section (LS) showing the midpiece of a tinamou spermatozoon. A, annulus; M, mitochondrion; F, fibrous sheath; G, glycogen; p, proximal centriole. ¥ 17400. B. LS showing portions of three spermatozoa of Thryothorus ludovicanus, the Carolina wren. The mitochondria (M) which spiral around the flagellum (FLA) have been referred to as the undulating membrane. Fig. 8.9 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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They also considered the Tinamou spermatozoon to be unusual in that there is a single distal centriole which extends the entire length of the midpiece. However, this has also been described for Struthioniformes (see Section 8.6.1), and for chelonians, and Sphenodon (Healy and Jamieson 1992; Jamieson and Healy 1992) and therefore appears to be a symplesiomorphy of Palaeognathae. Asa et al. (1986) observed that other birds such as fowl and ducks also have long distal centrioles, but that the midpiece of spermatozoa of these species is longer and the centriole only occupies the anterior portion of the midpiece. However, it is here noted that in addition to being shorter than the midpiece, the absolute length of the distal centriole in these is less than that in Struthioniformes and Tinamiformes. The presence of glycogen, around the fibrous sheath, appears to be unique to the Tinamou spermatozoon and therefore an autapomorphy insofar as tinamiforms have been studied. Soley (1993) reasonably considered that there is a trend to enlargement of axonemal dense fibers in birds. He therefore considered Tinamou to be the most primitive ratite in this respect in the erroneous belief that it lacked dense fibers in contrast with the rudimentary dense fibers observed in the proximal segment of the principal piece of Ostrich and Rhea (Phillips and Asa 1989) sperm. Therefore consideration of dense fibers does not contribute to phylogenetic analysis within the ratites. Furthermore, although (Soley 1993) rightly recognizes a trend to reduction in the length of the endonuclear canal and contained perforatorium in the non-passerines, with their loss in passerines, it is by no means certain that the fact that these structures are longer in tinamou than in struthioniforms indicates that Tinamou is more primitive. Doubt is cast on the latter inference by the fact that the canals and perforatoria are shorter in Crocodylia than in struthioniforms. As Soley (1993) argues, Ostrich and Rhea sperm appear to be more primitive than those of Tinamou in respect of the structure of the distal centriole. In these birds a central pair of microtubules occupies the centriolar cavity, a feature which is typical of chelonian (Furieri 1970; Hess et al. 1991; Healy and Jamieson 1992) and, we may add, crocodilian sperm (Jamieson et al. 1997). Tinamou sperm (and, apparently, see Section 8.6.1.5, Emu sperm) are said to lack this microtubular arrangement and to display an empty distal centriole (Asa et al. 1986; Asa and Phillips 1987) as in other non-passerine birds. However, central singlets appear to reach almost halfway into the Fig. 8.9 Contd. ...

A, acrosome; AP, lateral projection of acrosome; D, dense fibers; ED, electrondense material of the neck; N, nucleus. ¥ 17400. C. Transverse section (TS) through the principal piece of tinamou spermatozoa. E, endpiece; F, fibrous sheath; G, glycogen. ¥ 48700. D. TS of spermatozoa from the vas deferens of the Carolina wren. Viewed in cross section the dense fibers are approximately circular and are similar to one another. M, mitochondria. ¥ 45900. After Asa, C. S. and Phillips, D. M. 1987. Pp. 365-373. In H. Mohri (ed). New horizons in sperm cell research, Japan Science Society Press, Gordon and Breach Scientific Publications, Tokyo/New York, Figs. 1-4.

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Reproductive Biology and Phylogeny of Birds

Fig. 8.10 Drawings by light microscopy of spermatozoa of non-passerine birds. Rooster (Gallus gallus); Tufted duck (Aythya fuligula); Domestic duck (Anas platyrhynchos); Common guillemot (Uria aalge); Corncrake (Crex crex); Lesser black-backed gull (Larus fuscus); Alpine dunlin (Calidris alpine); Green sandpiper (Tringa ochropus); Lapwing (Vanellus vanellus); Woodcock (Scolopax rusticola); Common coot (Fulica atra). After Retzius, G. 1909. Biologische Untersuchungen, Neue Folge 14(10): 89-122 Taf XIX-XXXVII.

Avian Spermatozoa: Structure and Phylogeny

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centriole in Crested tinamou sperm (see Fig. 8.9A, above, from Asa and Phillips 1987). The phylogeny of ratites is discussed in Section 8.11.

8.7

NEOGNATHAE

According to Gauthier and de Queiroz (2001) “Neognathae” refers to the crown clade stemming from the last common ancestor of Charadrius pluvialis (Pluvialis apricaria) Linnaeus 1758 and all extant birds sharing a more recent common ancestor with that species than with Struthio camelus Linnaeus 1758 and Tetrao (Tinamus) major Gmelin 1789. Neognathae consists of two primary crown clades, Galloanserae and Neoaves.

8.8

GALLOANSERAE

This is the Parvclass Galloanserae of Sibley and Ahlquist (1990). Its monophyly versus paraphyly are discussed by Harshman in Chapter 1. Spermatozoal ultrastructure is consistent with monophyly. The ancestor of the Galliformes and Anseriformes was presumably a generalized form lacking the highly derived filter feeding apparatus of Anseriformes.

8.8.1 8.8.1.1

Order Galliformes Gallus gallus (=domesticus)

The spermatozoon of the rooster, Gallus gallus (= domesticus), Phasianidae, has been described ultrastructurally (Grigg and Hodge 1949; Bonadona 1954; Nagano 1960, 1962; McIntosh and Porter 1967; Nicander and Hillstrom 1967; Krustev and Danov 1968; Lake et al. 1968; Nicander 1970b; Tingari 1973; Bakst and Howarth 1975; Gunawardana and Scott 1977; Bakst and Sexton 1979; Bakst 1980; Xia et al. 1985; Xia et al. 1986; Bae and Kim 1987; Thurston and Hess 1987; Woolley and Brammall 1987; Sprando and Russell 1988; Xia et al. 1988; Jamieson 1999). The following account is based chiefly on a reexamination. Reference is made to other accounts where they are not in agreement or give additional or important supporting information. General morphology. The Gallus spermatozoon (Figs. 8.11, 8.12, 8.16J, P, R) has the usual non-passerine components in anterior-posterior sequence: an acrosome vesicle; perforatorium (apical spine of Grigg and Hodge 1949); acrosome spine of Lake et al. 1968; acrosomal spine of Tingari 1973) lying in the subacrosomal space and extending posteriorly into an endonuclear canal, here in the form of a deep anterior nuclear fossa; elongate nucleus; proximal and distal centrioles; midpiece, consisting of mitochondria encircling the 9+2 axoneme and ending posteriorly at the annulus; principal piece, consisting of the axoneme surrounded by a fibrous, here amorphous, sheath; and the endpiece, consisting of the axoneme surrounded by the plasma membrane but lacking a fibrous sheath. The endpiece contains a central dense ‘tip granule’

!&" Reproductive Biology and Phylogeny of Birds

Fig. 8.11 Gallus gallus. Transmission electron micrographs. A. Longitudinal section (LS) of the acrosome and adjacent nucleus, showing perforatorium in the subacrosomal space and extending into the nuclear fossa (endonuclear canal). B. LS basal portion of nucleus, midpiece and anterior portion of principal piece. C. LS Fig. 8.11 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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(terminology of Woolley 1995 for Coturnix), also observed in Gallus by Woolley and Brammall (1987). Grigg and Hodge (1949) give a total length for the sperm “head” (nucleus but contrary to usual definitions apparently excluding the acrosome) of 14 µm and a diameter not exceeding 0.5 µm, and a length for the sperm of more than 100 µm, c.f. 90 µm or more (Thurston and Hess 1987). Acrosome. The acrosome vesicle is approximately 2.2 µm long (present study), agreeing with 2 µm or greater (Thurston and Hess 1987). It has the form of an elongate cone, asymmetry of which correlates with a slightly curved shape. The base of the vesicle overlaps a short, abruptly narrowed anterior region of the nucleus (Figs. 8.11A, C, 8.12A). In the large, conical subacrosomal space which occupies a little more than the basal third of the acrosome, there lies a dense rod, the perforatorium, ca 1.2 µm long (1.0 µm, Thurston and Hess 1987), the tip of which closely abuts the apex of the subacrosomal space. Approximately the posterior fourth of the perforatorium is contained in, and closely fits, the anterior nuclear fossa which is here considered the homologue of the ratite endonuclear canal. Amorphous material in the subacrosomal space is said to be continuous with the perforatorium and to be part of it by Xia et al. (1988). This condition is seen in the present study in Fig. 8.11C but may represent a dissolution of the perforatorium as the latter is a distinct rod in the absence of amorphous material, except that closely ensheathing the rod, in Fig. 8.11A. Micrographs of cross sections at the base of the perforatorium of tannic acid-fixed rooster sperm demonstrate the plasmalemma, inner and outer acrosomal, and double nuclear membranes (Thurston and Hess 1987) (Fig. 8.16J). Nucleus. The nucleus is an elongate cylinder, slightly tapering anteriad, with a slight shoulder supporting the base of the acrosome vesicle (Fig. 8.11A, B, C). It is slightly curved, at least in fixation. The chromatin is strongly condensed and electron-dense. At its base there is a very shallow, slightly asymmetrical basal (implantation) fossa. Centrioles. The short proximal centriole lies at right angles to the long axis of the spermatozoon, with its anterior region in the shallow implantation fossa (Fig. 8.11B). It consists of 9 triplets embedded in a dense ring. It is linked by an unstriated connecting piece to the base of the nucleus (Bakst and Howarth 1975), termed the non-striated connecting piece and considered to include the proximal centriole by Thurston and Hess (1987). This is termed the capitulum Fig. 8.11 Contd. ...

base of acrosome vesicle, perforatorium and anterior nuclear fossa. D. Transverse section (TS) through anterior nuclear fossa, showing enclosed perforatorium. E. TS nucleus, showing circular profile. F. TS midpiece, showing four mitochondria and the axoneme with nine outer dense fibers. G. TS principal piece, distinguished by presence of a fibrous (amorphous) sheath around the axoneme. H. TS endpiece, distinguished by the axoneme lacking a fibrous sheath. I. TS terminal region of endpiece, showing doublets undergoing disruption and central singlets replaced by a dense ‘tip granule’. Original.

!&$ Reproductive Biology and Phylogeny of Birds

Fig. 8.12 Gallus domesticus. A. Longitudinal section of the acrosomal region. The acrosomal vesicle overlaps a perforatorium which inserts into a nuclear concavity. The perforatorium is not membrane bound and is surrounded by amorphous, granular material. B. Longitudinal section of the centrioles and midpiece, the proximal centriole (PC) is orientated at right angles to the distal centriole (DC) which is surrounded by mitochondria (M). Bars: 0.1 µm. From Thurston, R. J. and Hess, R. A. 1987. Scanning Microscopy 1: 1829-1839, Figs. 2b, 4b.

by Tingari (1973) who states that it forms in the epididymis by fusion of previously existing dense masses and that in the excurrent ducts the reduction in size of the central cavity of the proximal centriole may be due to deposition of dense matrix. The distal centriole, contiguous with the proximal centriole, lies in the longitudinal axis of the sperm and is continuous with the axoneme. It is surrounded by the most anterior mitochondria of the midpiece. Unlike the condition in struthioniforms, the central singlets do not extend to the proximal end of the centriole. It is considered that these commence at its posterior end (Nagano 1962; Bakst and Howarth 1975; Gunawardana and Scott 1977) but some micrographs suggest a considerable penetration of the singlets into the distal end of the centriole (Fig. 8.12B). From a micrograph of Bakst and Howarth (1975) the length of the distal centriole is ca 1.8 µm and it extends for ca 0.4 of the length of the midpiece. It has nine triplets embedded in a ring of dense material (Lake et al. 1968; Bakst and Howarth 1975; present study). The distance from the proximal end of the distal centriole to the commencement of the inner paired microtubules is 0.9 µm (Thurston and Hess 1987).

Avian Spermatozoa: Structure and Phylogeny

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Tannic acid fixation reveals for the proximal (Fig. 8.16J) and distal centrioles 13 protofilaments for subtubule A and 10 for each of B and C (Thurston and Hess 1987). In the round spermatid the two centrioles are said to be of the same size and to lie end to end and almost in a straight line (Nagano 1962), the condition seen in the mature Coturnix sperm. However, Xia et al. (1986) clearly indicate a small proximal centriole at right angles to a long distal centriole in the round spermatid, as confirmed here for the spermatozoon in Fig. 8.11B. Midpiece. The midpiece is ca. 3.7 µm long (agreeing with 4 µm, Grigg and Hodge 1949). It shows, in transverse section, four mitochondria encircling the axoneme (Fig. 8.11F) and 7 or 8 along its length (Fig. 8.11B), totaling ca 28-32, agreeing with approximately 30 according to Bakst and Howarth (1975). Their arrangement is helical (Lake et al. 1968; Bakst and Howarth 1975; Thurston and Hess 1987). The cristae form stacks of plates orientated longitudinally (Figs. 8.11B, 8.12B). The mitochondria in tangential section appear as closely fitting slightly elongate polygons (usually hexagons) (Bakst and Howarth 1975). The axoneme in the midpiece, posterior to the distal centriole, has nine conspicuous dense fibers (osmiophilic masses of Bakst and Howarth 1975); each fiber being in the radius of its doublet and enveloping the latter in its inner extremity. The dense fibers do not extend into the principal piece (Lake et al. 1968; Bakst and Howarth 1975; present study). Annulus. A small, compact annulus marks the posterior limit of the midpiece (Fig. 8.11B). Principal piece. This commences at the annulus and is defined by the presence, encircling the axoneme, of a fibrous sheath (Figs. 8.11B, G, 8.16P). This is amorphous in that it does not show the annulation or ribbing seen in ratites. Endpiece. The endpiece (Figs. 8.11H, 8.16R) consists of the axoneme and plasma membrane and lacks the fibrous sheath. As in the principal piece, the A subtubule of each of the 9 doublets has dense contents, as first noted by Nagano (1962), and the outer dynein arms are more conspicuous than the inner arms. Posteriorly the two central singlets are replaced by a large dense structure, the so-called ‘tip granule’ (Fig. 8.11I). At this level and posteriorly the doublets are progressively disrupted. Location of the tip granule at the end of the two central singlets and not terminally is deduced from the observations of Woolley (1995) for Coturnix coturnix.

8.8.1.2Coturnix japonica In the present account the valid name Coturnix japonica is used in place of Coturnix coturnix, Coturnix coturnix japonica and Coturnix coturnix var. japonica employed in the various accounts summarized. The spermatozoon of C. japonica (Phasianidae) has been described by Marquez and Ogasawara (1975); Saita et al. (1980); Maretta et al. (1982); Lin and Jones (1993) and Woolley (1995). The dynamics of spermatozoal motility are described and illustrated

!&& Reproductive Biology and Phylogeny of Birds

Fig. 8.13 Coturnix japonica. Light microscopy of spermatozoon. A. Immotile spermatozoon, wet preparation. Upper arrow indicates the midpiece-principal piece boundary. Lower arrow, the distal limit of the principal piece and its sheath. The long unsheathed more posterior region is here termed the endpiece. B. Flagellum after trypsin digestion, showing helical midpiece. C. The acrosome, nucleus, and proximal flagellum contrasted supravitally with eosin-yellow. After Woolley, D. M. 1995. Acta Zoologica (Copenhagen) 76: 45-50, Figs. 1-3.

Avian Spermatozoa: Structure and Phylogeny

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by Vernon and Woolley (1999) who repeat the ultrastructural data of Woolley (1995). The following account is drawn chiefly from Woolley (1995). General morphology. Eosin-yellow in vivo staining of the spermatozoon reveals a pointed conical acrosome many times shorter than the cylindrical nucleus (Fig. 8.13C). Light micrographs of the flagellum revealed three zones of decreasing thickness, later confirmed to be the midpiece, the proximal (sheathed) principal piece and the “distal principal piece” (endpiece) (Fig. 8.13A). The mean overall length of the flagellum was 207.6 mm. The mean lengths of the three zones (± one standard deviation) were: midpiece 161.4 (± 2.8) mm, proximal (sheathed) principal piece 5.4 (± 0.7) mm and endpiece 40.8 (± 1.7) mm (n = 10, a single bird). Acrosome. The acrosome is conical, about 2.6 mm (Fig. 8.13C). The mode of attachment of the acrosome (Fig. 8.14A) involves an overlapping joint, as is usual in galliforms, with a perforatorium (length 1.5 mm) engaged in conical depressions in both the acrosome and the nucleus, and connected to each through a granular matrix. Nucleus. The sperm nucleus is a curved cylinder, 20.6 mm long. The nuclear envelope is thickened where it lies against the inner acrosomal membrane. Centriolar region. The neck of the spermatozoon contains two separate centrioles lying almost on the same axis. The proximal centriole is thickened abaxially to support an implantation plate; distally it ends in an annular thickening into which the base of the longer distal centriole is anchored (Fig. 8.14F-H). The triplet microtubules are not continuous between the two centrioles (Fig. 8.14F). Beyond the distal centriole there is a transition region characterized by the appearance of the central pair, by electron dense material peripheral to the nine doublets and by additional densities associated with the radial spokes, showing their 96 nm periodicity (Fig. 8.14F, I). The transition region is within the mitochondrial sheath. Midpiece. Each mitochondrion is disc-shaped, diameter about 0.25 mm, and curved against the axoneme, with the packing of adjacent ones often making the profiles slightly hexagonal (Fig. 8.14J). Transverse sections typically show four mitochondria at any level (Fig. 8.14K). The arrangement can be modeled as four parallel ‘out-of-register’ chains of mitochondria surrounding the axoneme. The total number of mitochondria per sperm is estimated as ca 2,500. The midpiece terminates at a thin annulus (Fig. 8.14M). Principal piece. The short “proximal principal piece” consists of the axoneme encased in a cylindrical fibrous sheath that tapers distally (Fig. 8.14N). Beyond this sheath, the axoneme is simple (Fig. 8.14O). Although the fibrous sheath is amorphous in the mature sperm, it develops as a series of circumferentially orientated hoops in the stage 2 spermatid (Lin and Jones 1993). Endpiece. The endpiece exceeds 1.5 mm in length. In it the central pair of microtubules terminates first. Posterior to this, for about 0.4 mm, the center of

!' Reproductive Biology and Phylogeny of Birds

Fig. 8.14 Coturnix japonica. TEM of sperm. A. Longitudinal section (LS) through the junction between the acrosome vesicle (av) and the sperm nucleus (n). The perforatorium (p) inserts into each structure. B. Transverse section (TS) of nucleus, Fig. 8.14 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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the axoneme is occupied, as in Gallus, by an electron dense body tip granule (Fig. 8.14L, P). Thereafter the nine doublets become progressively simplified and end as singlets attached to the plasmalemma (Fig. 8.14Q, R) (Woolley 1995). Remarks. Woolley (1995) gives an interesting discussion of the sperm of Coturnix coturnix, some points of which will be discussed here. The morphology of the acrosome and sperm nucleus is typical of galliform birds. Furthermore a conical acrosome, with a perforatorium occurs in most of the other non-passerine orders that have been examined ultrastructurally (reviewed by Asa and Phillips 1987). As Woolley (1995) notes, the ultrastructure of the sperm neck in Coturnix has an unusual feature. In spermatozoa generally, there are two centrioles (at least, during development): a distal centriole that gives rise to the flagellum and a proximal one at right angles to it. This arrangement is also general in birds. In Coturnix japonica, however, the two centrioles lie almost in line. This condition is also seen in C. chinensis (8.8.1.3) and Numida meleagris (8.8.1.6). As also noted by Woolley (1995), the general features of the flagellum in Coturnix japonica spermatozoa are qualitatively typical of non-passerine birds. Its dimensions, however, are unusual. First, at 208 mm, the flagellum is more than twice as long as in the other galliforms noted. In passerine bird sperm the flagella range up to 263 mm in length, in Dendroica petechia, the yellow warbler (Briskie and Montgomerie 1992). Flagellar length is useful taxonomically within avian genera, as shown for Dendroica by McFarlane (1963). The significance, taxonomic or physiological, of the flagellar

Fig. 8.14 Contd. ...

which has a circular profile at all levels. C. TS of the region where acrosome and nucleus are interlocked. D. TS of caudal acrosome, showing the perforatorium centrally. E. TS of rostral acrosome. F. LS through the neck region, which consists of a proximal centriole (pc), a distal centriole (dc) and a transitional region with periodic densities suggestive of mechanical re-enforcement. G. TS through one of the centrioles, probably the distal one. H. A further LS of the neck to show the separate identity of the centrioles, indicated by their lack of continuity and slightly different axes. I. TS of the transition region. The microtubular triplets have been reduced to doublets, a central singlet has appeared and there are extra densities both peripherally and centrally. J. Tangential section of the midpiece to show the shape and arrangement of the mitochondria. K. TS of midpiece showing the most common arrangement of the mitochondria. L. LS of the flagellar tip showing particularly the tip granule. M. LS showing annulus (arrows) at the distal limit of the midpiece. N. TS proximal principal piece, where fibrous sheath occurs between the axoneme and the cell membrane. O. TS distal principal piece. P. TS flagellum at the level just beyond the termination of the central pair, showing the central tip granule (tg). Q, R. TS showing progressive reduction of the axoneme in the flagellar tip. After Woolley, D. M. 1995. Acta Zoologica (Copenhagen) 76: 45-50, Figs. 4-21.

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Reproductive Biology and Phylogeny of Birds

elongation in Coturnix is unknown. However, the physiological explanation may lie in the positive correlation shown by Briskie and Montgomerie (1992) between sperm flagellar length in birds and the length of the sperm storage tubules in the female of the species. The length of the midpiece in Coturnix japonica is noteworthy (Woolley 1995) and requires confirmation. Thurston and Hess (1987) estimated that the turkey, the domestic fowl and the Guineafowl all have 25-30 mitochondria in each spermatozoon (confirmed here for Gallus). For Coturnix, Saita et al. (1980), using electron micrographs of testicular tissue, estimated ca 350 mitochondria per sperm. The estimate of midpiece length (161 mm) in Coturnix japonica (=Coturnix coturnix var. japonica) by Woolley (1995) was made from light micrographs, supported by the very high frequency of midpiece profiles seen in the thin sections. This length measurement led to an estimate of mitochondrial number of ca 2500 per sperm. This would suggest a surprising 616 tiers of mitochondria. The only report of a midpiece approaching this length in non-passerine sperm is for the Order Columbiformes. In contrast, in the Passerida the mitochondria make a single spiral thread along much of the axoneme (reviewed by Asa and Phillips 1987, and this chapter).

8.8.1.3 Coturnix chinensis General morphology. The ultrastructure of the spermatozoon of Coturnix chinensis, the Blue-breasted quail (Fig. 8.15A-L) is closely similar, excepting some dimensions, to that of Gallus and, particularly, of Coturnix japonica. It has the usual non-passerine components described above for Gallus. By light microscopy, approximate dimensions are: total length of the spermatozoon 91 µm; head (acrosome + nucleus) 9.7 µm; midpiece 24.4 µm (n=1). Acrosome. The acrosome vesicle, by TEM, is approximately 1.9 µm long (n=2). It has the form of an elongate, straight cone. The base of the vesicle overlaps a short, narrower anterior region of the nucleus (Figs. 8.15C, E). The nuclear shoulders are rounded and not as distinct as in Gallus. In the large, conical subacrosomal space which occupies more than half the length of the acrosome vesicle, there lies a dense rod, the perforatorium, ca 1.3 µm long. This agrees with C. japonica and Gallus in abutting the apex of the subacrosomal space but differs from that of Gallus in more closely fitting the sides of the space, there being a relatively thin later of granular material between it and the vesicle laterally. Approximately the posterior third of the perforatorium is contained in, and closely fits, the anterior nuclear fossa (reduced endonuclear canal). Nucleus. The nucleus is an elongate cylinder, ca 6.4 µm long (cf 20.6 µm for C. japonica), circular in cross section (Fig. 8.15B), slightly narrowing anteriad, but conspicuously clubbed basally on one side (Fig. 8.15C). Seen in the plane of the clubbing (Fig. 8.15C, F) the nucleus is tilted at an angle to the long axis of the midpiece and flagellum. In a plane at right angles to this (Fig. 8.15G) clubbing is absent and the nucleus appears to be in the long axis. The chromatin is homogeneous and electron-dense (Figs. 8.15C, F, G, H). Where it

Avian Spermatozoa: Structure and Phylogeny

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appears granular (Fig. 8.15E) this may be due to incomplete maturity, as a transition from granular to condensed is noted by Maretta et al. (1982) for C. japonica. At its base there is a very considerable, slightly asymmetrical basal (implantation) fossa (Figs. 8.15F, G, H). As in C. japonica, the extreme tip of the nucleus is attenuated around the anterior half of the anterior fossa (Fig. 8.15E), being thinner on each side than in Gallus, and is asymmetrical so that in transverse section it may appear interrupted on one side (Fig. 8.15A). Centriolar region. The neck of the spermatozoon contains two separate centrioles lying almost on the same axis (Fig. 8.15G, D), as in C. japonica. The triplet microtubules are not continuous between the two centrioles. The posterior limit of the distal centriole is difficult to define but appears to be three or four times the length of the proximal centriole. It is penetrated for about half of its length by a central axonemal element. This consists of a dense sheath around and capping the two central singlets (Fig. 8.15G). At the level of the tip of this sheath, the nine triplets of the centriole lie in a dense ring and their triplet structure is almost obscured though persistent (Fig. 8.15I). The ring is surrounded by the mitochondria of the midpiece. Midpiece. Transverse sections of the midpiece typically show four mitochondria at any level (Fig. 8.15I, J). The length of the midpiece has not been determined by TEM, nor has its posterior limit been sectioned. Small outer dense fibers, one in the radius of and contiguous with, each doublet are present in the midpiece (Fig. 8.15J); in some sections they are considerably larger than those shown in this figure. Principal piece. This is defined by the presence, encircling the axoneme, of a fibrous sheath (Fig. 8.15K). This is amorphous in that it does not show the annulation or ribbing seen in ratites. Endpiece. The endpiece (Fig. 8.15L) consists of the axoneme and plasma membrane and lacks the fibrous sheath. As in the principal piece, the A subtubule of each of the 9 doublets has dense contents. A tip granule has not been demonstrated in the few sections obtained. Remarks. The difference in length of the nucleus between Coturnix coturnix and C. chinensis, 20.6 µm versus 6.4 µm is remarkable. Even more disparate is the midpiece length of 161 µm contrasting with approximately 24 µm determined by light microscopy for C. chinensis. The two species agree, and differ from Gallus, in the much greater length of the midpiece, and in the almost co-linear arrangement of the proximal and distal centrioles, the proximal centriole being at right angles to the distal in Gallus as in other birds studied, with the exception of Numida meleagris. The clubbing and angular deflection of the nucleus in C. chinensis is echoed in a light micrograph by Woolley (1995) of the sperm of C. japonica (Fig. 8.13A) but may reflect a dynamic situation as it is not evident in a second micrograph (Fig. 8.13C).

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Fig. 8.15 Coturnix chinensis. TEM of sperm. A. Transverse section (TS) acrosome through the tip of the nucleus, showing asymmetry of the anterior nuclear fossa containing the base of the perforatorium. B. TS nucleus showing circular profile. Fig. 8.15 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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Fig. 8.15 Fig. 8.15 Contd. ...

C. LS of the entire acrosome and nucleus, with centriolar and anterior midpiece. D. Detail of centriolar region, showing proximal centriole almost in line with the distal Fig. 8.15 Contd. ...

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8.8.1.4 Meleagris gallopavo The spermatozoon of the turkey, Meleagris gallapavo (Phasianidae), has been described ultrastructurally (Marquez and Ogasawara 1975; Bakst and Sexton 1979; Baccetti et al. 1980; Bakst 1980; Bradley et al. 1986; Thurston and Hess 1987). The account of Thurston and Hess (1987) gives a valuable comparison of the sperm of Turkey, Guineafowl and rooster and is summarized here for the turkey, with some reference to the other species and other accounts. General morphology. As shown by SEM, the general shape of turkey, Guineafowl (‘guinea’) and rooster spermatozoa is remarkably similar. As in Gallus, the spermatozoa are long and narrow with a vermiform appearance, and an acrosome, nucleus, midpiece (Fig. 8.16A). The nucleus is usually curved. The anterior end of the sperm consists of a conical acrosome which is most prominent in Guineafowl sperm (Fig. 8.16B). The acrosome of the turkey is 1.0-2.6 µm (mean 1.8) µm long (Marquez and Ogasawara 1975), compared to 2 µm or greater (Thurston and Hess 1987), reaching 2.5 µm (Marquez and Ogasawara 1975) for the rooster acrosome. The nucleus gradually increases in diameter from its junction with the acrosome to its distal end at the beginning of the midpiece. The turkey sperm nucleus is said to be shorter (7 to 9 µm) than that of Guineafowl or rooster (10 to 14 µm in length) (Thurston and Hess 1987). However, there is considerable overlap between species as Marquez and Ogasawara (1975) give a length of 7.2-11.0 µm, with a mean width of 0.8, for the turkey. Junction of the nucleus with the midpiece at the neck region is not as conspicuous as in Guineafowl sperm (Thurston and Hess 1987). The flagellum comprises most of the length of the spermatozoon, although the junction between the principal and end piece could not be discerned with SEM. Turkey and Guineafowl sperm flagella are usually 60-65 µm long (Thurston and Hess 1987) (61 µm, Marquez and Ogasawara 1975), cf often more than 70 µm in rooster spermatozoa. The overall sperm length of 75-80 µm in turkey and Guineafowl sperm is also less than that of rooster sperm (90 µm). For all three species, the spermatozoa increased in width from the acrosome to a maximum of 0.5-0.7 µm at the junction of the nucleus with the midpiece. The width then decreased to 0.1-0.2 µm at the end of the flagellum (Thurston and Hess 1987).

Fig. 8.15 Contd. ...

centriole. E. LS acrosome and adjacent nucleus. F. LS base of nucleus, centriolar region and anterior midpiece. G. Same, in a plane approximately at right angles. H. Same but showing some mitochondria around the nucleus. I. TS distal centriole, within the midpiece, and through dense sheath at commencement of central singlets. J. TS midpiece and contained axoneme showing minute outer dense fibers. K. TS principal piece, showing amorphous sheath surrounding axoneme. L. TS endpiece. Original.

Avian Spermatozoa: Structure and Phylogeny

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Acrosome. As seen by TEM, the membrane-bound cap-like acrosomal vesicle (Fig. 8.16C, D) contains a granular, amorphous material which surrounds the perforatorium, and adjacent to the perforatorium is fine, granular material of moderate density (more abundant in rooster spermatozoa. At its distal end, the acrosomal cap encircles projections of chromatin from the apical portion of the nucleus. At its posterior end the perforatorium inserts into a concavity of the nucleus and extends obliquely forward approximately half the length of the acrosomal cap in turkey as in rooster, contrasting with nearly the entire length of the cap in Guineafowl sperm. Thus, the perforatorium of the turkey and rooster, at 1.0 µm, is appreciably shorter than that of the Guineafowl (1.9 µm). The base of the perforatorium of turkey is narrower than that of rooster and Guineafowl sperm (Fig. 8.16A). The substance of the perforatorium is dense and amorphous, and often interrupted by lucent channels which contain granular material similar to that adjacent to the perforatorium (Fig. 8.16D, E). Nucleus. A longitudinal section of the base of the nucleus and anterior portion of the midpiece is shown in Fig. 8.16F. The nuclear chromatin is dense and granular with occasional small lucent areas giving it a mottled appearance (Fig. 8.16G). The distal end of the nucleus terminated in a concavity, the implantation fossa (Fig. 8.6F, H). Centrioles and neck. For turkey (Figs. 8.16F, H) and rooster sperm, dense processes extended radially from the proximal centriole wall to abut against the nuclear membrane in the implantation fossa. The centriole complex plus the projections constitute the non-striated connecting piece of the neck of the spermatozoon (Bakst and Howarth 1975). Turkey and rooster have a proximal centriole orientated perpendicular to the distal centrioles (contrast Guineafowl below). In Numida, as for all three species, cross sections of the centrioles have the typical ‘pinwheel’ arrangement of nine triplet microtubules embedded in a cylindrical, dense wall (Fig. 8.16H). Each projection of the non-striated connecting piece is associated with one set of the triplet microtubules (Fig. 8.16H). Midpiece. From SEM micrographs of turkey sperm (Fig. 8.16D) and those of midpiece cross sections (Fig. 8.16K) where the mitochondrial length varied progressively from long to short, it is ascertained (Thurston and Hess 1987) that the midpiece had 25-30 mitochondria arranged in an helical pattern. Marquez and Ogasawara (1975) observed lengths for the midpiece of 4.0-6.0 µm; plate-like mitochondria numbering four per turn surround the axoneme; this arrangement is repeated seven times to give approximately 28 mitochondria per midpiece. In surface view (Thurston and Hess 1987), the mitochondria are polygonal with the dimensions of approximately 0.8 ¥ 0.11 ¥ 0.3 µm. In contrast to Guineafowl, cristae of turkey and rooster sperm mitochondria are parallel to the outer membrane (Fig. 8.16F).

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Fig. 8.16 A-H, K-M, O, Q-T. Meleagris gallopavo. Turkey spermatozoa. I, N. Numida meleagris, Guineafowl. J, P, R. Gallus gallus, Rooster spermatozoa. A, B. SEM of turkey sperm. A. The narrow, vermiform shape of the turkey spermatozoon is typical Fig. 8.16 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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Stages in transition from the distal centriole caudally are as follows (Thurston and Hess 1987) (Fig. 8.16K-M). The distal centriole microtubules come to lie in a circle and the dense wall material disintegrates (Fig. 8.16K). Central, singlet microtubules then become visible (Fig. 8.16L). Further caudally, the 9 + 2 microtubular pattern of the axoneme is visible and then (Fig. 8.16M) vestiges of dense material remain with the A doublet microtubule and (not mentioned) dense fibers are present. The centriole has a lucent center mottled with sparse granular material which extends from the apical end of the centriole caudally to the origin of the inner paired microtubules (Fig. 8.16F) of the axoneme. This distance varied, being 2.2, 0.9 and 0.65 µm for turkey, rooster and Guineafowl sperm, respectively. Fig. 8.16 Contd. ...

for sperm of non-passerine birds. Apical tip = acrosome, N, nucleus, M, midpiece, F, flagellum. B. A damaged turkey sperm with a denuded midpiece demonstrates helical arrangement of mitochondria (M). N, nucleus. Bars: A 5 µm; Fig. B 0.5 µm. C. Acrosome vesicle (AV) ensheathing the perforatorium which projects into the short endonuclear canal. Bar: 0.1 µm. D, E. The perforatorium has lucent areas (L) containing granular material. Transverse section of a turkey sperm nucleus. Dense chromatin (NC) granules are surrounded by lucent areas. Bars: D 0.1 µm, E 0.05 µm. F. Anterior portion of the midpiece; the proximal centriole is at right angles to the long distal centriole. M: mitochondria. G. TS nucleus. Dense chromatin (NC) granules are surrounded by lucent areas. H. The non-striated connecting piece consists of dense projections from the proximal centriole (PC) in turkey sperm and I. what appears to be the distal centriole (DC) in Guineafowl sperm. One projection is associated with one set of the nine triplicate centriolar microtubules. Bars: F-H, I 0.1 µm. J. Triplet microtubules of the proximal centriole of rooster sperm fixed with tannic acid. The tubulin protofilaments are visible: 13 for microtubule A, 10 for B and C. Bar: 0.05 µm. K-M. Turkey sperm midpiece at progressively distal levels. K. The distal centriole microtubules become circular and the dense wall material disintegrates (DC lumen of the distal centriole). L. Central, singlet microtubules then become visible (arrow). M. Further caudally, appearance of the 9 + 2 microtubular pattern of the axoneme. Vestiges of dense material remain with the A doublet microtubule (arrow). M – mitochondria. N. Annulus (arrows) at the termination of the Guineafowl sperm midpiece and beginning of the flagellum. Bars in K-N 0.1 µm. O-R. Figs. O and Q represent turkey sperm flagella fixed while motile. The axonemal doublets have a complete, dense A (A) microtubule connected to dynein arms (D) and radial links (R), and an incomplete, lucent B microtubule (B). The outer amorphous sheath (Fig. O) disappears in the distal end of the flagellum (Fig. Q). The inner matrix is coalesced in rooster flagella fixed after being immobilized by hypertonicity (Fig. P), obliterating the radial links, but the double microtubules, with 73 protofilaments in A and 10 in B, are intact (Fig. R, A and B; see also inset above). Bars: O-R 0.05 µm. S, T. LS of turkey sperm flagellum. S. The central tubules are bridged by material spaced 12 nm (arrow). T. Axonemal microtubules extend to the end of the flagellum. Bars 0.1 µm. Adapted from Thurston, R. J. and Hess, R. A. 1987. Scanning Microscopy 1: 1829-1839, Figs. 1a, d; 2a; 3a, b; 4a; 5a-c; 7a-d; 8a-d; 9a, b.

" Reproductive Biology and Phylogeny of Birds Cross sections of the flagella showed typical 9 + 2 microtubular axonemes (Fig. 8.16O). As in other galliforms, the A microtubule of the outer doublets is completely circular and filled with dense material. Surrounding the outer doublets is an amorphous sheath (Lake et al. 1968), defining the principal piece. Endpiece. The portion of a flagellum in which the cell membrane is in juxtaposition to the doublet microtubules, in the absence of the amorphous sheath, is the end piece (Fig. 8.16Q). The dense A microtubule of the doublet extends uninterrupted along the length of the flagellum (Fig. 8.16S, T), but doublet microtubules probably become single near the end of the flagellum as shown for rooster. Remarks. The structure of the sperm of the turkey is typical of the Galloanserae.

8.8.1.5

Tragopan caboti

The spermatozoon of Cabot’s tragopan, Tragopan caboti (Phasianidae) has been described ultrastructurally by Wen et al. (1997). The following account is augmented by examination of their micrographs. General morphology. The ultrastructure of the spermatozoon of Cabot’s tragopan is similar to that of the rooster. Acrosome. The conical acrosome is located in front of the nucleus, its caudal part slightly overlaps the anterior end of the nucleus. The perforatorium (‘acrosomal spine’) lies in the subacrosomal space with its base in the anterior nuclear fossa (‘nuclear pocket’). It is not homogeneous in electron density. Nucleus. The nucleus narrows abruptly for a short length anteriorly and forms shoulders supporting the base of the acrosome vesicle. It is a slender cylinder widened basally and has a length of ca 12 µm. The implantation fossa is occupied by the centriolar complex and non-striated connecting piece, which makes up the neck of the spermatozoon. Centrioles. The short proximal centriole is at right angles to the long axis of the sperm. The distal centriole, contiguous with it, is in the long axis and is elongate, being ca 1.3 µm long, approximately half the length of the midpiece. It continues as the axoneme. It is not penetrated by the central axonemal singlets. Midpiece. In transverse section of the midpiece, there are a few mitochondria encircling the axonemal complex. The axonemal complex has the typical 9+2 structure. There is said to be no annulus between the midpiece and principal piece but this requires confirmation; in micrographs a typical annulus is not distinguishable, though presence as a rudiment cannot entirely be rejected. Within the midpiece the doublets have each an outer dense fiber. Principal piece. The principal piece is demarcated by the presence of an amorphous fibrous sheath surrounding the axoneme. The A subtubule of each doublet has dense contents as in the endpiece.

Avian Spermatozoa: Structure and Phylogeny

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Endpiece. The endpiece is demarcated by absence of the fibrous sheath. Terminally, the typical axonemal sequence gradually becomes disorderly and unsystematic and the A subtubules have lost their dense contents. The B subtubules become open crescents in transverse section. There is no evidence for the tip granule seen in Gallus sperm.

8.8.1.6

Numida meleagris

The spermatozoon of the Guineafowl (Numida meleagris, Numididae), has been described ultrastructurally (Thurston et al. 1982; Hess et al. 1986; Thurston and Hess 1987; Aire and Soley 2003). General morphology. The general structure of the Numida spermatozoon (Fig. 8.17A, B) is similar to that described for turkey and rooster spermatozoa. The apex of the spermatozoon consists of a conical acrosome ca 1.8 mm long and 0.47 mm wide at the point of articulation with the nucleus (Fig. 8.18A, B). The acrosome is a homogenous, membrane-bound, “cap-like” structure superimposed on a lanceolate spine (here termed the perforatorium), and

Fig. 8.17 Numida meleagris, Guineafowl. A. Scanning electron micrograph of the entire spermatozoon. B. Transmission electron micrograph of a longitudinal section through the posterior nucleus to anterior tail. Approximately six mitochondria span the length of the midpiece. A, acrosome; AX, axoneme; N, nucleus; M, midpiece; T, flagellum. Thurston, R. J., Hess, R., Hughes, B. L. and Froman, D. P. 1982. Poultry Science 61: 1738-1743.

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Reproductive Biology and Phylogeny of Birds

Fig. 8.18 Numida meleagris, Guineafowl. A. SEM of sperm apex. ¥ 22000. B. TEM of a longitudinal section (LS) of the acrosome and anterior nucleus. ¥ 42000. C. SEM midpiece; arrow marks articulation between nucleus and midpiece. ¥ 19000. Fig. 8.18 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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distally it extends over the nuclear chromatin (Fig. 8.18B). The nucleus and acrosomal cap are concentrically contiguous where they overlap (Fig. 8.18G) (Thurston et al. 1982). The total length of the spermatozoon is 75-80 µm, compared with 90 µm or more for the rooster (Thurston and Hess 1987). Perforatorium. The perforatorium (Fig. 8.18B) is similar to, but longer than, that of rooster sperm but it was not discerned whether the perforatorium was hollow as in turkey spermatozoa. It is 1.9 µm long, versus 1.0 µm for turkey and rooster. In most spermatozoa, the perforatorium is inserted into a deep anterior nuclear concavity or fossa (here considered a reduced endonuclear canal) and projects angularly, terminating near the end of the cap (Fig. 8.18B). Granular material separates the perforatorium from acrosomal material and nuclear chromatin (Fig. 8.18F, H) (Thurston et al. 1982; Thurston and Hess 1987). Nucleus. Distal to the acrosome is the nucleus, approximately 12.8 mm long and 0.49 mm wide (Figs. 8.17A, 8.18A). The plasmalemma and nuclear membranes are not distinguishable as they are intermeshed to form a wavy network (Fig. 8.18E, I). However, a double nuclear membrane is discernible in the subacrosomal space and where the perforatorium inserts into nuclear fossa (Fig. 8.8B, G). At the distal end of the nucleus there is an implantation fossa (Fig. 8.18E). Midpiece. The midpiece, approximately 3.9 mm long, and 0.59 mm wide, consists of a distal centriole and anterior axonemal complex circumferentially encased by helically arranged mitochondria and the plasmalemma (Figs. 8.17B, 8.18D). The 25-30 mitochondria project from distal extensions of the nucleus caudally to the annulus (Fig. 8.18D, E), which marks the terminus of the midpiece. They are said to be arranged in a helix. This is consistent with the arrangement of mitochondria in rooster and turkey sperm (Thurston et al. 1982; Thurston and Hess 1987). Outlines of mitochondria are visible on the surface of the midpiece as a result of possibly artefactual depressions (Fig. 8.18C). In most species, Fig. 8.18 Contd. ...

D. TEM distal midpiece. E. TEM of LS of distal midpiece and anterior tail; arrow indicates concave implantation fossa. ¥ 34700. F. Transverse section (TS) of acrosome tip ¥ 42000. G. TS region where distal extension of acrosome vesicle overlaps the nucleus ¥ 33600. H. TS. Where perforatorium projects into the nucleus. ¥ 42000. I. TS of the nucleus. J. TS distal centriole. ¥ 42000. K. TS of midpiece immediately distal to point of origin of the central microtubules of the axoneme; arrow indicates dense fiber. ¥ 42000. L. TS flagella at their proximal (P) and distal (D) ends. ¥ 42000. Abbreviations: A, acrosome; An, annulus; Ar, arms extending from the dense material of the centriole and terminating in juxtaposition to the nuclear membrane; B, double bridge across central microtubules; C, acrosome vesicle; CM, central singlet microtubules of axoneme; DC, distal centriole; M, midpiece and mitochondria; N, nucleus; S, perforatorium; Sp, spaces between mitochondria. Partly relabeled after Thurston, R. J., Hess, R., Hughes, B. L. and Froman, D. P. 1982. Poultry Science 61: 1738-1743, Figs. 1-16.

"" Reproductive Biology and Phylogeny of Birds including roosters and turkeys, the cristae of germinal cell mitochondria are arranged parallel to the outer limiting mitochondrial membrane, but in Guineafowl sperm, they are both parallel and obliquely aligned (Fig. 8.18D, E, K). A fine, granular amorphous material occupies the cisternae created by the cristae, often making it difficult to resolve the inner membranes (Fig. 8.18K). Usually six mitochondria are visible in longitudinal section (Fig. 8.17B) and four in cross section (Fig. 8.18K), totaling 24 for the spermatozoon. Their three-dimensional structure is probably that of a polygonal plate as for rooster spermatozoa (Bakst and Howarth 1975), but dimensions measured from cross and longitudinal sections are approximately 0.6 mm ¥ 0.4 mm ¥ 0.16 mm. The distal termination of the midpiece is marked by an annulus which appeared dense and triangular in longitudinal sections (Fig. 8.18D). Guineafowl sperm differ from those of turkey and rooster in having mitochondrial cristae that are often oblique and the inner matrix is more dense. Centriole. Although rooster and turkey spermatozoa contain a proximal centriole orientated transversely to the distal centriole, (Thurston et al. 1982) states that in the Guineafowl spermatozoon, the midpiece contains only a distal centriole (Fig. 8.18E). However, it was later considered that in-line orientation of a proximal centriole with the distal centriole could not be discounted (Thurston and Hess 1987) and presence of both centrioles in line has been confirmed by Aire and Soley (2003). The non-striated connecting piece of the Guineafowl spermatozoon consisted of projections originating from the wall of the distal centriole. To accommodate this arrangement, the implantation fossa is deeper and more curved, the caudal end of the nucleus forming a semicircular concavity (Fig. 8.18E). Dense material similar to the nonstriated connecting piece of rooster and turkey sperm projects laterally as “arms” from the wall of the distal centriole toward the nuclear membrane (Fig. 8.18J). Although not clearly discernible, there appears to be one “arm’’ per group of three centriolar microtubules. When the region of the implantation fossa is sectioned transversely, the nine groups of three microtubules of the distal centriole, arranged in a “pinwheel” fashion, are observed embedded in dense material contiguous with the “arms” of the nonstriated connecting piece (Fig. 8.18J). For rooster and turkey spermatozoa, where the centrioles are mutually perpendicular, such a section would longitudinally bisect the proximal centriole and connecting piece. The distance from the apical end of the centriole to the commencement of the central singlets is 0.65 mm (Thurston and Hess 1987). Flagellum. The flagellum averages 59 mm (Thurston et al. 1982) but reaches 65 mm (Thurston and Hess 1987) in length and 0.52 mm wide (at the junction with the midpiece), and its ultrastructure is similar to that described for rooster and turkey spermatozoa. As in those species, dense outer fibers are absent from the axoneme. The central region of the A doublet microtubule retains dense material throughout the flagellum similar to that of the centriolar wall (Fig. 8.18L). A wide amorphous fibrous sheath (Fig. 8.18L) surrounds the axoneme

Avian Spermatozoa: Structure and Phylogeny

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in the principal piece (Thurston and Hess 1987). An endpiece is illustrated (Fig. 8.18L) demarcated by absence of the fibrous sheath. The central singlet microtubules originated from the basal granule (distal centriole) approximately 1.0 mm from the implantation fossa (Fig. 8.18E). Thus, the distal centriole in guinea spermatozoa is stated by Thurston et al. (1982) to be shorter than of rooster or turkey sperm. From the illustration this appears to be the case but it must be remembered that in ratites the central singlets deeply penetrate the distal centriole. At the terminus of the endpiece the doublets “probably become singlets”. Remarks. The spermatozoon of Guineafowl is closely similar to, but shorter than, that of the rooster and Turkey. The distal centriole, or at least the length from the apical end of the centriole to the commencement of the central singlets, is shorter, at 0.65 mm, compared with 2.2 mm and 0.9 mm for Turkey and rooster respectively (Thurston and Hess 1987). As the terminal region of the endpiece is not described it is not known whether the central “tip granule” seen in Gallus and Coturnix is present.

8.8.2 Order Anseriformes Ultrastructural accounts of anseriform sperm are those for Anas platyrhynchos, Mallard (Humphreys 1972; Maretta 1975a, b) and Branta sandvicensis, Hawaiian goose (Humphreys 1972). Light microscope accounts exist for the spermatozoa of Tadorna tadorna, Common Shelduck (Ballowitz 1888), Anas platyrhynchos, Mallard, Aythya fuligula, Tufted duck (Retzius 1909) (Fig. 8.10), and Somateria mollissima, Eider (McFarlane 1963) (Fig. 8.27C). General morphology. Like the Galliformes, Trogoniformes and Charadriiformes, inter alia, the anseriform sperm, as seen by light microscopy, is not helical and the acrosome and midpiece are shorter than the nucleus. As in galliforms, the acrosome is conical and the midpiece only a small fraction of the length of the elongate nucleus (Ballowitz 1888; Retzius 1909; McFarlane 1963).

8.8.2.1

Anas platyrhynchos

The following ultrastructural account is taken chiefly from Maretta (1975a, b) but employs current terminology. The drake spermatozoon (Fig. 8.19A-M) has the form of a cylinder, apically terminated by the pointed acrosome. The length and diameter [of the head] range within 10-11 mm and 0.6-0.7 mm, respectively. Acrosome. The acrosome vesicle (Fig. 8.19A-D) consists of homogeneous material of moderate density surrounded by a single membrane. It is inverted V-shaped and reaches 2.2 mm in length. The thickness of its wall is 0.1-0.2 mm, being thinnest at the site where the nucleus is attached. The perforatorium (termed the acrosome spine), composed of electron-dense material, is needleshaped, reaching an average length of 2.6 mm and a diameter of 0.2 mm. Of this length, 1 mm lies in an invagination (anterior nuclear fossa or endonuclear

"$ Reproductive Biology and Phylogeny of Birds

Fig. 8.19 Anas platyrhynchos, Mallard. A. Longitudinal section (LS) of an acrosome. The acrosome vesicle (AC) covers the perforatorium (AS). Along the inner part of the perforatorium vacuoles are seen. The space between the perforatorium and the vesicle is filled with granular material (GM). B-E. Transverse sections (TS) of an acrosome at different levels. The vacuoles (V) inside the perforatorium are clearly visible. F. LS of the caudal region of the head. The chromatin is coarsely granular with a small vacuole present in the middle. Cytoplasmic membrane closely applied Fig. 8.19 Contd. ...

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canal in current terminology) of the proximal portion of the nucleus. The remaining, more anterior portion of the rod projects into the subacrosomal space and is covered by the acrosome vesicle. Longitudinally running vacuoles are present within the rod along its entire length, their number varying from two to five and sometimes more, and varying in size (Fig. 8.19BD). The vacuoles communicate with the space between the acrosome vesicle and the rod. They are empty or filled with a moderately dense substance. The acrosome vesicle and the perforatorium are separated by a 10-40 nm wide space, widest around the processes of the nucleus (nuclear rostrum). The space is filled with a granular substance of low electron-density. Anteriorly the perforatorium touches the posterior membrane of the acrosome vesicle whereas basally it may or may not reach the base of the short endonuclear canal. Within the endonuclear canal the perforatorium is closely applied to the inner nuclear membrane lining the canal (Fig. 8.19A). Nucleus. The chromatin of the nucleus is of uniform density and composed of large granules of about 60 nm which are closely attached to each other. The nucleus is surrounded by a double membrane which in mature spermatozoa is so closely applied to the chromatin that it can hardly be differentiated. Sometimes the nuclear membrane becomes obvious only in the anterior region of the head at the site where the acrosome vesicle is attached, at the endonuclear canal, and at the base of the nucleus where it articulates with the tail (Maretta 1975a). Posteriorly there is a broad, shallow basal nuclear fossa (implantation fossa) (Fig. 8.19F). Neck, midpiece and centrioles. The tail of the drake spermatozoon is composed of four parts: a neck, a midpiece, a principal piece and an endpiece. The neck is the place of articulation between the head and the tail. In contrast to the distal centriole, the proximal centriole (Fig. 8.19F) maintains its original size. At one end of its lumen an electron-dense material can be seen. Its wall consists of nine triplet tubules and forms a right angle to the distal Fig. 8.19 Contd. ...

to the nuclear surface (N). G. TS sperm head. At bottom is a TS of the basal fossa with remnants of the proximal centriole. H. LS of the neck and part of the mitochondrial sheath. Mitochondria (M) are of irregular subspheric shape. I. TS of the caudal part of the head and anterior region of the midpiece. The proximal centriole is surrounded by electron-dense material. J. TS neck in the region of the proximal centriole (PC). K. LS midpiece. Central singlets (CF) arise from the caudal end of the distal centriole (DC). The cristae are arranged parallel to the flattened mitochondrial wall (M). The midpiece terminates at an annulus. L. TS midpiece through proximal part of distal centriole. M. TS midpiece at different levels. Outer dense fibers are present (F). A. ¥ 47700. B. ¥ 26000. C. and D. ¥ 27300. E. ¥ 33500. F. ¥ 27000. G. and H. ¥ 27000. I. ¥ 36500. J. ¥ 37700. K. ¥ 28200. L. ¥ 26100. M. ¥ 22800. A-G. Relabeled after Maretta, M. 1975. Acta Veterinaria Academiae Scientiarum Hungaricae 25(1): 47-52, Figs. 1-7. H-M. Relabeled after Maretta, M. 1975. Acta Veterinaria Academiae Scientiarum Hungaricae 25(1): 5360, Figs. 1-6.

"& Reproductive Biology and Phylogeny of Birds centriole. In cross sections it appears as a thick-walled ring in which some tubules of the triplets can be partly identified (Fig. 8.19H, I, J). The midpiece (Fig. 8.19H-M) is defined by the length of the mitochondrial sheath measuring in the drake sperm 3-4 mm, and having a diameter of 0.60.7 mm. The main portion of the midpiece consists of’ the centrally placed distal centriole and part of the axial filament complex. In mature spermatozoa the distal centriole reaches a length of up to 2 mm and a diameter of 0.2 mm. It appears to be 3-4 times longer than the proximal centriole, the latter being shown in Fig. 8.19F, J, K). The lumen of the proximal portion of the centriole is usually filled with electron-dense material (Fig. 8.19L). The proximal centriole is orientated with its long axis transverse relative to the longitudinal axis of the sperm (Fig. 8.19F, H, K). The distal centriole is connected with the proximal centriole by dense material, which also fills the space under the projecting parts of the proximal centriole. Within the caudal end of the centriole a small amount of electron-lucent material is placed where the central fibrils arise. Annulus. The annulus is a ring-shaped formation (Fig. 8.20B) located at the junction of the mitochondrial sheath of the midpiece and the amorphous sheath of the principle piece (Figs. 8.19K, 8.20A). In cross section it is almost triangular with a wall length of 100-150 nm. It is composed of homogeneous material suggesting sometimes a fibrous structure. The cytoplasmic membrane covering the entire length of the tail is closely applied to the underlying structures. It is fixed only to the annulus. In the posterior portion of the principal piece the membrane is separated from the amorphous sheath by a narrow lighter space. Axoneme. The axoneme has the usual 9+2 pattern. The denser subtubule of the each doublet (subtubule A) is smaller and lies nearer to the axis of the tail than does the larger, less dense subtubule B. Subtubule A bears two dynein arms. Outer dense fibers are present (Fig. 8.19M), they reach a diameter of about 40-50 nm and are closely attached to the doublets. They are visible only in the posterior region of the midpiece reaching as far as the beginning of the principal piece. Further externally, mitochondria are attached to the distal centriole and part of the axoneme. In the anterior part of the midpiece mitochondria closely approach the head and are caudally defined by the annulus. They are of irregular spherical shape (Fig. 8.19H) and their total number ranges between 24-30. They are flattened at the sides and about 0.2 mm thick. Mitochondrial cristae run parallel to the flattened mitochondrial wall (Fig. 8.19 J, K, L). Principal piece. The principal piece of the tail (Fig. 8.20A, C, D) is defined by the length of the amorphous sheath. Cranially, it starts at the annulus and passes caudally into the endpiece. Remnants of the outermost dense fibers are visible only in its anterior region. Amorphous sheath. The amorphous sheath consists of moderately electrondense material and its wall has a diameter of 0.1 mm in its proximal portion. It is separated from the axoneme by a 20 nm space. It is composed of an inner

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Fig. 8.20 Anas platyrhynchos, Mallard. A. Longitudinal section of principal piece in the anterior and posterior region. The midpiece and principal piece are separated by an annulus (An). An amorphous sheath (AS) surrounds the axoneme (AFC). B. Transverse section (TS) of the tail at the annulus (An). Outer dense fibers (F) are present. C. TS of the tail in the proximal region of the principal piece. The amorphous sheath (AS) is composed of an outer less dense and an inner dense layer. D. TS posterior region of principal piece. The amorphous sheath (AS) here consists only of the inner dense layer. E. TS endpiece. F. TS termination of endpiece, showing disruption of 9+2 pattern. A. ¥ 3200. B-D. ¥ 39700. E. ¥ 38000, F. ¥ 37900. After Maretta, M. 1975. Acta Veterinaria Academiae Scientiarum Hungaricae 25(1): 53-60, Figs. 7-12.

" Reproductive Biology and Phylogeny of Birds more dense layer and an outer less dense layer. Candally, the sheath gradually becomes narrower; the outer layer disappears first, then, gradually, the inner layer. The site where the amorphous sheath disappears is the junction with the endpiece. Endpiece. The endpiece (8.20E) is composed of the axial filament complex (axoneme). Near the tip of the endpiece the arms of the doublets disappear and subtubule A loses it dense contents. The doublets are reduced to single tubules and gradually decrease in number (Maretta 1975b) (Fig. 8.20F).

8.8.2.2 Branta sandvicensis The sperm of Branta sandvicensis, the Hawaiian goose, compared with those of the Mallard by Humphreys (1972) had an identical appearance which conformed closely to that of Gallus sperm. Each spermatozoon was about 100 mm long, with a nuclear diameter of about 0.5 mm. Humphreys (1972) gave a graphical comparison of a mallard and canary sperm.

8.8.2.3 Conclusion for Galloanserae The spermatozoa of the Anseriformes are closely similar to those of the Galliformes but a notable distinction appears to be that in anseriform sperm, as exemplified by Anas branchyrhynchos, the perforatorium extends almost to the tip of the spermatozoon, as the acrosome vesicle is apically very narrow, whereas in galliforms a large amount of material is present in the acrosome vesicle anterior to the tip of the perforatorium, correlated with a much shorter subacrosomal space. Demonstration that this difference is constant would require examination of a larger sample of species. The galloanseran spermatozoon resembles that of crocodile and paleognaths in the conical acrosome vesicle, much shorter than the nucleus; perforatorium penetrating the nucleus in an endonuclear canal; mitochondria in tiers surrounding the distal centriole; and presence of an annulus and of a fibrous sheath. It differs in the short stout form of the perforatorium; short endonuclear canal; and the amorphous, not ribbed, fibrous sheath. Anseriforms possess what appears to be the most primitive avian spermatozoon above the paleognaths.

8.9

METAVES

As noted by Harshman (Chapter 1 of this volume), Fain and Houde (2004) divided Neoaves into two basal clades: Metaves, consisting of Caprimulgiformes, Apodiformes, Podicipedidae, Phaethontidae, Phoenicopteridae, Opisthocomidae, Mesitornithidae, Rhynochetidae, Eurypygidae, Pteroclidae, and Columbidae; and Coronaves, consisting of the remaining Neoaves. This subdivision of the Neoaves requires confirmation from other analyses as it was based solely on an analysis of intron 7 of the b-fibrinogen gene. It cannot be said to be supported by spermatozoal ultrastructure (see section 8.11).

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In the Metaves, only the Apodiformes, Caprimulgiformes and Columbiformes have been examined for spermatozoal ultrastructure.

8.9.1

Apodiformes

Spermiogenesis in Apus (=Cypselus) apus, the Common swift was briefly treated by Baccetti et al. (1980), who reported it as a passerine, in a valuable work on the vertebrate perforatorium and by Tripepi et al. (1984) in an abstract. Jamieson and Tripepi (unpublished; see also 2005) made a more detailed investigation of the late spermatid. The only other ultrastructural account for the Apodiformes is a brief account of microtubules in the spermatid of Apus melba, the Alpine swift (Tripepi et al. 1991).

8.9.1.1

Apus (=Cypselus) apus

Acrosome. The acrosome vesicle (Fig. 8.22A, E-H) forms a slender, smooth, pointed cone approximately 3 µm in length and 0.75 mm at its greatest, basal width. Its base is rounded and closely fits a depression of the anterior end of the nucleus. A perforatorium is absent. From a light microscope drawing by Retzius (1911) (Fig. 8.21) the acrosome: nucleus ratio is very approximately 0.03. Nucleus. The nucleus is an elongate cylinder (Fig. 8.22A, B, E, I, K, P) tapering only slightly towards its tip. Its full length has not been determined but it exceeds 8 µm, with a basal width of 0.6 µm. In young spermatids the chromatin is finely granular and the nuclear membrane is surrounded by microtubules, in single layer, which lie under the plasma membrane (Fig. 8.22J). In the more mature nucleus (Fig. 8.22K) the chromatin forms dark clumps interspersed sporadically throughout its length with pale areas, some of which impinge on its surface, and few microtubules remain beneath its investing membrane (Fig. 8.22B). In the mature nucleus the chromatin is electron dense and almost homogenous and microtubules are absent (Fig. 8.22I). The nuclear surface is almost smooth. The anterior nuclear fossa is matched by a concave posterior fossa, the implantation fossa (Fig. 8.22B, P). Midpiece and centrioles. The elongate cylindrical midpiece in which the mitochondria are located is wider, at 1.1 µm, than the nucleus. Its length is approximately 3.5 µm (Fig. 8.22M). Its central axis is occupied by the proximal centriole, which lies partly within the implantation fossa, and by the distal centriole. The proximal centriole is short, with its longitudinal axis perpendicular to the sperm axis. It shows the usual nine triplets of microtubules (Fig. 8.22B, N-P) but its central space is occupied by a structure which is annular in transverse section (Fig. 8.22B, N-P). The distal centriole, perpendicular to the proximal centriole and in the long axis of the cell, also shows a triplet configuration (Figs 8.22L, Q). It extends for the whole length of the midpiece. Its axis is empty except for intrusion of the central singlets of the axoneme a very short distance into its base (Figs 8.22C, D, M).

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Reproductive Biology and Phylogeny of Birds

Fig. 8.21 Drawings of some non-passerine sperm by light microscopy. Philomachus pugnax, Ruff; Psittacus sp.; Strix aluco, Tawny owl; Dendrocopos (=Picoides) major, Great spotted woodpecker; Columba livia, Domestic pigeon. Fig. 8.21 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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The mitochondria form a circle around the distal centriole, numbering five or six in a transverse section of the cell (Fig. 8.22L, Q). There are six or seven in longitudinal sequence (Fig. 8.22D, M) but some of these may be partly conjoined (Fig. 8.22M). Posteriorly the midpiece narrows slightly but is not demarcated by a recognizable annulus (Fig. 8.22 D, M). Each mitochondrion has several cristae which appear transverse in cross- and oblique in longitudinal section of the midpiece. They are initially subspherical (Fig. 8.22D) but become more elongate nearer maturity (Fig. 8.22B, C, D, M). Axoneme. Immediately behind the midpiece, the axoneme commences as indicated by the presence of central singlets. A moderately electron-dense mass at the anterior end of these protrudes a little into the midpiece (Fig. 8.22C, D, M). An amorphous sheath (Fig. 8.22C, D, M) surrounds the axoneme behind the midpiece and the long ensheathed region constitutes the principal piece. Dense fibers have not been observed. A presumed endpiece, with axoneme lacking the amorphous sheath is surrounded by a transient cytoplasmic canal and sheath during development (Fig. 8.22R). Remarks. The long distal centriole is a remarkably plesiomorphic feature of the swift sperm, being seen only in paleognaths, as it is somewhat shortened even in the Galloanserae. In the Palaeognathae, struthioniforms differ, however, in penetration of the distal centriole by the two central axonemal singlets (Baccetti et al. 1991; Phillips and Asa 1989; Soley 1993, 1999), though these reach only about halfway into the centriole in Crested tinamou (see Fig. 1 of Asa and Phillips 1987). On the other hand, loss of the perforatorium in Apus is a notable apomorphic departure from paleognaths and Galloanserae. The phylogenetic implications of the long centriole and other features of the sperm are further discussed in section 8.11. In Apus apus, microtubules in the spermatid are restricted to a transient layer encircling the cell, though longitudinal microtubules are also present in the Sertoli cell which invests the spermatid, as also seen in A. melba (Tripepi et al. 1991). This condition contrasts with passerines in which, in the spermatid, an ‘helical membrane’ consists of multiple microtubules forming a thick strand helically coiled around at least the flagellum (e.g. Asa and Phillips 1987; Jamieson 2006).

8.9.1.2

Apus melba

Tripepi et al. (1991) comment on, and illustrate, the microtubular arrangement in the spermatid of the Alpine swift (Apus melba). Many microtubules, arranged parallel to the longitudinal axis of the elongating spermatid appear in the cytoplasm of the Sertoli cell and surround the head of the spermatid. Fig. 8.21 Contd. ...

After Retzius, G. 1909. Biologische Untersuchungen, Neue Folge 14(10): 89-122 Taf XXX, Fig. 14, XXXI, 13, 23, 27. Apus (=Cypselus) apus, Common swift. After Retzius, G. 1911. Biologische Untersuchungen, Neue Folge 16: 89-92 Taf XXVII, Fig. 24.

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Fig. 8.22 Apus apus, Common swift. Transmission electron micrographs of spermatids. A. Oblique longitudinal section (LS) of acrosome on tip of nucleus. B. LS late spermatid showing elongate nucleus, with scattered uncondensed areas, and anterior portion of midpiece with enclosed proximal and distal centrioles. C. Same cell, showing posterior end of midpiece and anterior portion of principal piece. D. LS of the midpiece of a younger spermatid in which mitochondria are subspherical. Arrow indicates absence of an annulus. E. LS of two acrosomes. F-H. Progressively posterior transverse sections (TS) of acrosome vesicle, H, at the level of the anterior nuclear fossa. I. TS of an advanced nucleus with strongly condensed chromatin and lacking peripheral microtubules. J. TS nucleus of younger spermatid with uncondensed granular chromatin and peripheral single layer of microtubules. K. TS nucleus at intermediate stage still showing pale uncondensed areas. L. TS immature midpiece, showing six mitochondria surrounding the distal centriole. M. Longitudinal section (LS) of an advanced spermatid through the entire length of the midpiece and through the Fig. 8.22 Contd. ...

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There is therefore a simultaneous occurrence of the longitudinal manchette in the spermatid and the microtubules of the Sertoli cell, as in Crested tinamou.

8.9.2

Order Caprimulgiformes

The Caprimulgiformes were subsumed in the Strigiformes by Sibley and Ahlquist (1990), as endorsed by ITIS (http://www.itis.usda.gov/). However, as relationship of the aegolothelids (owlet-nightjars) is closer to apodiforms than to strigeids, inclusion of caprimulgiforms in the Strigiformes is contraindicated. The order Caprimulgiformes is therefore recognized here while recognizing that paraphyly or even polyphyly of the order (see Chapter 1) may preclude association of caprimulgids with the aegoleothelids. Strigidae are now placed in the Coronaves (see Harshman, Chapter 1) and this is supported by sperm ultrastructure (Section 8.10.2) We owe to Ballowitz (1888) a drawing of a spermatozoon of Caprimulgus europaeus, the European Nightjar. The spermatid of C. europaeus has been the subject of a paper on microtubules by Tripepi et al. (1991) and has been described in some detail by Tripepi, Jamieson and Brunelli (unpublished) from whose account the following description is largely drawn.

8.9.2.1

Caprimulgus europaeus

The structure of the spermatozoon of Caprimulgus europaeus has been deduced from the description by TEM (Fig. 8.23) of the late spermatid (Tripepi unpublished) considered in conjunction with the light microscope description of the mature spermatozoon by Ballowitz (1888). Acrosome. The acrosome consists of the acrosome vesicle and associated material of a presumed perforatorial nature. Condensation of dense material at the nuclear surface at the base of the subacrosomal space results in compaction to form an electron dense, stout, spine-like structure, the putative perforatorium. This penetrates a short distance into the tip of the nucleus within an endonuclear canal and protrudes into the subacrosomal space (Fig. 8.23A). In transverse section, the acrosome vesicle forms a thick ring, with moderately electron-dense contents, around the tip of the nucleus with its central endonuclear canal and perforatorium (Fig. 8.23B). The maturing acrosome vesicle (Fig. 8.23A) assumes a conical form; the perforatorium becomes less electron-dense than formerly and the portion within the endonuclear canal is considerably longer than the ill-defined portion in the now conical subacrosomal space. Fig. 8.22 Contd. ...

principal piece. N,O. TS proximal centriole and adjacent midpiece, N being detail of O, showing the central structure of unknown significance. P. TS proximal centriole and adjacent nucleus and midpiece. Q. Transverse sections (TS) through two midpieces. R. TS endpiece still surrounded by a transient cytoplasmic canal and cytoplasmic sheath. as = amorphous sheath; av, acrosome vesicle; ax, axoneme; dc, distal centriole; m, mitochondrion; mt, microtubules; n, nucleus; pc, proximal centriole. From Jamieson and Tripepi, unpublished, see also Acta Zoologica (Stockholm) 86: 239-244, Figs. 1 and 2.

"$ Reproductive Biology and Phylogeny of Birds

Fig. 8.23 Caprimulgus europaeus. A. LS more mature acrosome, showing perforatorium lying in the endonuclear canal and extending somewhat amorphously into the subacrosomal space. The acrosome vesicle is assuming a conical form. B. Transverse section of an advanced acrosome showing the base of the acrosome Fig. 8.23 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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By light microscopy, the mature spermatozoon of Caprimulgus europaeus appears in a drawing (Fig. 85 in Ballowitz 1888) to have a conical acrosome tapering evenly from the much longer but fairly stout, curved nucleus. However, this species is not specifically mentioned in this regard in the description of the two types of sperm head recognized for birds by Ballowitz, though in the legend for Fig. 86 the head is described as “an der Spitze deutlich das kleine blasse Spitzen-stueck zeigend”. This pale point is consistent with the conical form of the maturing acrosome observed here. It is in marked contrast to the button-like acrosome of the only described member of the Strigiformes, the Tawny owl (Retzius 1909), the order in which caprimulgids were subsumed by Sibley and Ahlquist (1990). Nucleus. In the elongating spermatid the chromatin comes to form coarse, electron-dense granules (Fig. 8.23A, C). At maturity, as seen in the advanced spermatid, the nucleus is cylindrical with strongly condensed, homogeneous, electron-dense chromatin (Fig. 8.23D). Nuclear elongation is accompanied by growth of a longitudinal manchette of microtubules in the surrounding cytoplasm. No circular manchette is present. Midpiece. The Caprimulgus midpiece, about half the length of the nucleus, has some 10 tiers of mitochondria; the free axoneme is about the same length as the head plus midpiece (see Ballowitz 1888). In the advanced spermatid, mitochondria group around the proximal axoneme. A maximum number of six mitochondria has been observed, in transverse section, at one level (Fig. 8.23C, top left). Centrioles. Each spermatid has a proximal and distal centriole, mutually perpendicular, and the distal centriole is continuous with the flagellum. The centrioles have the normal configuration of nine triplets of microtubules. The lumen of the distal centriole is not, initially, penetrated by the central singlets of the axoneme but in the young elongating spermatid the distal centriole is penetrated by these singlets. Axoneme. The axoneme forms an extension of the distal centriole. In transverse section, the axoneme has the typical ‘9+2’ arrangement of Fig. 8.23 Contd. ...

vesicle enclosing the tip of the nucleus with its endonuclear canal and perforatorium. C. Transverse sections (TS) of midpieces and nucleus of advanced elongating spermatids. Four to six mitochondria surround the proximal axoneme as the midpiece. Dense fibers associated with the axonemal doublets are small in these sections. The chromatin of the nuclei has become clumped as condensation proceeds. Longitudinal but no circularly running cytoplasmic microtubules are present. D. LS nucleus fully condensed but still accompanied by the manchette. E. TS elongating spermatids. Left through the transient cytoplasmic canal and contained axoneme. Center right, TS through a developing midpiece which has assembled only four mitochondria at this level. F. TS axoneme with weakly developed amorphous sheath. av, acrosome vesicle; ax, axoneme; cytoplasmic canal; ec, endonuclear canal; m, mitochondrion; mp, midpiece; mt, microtubules of manchette; n, nucleus; p, putative perforatorium. Tripepi, Jamieson and Brunelli, unpublished.

"& Reproductive Biology and Phylogeny of Birds microtubules: Within the midpiece, a dense fiber is attached to the outer aspect of each of the nine doublets (Fig. 8.23C). The free axoneme is surrounded by a weakly developed amorphous sheath, defining the principal piece (Fig. 8.23F). Phylogenetic considerations. The structure of the spermatozoon of Caprimulgus europaeus can be deduced from the description by TEM of the late spermatid given above when considered in conjunction with the light microscope description of the mature spermatozoon by Ballowitz (1888). It is typical of other non-passerines in many respects. Features shared with paleognaths (ratites and tinamous) and the Galloanserae (e.g. rooster and duck) are the conical acrosome, shorter than the nucleus; presence of a perforatorium and endonuclear canal; presence of a proximal as well as distal centriole; the elongate midpiece with mitochondria grouped around a central axis (here maximally six mitochondria in approximately 10 tiers); and presence of a fibrous or amorphous sheath around the axoneme. Most of these features characterize non-passerines in general. A major (apomorphic) difference from paleognaths and galloanserans is the short distal centriole, the midpiece being penetrated by the axoneme and not by the centriole. In lacking an appreciable annulus, the sperm of Caprimulgus, like those of Psittaciformes (Jamieson 1999; Jamieson et al. 1995), Gruiformes (Grus vipio, Phillips et al. 1987), Apodiformes (Jamieson and Tripepi 2006), and passerines (e.g. Asa and Phillips 1987; Jamieson 1999), differ from those of paleognaths (e.g., Baccetti et al. 1991), galloanserans, and charadriiforms as represented by Jacana (Saita et al. 1983). An annulus is basal to paleognaths and these nonpasserines. Absence of the annulus is therefore an apomorphic feature of caprimulgid sperm. In Caprimulgus europaeus, the circular manchette has been lost and only a longitudinal manchette is present in the developing spermatid (Tripepi et al. 1991). A similar arrangement is seen in Jacana jacana and (Jamieson and Tripepi 2005) in the apodiform Apus apus. In contrast, passerines differ from non-passerines in possessing, in the spermatid, an ‘helical membrane’, consisting of multiple transverse and longitudinal microtubules forming a thick strand helically coiled around at least the flagellum (e.g. Asa and Phillips 1987; this study). Tripepi et al. (1991) consider the arrangement of microtubules in C. europaeus to be the second above a “reptilian” level. For further phylogenetic considerations see section 8.11.

8.9.3

Order Columbiformes

Columbiforms for which sperm ultrastructure has been described are: ‘Pigeon’ (presumably Columba livia) (Fawcett et al. 1971), restricted to a consideration of nuclear shaping in the spermatid; Columba livia, Domestic pigeon (Yasuzumi and Yamaguchi 1977), spermiogenesis, mainly concerning development of the acrosome and nuclear microtubules; Turtle dove (Streptopelia roseogrisea), chiefly spermiogenesis (Mattei et al. 1972); and brief descriptions of mature spermatozoa of Peaceful dove (Geopelia striata, Jamieson et al. 1995, 1999); and of Crested pigeon (Ocyphaps lophotes, Jamieson et al.

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1995, 1999). Sperm of G. striata and O. lophotes are illustrated (Figs. 8.24, 8.25) and described more fully below. Valuable accounts by light microscopy are those of Ballowitz (1888), Retzius (1911, 1912) and Vernon and Woolley (1999; also giving ultrastructure) for Columba livia. Five genera, with six species, were examined by McFarlane (1963); all except Blue Ground-Dove (Claravis pretiosa) were unnamed. These results are, however, summarized in his drawing showing a pointed acrosome much shorter than the very elongate cylindrical nucleus, and a long cylindrical midpiece, stated to extend far down the tail, in a nonhelical sperm (Fig. 8.27D). Acrosome. The acrosome vesicle forms an elongate blunt-tipped cone which is shorter, at ~1.0-1.2 mm (n=2), in Ocyphaps lophotes (Fig. 8.24A, B) than the ~2.6-2.8 mm (n=2) in Geopelia striata (Fig. 8.25A, B). The subacrosomal space forms a convex sided cone less than half the length of the vesicle. Transverse sections of the vesicle reveal a circular profile (Figs. 8.24E, F, 8.25G-J). A striking contrast with Galloanserae, psittaciforms and most other nonpasserines is the absence of a perforatorial rod. Instead of the subacrosomal space being occupied by a perforatorium, it is filled by a narrow anterior extension of the nucleus, the nuclear rostrum. However, the electron-dense chromatin does not fully occupy the investing conical nuclear membrane but leaves a varying length of the tip of the rostrum free of chromatin (O. lophotes, Fig. 8.24A, B; G. striata, Fig. 8.25A, B). In some cases (Fig. 8.25B) the pale tip of the rostrum has an appearance reminiscent of a perforatorium but this seems to be an artifact of eccentric sectioning of the rostrum as the electron dense rostrum extends far into the subacrosomal space in other sections (Fig. 8.25A). Nevertheless, it is possible that some of the finely granular material at the tip of and surrounding the rostrum represents a reduced perforatorium. In support of this it is noteworthy that some transverse sections of the rostrum reveal a central pale core (O. lophotes, Fig. 8.24E; G. striata, Fig. 8.25I) which cannot merely be dismissed as uncondensed chromatin as in some sections (Fig. 8.25J) it is in continuity with the subacrosomal matrix. This central lacuna in the rostrum, also seen in longitudinal section (Figs. 8.24, 8.12A), may, therefore, be a vestigial endonuclear canal. Nucleus. The nucleus forms a stout cylinder, circular in cross section (Figs. 8.24C, 8.25E) with shoulders, supporting the base of the acrosome vesicle, where it narrows to form the rostrum. The chromatin is electron-dense and, at maturity, almost homogeneous. The base of the nucleus widens slightly and is indented by a shallow but distinct basal fossa which forms an arc in longitudinal sections of the nucleus and houses the anterior part of the proximal centriole (O. lophotes, Fig. 8.24C, D; G. striata, Fig. 8.25K, L, M). The nuclei illustrated by light microscopy for Columba livia by Retzius (1909) (Fig. 8.21) and Claravis pretiosa by McFarlane (1963) are moderately elongate, though that for C. livia by Ballowitz (1888) appears shorter than that illustrated by Retzius.

"  Reproductive Biology and Phylogeny of Birds

Fig. 8.24 Ocyphaps lophotes, Crested pigeon. Transmission electron micrographs of spermatozoa. A and B. Longitudinal section (LS) acrosome and anterior portion of nucleus. C. LS base of nucleus, centrioles and anterior midpiece. D. LS almost mature, testicular showing same regions. E. Transverse section (TS) of base of acrosome vesicle surrounding the nuclear rostrum, the latter showing a central vacuole or canal. F. TS showing nuclear rostrum nearer its base. G. TS nucleus. H. TS distal centriole at the transition from 9 triplets to doublets, surrounded by mitochondria of the anterior midpiece. I. TS midpiece through 9+2 axoneme. J. LS midpiece, showing mitochondria aligned in single file and occasional dense bodies. K. TS axoneme posterior to midpiece. Original.

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Columbiforms appear plesiomorphic in retaining the nuclear rostrum, though it is possible that presence is a reversal (Jamieson 1999). Midpiece. In transverse section the midpiece has four to six mitochondria surrounding the distal centriole (Fig. 8.24H) and axoneme (Figs. 8.24I, 8.25C, N, O). In both species compact electron-dense bodies are interspersed between or lie medially to the mitochondria (Figs. 8.24J, 8.25M, N), an unusual condition for Aves but common in squamates. This co-occurrence, although appearing homoplastic, may well indicate persistence of a genetic basis laid down in early amniotes (Jamieson 1999). In longitudinal section (Figs. 8.24D, K, 8.25M) the mitochondria are seen to lie end to end in single file. Four or five parallel chains of mitochondria were described, and illustrated in transverse section, for Columba livia by Vernon and Woolley (1999). The length of the midpiece in Ocyphaps lophotes and Geopelia striata was not ascertained. Woolley (1995) estimated that in Columba livia “var. domestica”, the flagellar length is ca 150 µm, but that the midpiece occupies the proximal 114 mm (dimensions estimated from Ballowitz 1888, proportions confirmed from drawings by Retzius 1909) (see Fig. 8.21 of this chapter). He notes that this is the only record of a midpiece of an avian spermatozoon approaching the great length (161 mm) of that of Coturnix japonica, though here considered to require confirmation in the latter. Slightly different dimensions were given for the domestic pigeon by Vernon and Woolley (1999). The sperm head was about 16 µm long. The mean flagellar length with SD was 132.0 ± 11.1 µm; length midpiece 98.1 ± 11.2 µm (n = 13 sperm from 1 individual). Midpiece length and flagellar length were closely correlated: r = 0.89, P < 0.001. An annulus terminating the midpiece, basic to bird sperm, and seen in paleognaths, rooster and Guineafowl, has been reported for columbiforms (Asa and Phillips 1987) and specifically for Columba livia (Vernon and Woolley 1999). Centrioles. The proximal centriole is orientated with its long axis at right angles to the sperm axis (Figs. 8.24C, D, 8.25K, L, M), as stated and illustrated by Vernon and Woolley (1999) for Columba livia. Three projections (probably equivalent to striated columns) are seen to link it to the thickened basal membrane of the nucleus in Ocyphaps lophotes (Fig. 8.24C). The distal centriole forms the basal body orientated in the long axis of the axoneme with which it is continuous (Figs. 8.12C, D, 8.25C, D, L, M). Its posterior limit is difficult to determine but it is only a little longer than the proximal centriole, its length being estimated as 0.5 mm in Geopelia striata (Jamieson 1995). Both centrioles have the usual 9 triplets but where the distal centriole enters the midpiece the triplets progressively reduce to doublets (Figs. 8.24H, 8.25C). Each centriole is embedded in a ring of dense material. Axoneme. The axoneme shows the usual 9+2 pattern of microtubules. Within the proximal region of the midpiece of apparently mature sperm of Geopelia striata (Fig. 8.25N) and Ocyphaps lophotes (not illustrated) the axoneme has outer dense fibers (Jamieson 1995, 1999). These fibers are said by Mattei et al.

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Reproductive Biology and Phylogeny of Birds

Fig. 8.25 Geopelia striata, Peaceful dove. Transmission electron micrographs of spermatozoa. A and B. Longitudinal section (LS) of acrosome and anterior portion of nucleus. B cut off center through the nuclear rostrum. C. TS distal centriole at the Fig. 8.25 Contd. ...

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(1972) to be present in the spermatid but to be lost by maturation of the sperm in Streptopelia roseogrisea and they have not been observed in the sperm of O. lophotes. In the more distal region of the midpiece dense fibers are absent (Fig. 8.2O) and the axoneme posterior to the midpiece is simple (Figs. 8.24K, 8.25F). Vernon and Woolley (1999) state, for Columba livia, that accessory fibers occur proximally on the flagellum, in what were clearly mature sperm, but are extremely short. Fibrous sheath. A fibrous or amorphous sheath which surrounds the principal piece in lower non-passerines is absent from mature sperm of those columbiforms examined for this feature ultrastructurally (Streptopelia roseogrisea, Mattei et al. 1972; Geopelia striata and Ocyphaps lophotes, this chapter, and Columba livia, Vernon and Woolley 1999) as from psittaciforms. It was, however, demonstrated by Mattei et al. (1972) for the spermatid of S. roseogrisea. Remarks. The spermatozoa of columbiforms have many features of those of Galloanserae but differ notably in loss of the perforatorium and the fibrous sheath by maturity and the exceptionally great length of the midpiece, though the latter is possibly also greatly elongate in Coturnix japonica (section 8.8.1.2).

8.10

CORONAVES

The Coronaves includes the majority of taxa in the division of the Neoaves into Metaves and Coronaves by Fain and Houde (2004).

8.10.1

Order Piciformes

These constituted the Parvclass Picae in the system of Sibley and Ahlquist.

8.10.1.1

Melanerpes carolinus

The spermatozoa of the Piciformes are known only from the light microscope accounts of Ballowitz (1888) and Retzius (1909) for the Great spotted woodpecker (Dendrocopos (=Picus) major) (Fig. 8.21) and the brief ultrastructural account of Henley et al. (1978) for the Red-bellied woodpecker (Melanerpes carolinus) (Fig. 8.26A). Picoid sperm are markedly different from those of oscines and are considered by Henley et al. (1978) to be referable to Fig. 8.25 Contd. ...

transition from 9 triplets to doublets, surrounded by mitochondria of the anterior midpiece. D. Detail of same. E. TS nucleus. F. TS axoneme posterior to midpiece. G. TS anterior region of acrosome vesicle. H. TS basal region of acrosome through the nuclear rostrum. I and J. Same showing central vacuole in nuclear rostrum which in J communicates with the material in the subacrosomal space. K. LS showing base of nucleus, proximal centriole and anterior midpiece. L. Detail of proximal centriole. M. LS cut at right angles to K showing base of nucleus, proximal centriole and anterior midpiece. N. TS Midpiece showing a dense body between mitochondria and axoneme with nine uniform dense fibers. O. TS midpiece putatively further posterior, with the axoneme lacking dense fibers. Original.

" " Reproductive Biology and Phylogeny of Birds

Fig. 8.26 Melanerpes and Crotophaga A. M. carolinus, Red-bellied woodpecker. Transverse section through a group of maturing spermatozoa. The section passes Fig. 8.26 Contd. ...

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the type seen in ‘various fowls’ by Retzius (1909), McFarlane (1963), and in a duck by Henley et al. (1978). The acrosome in such sperm is said by Henley et al. (1978) to be “buttonlike”, rather than tapering and long as in oscines. However, the galliform and anseriform acrosome is here considered elongate conical, though less elongate than in oscines. The acrosome of Dendrocopos illustrated by Retzius (1909) (Fig. 8.21) is also a pointed cone though shorter than that of the latter orders. It does appear to be a small knob in the less clearly defined drawing by Ballowitz (1888) but is said (p. 449), like Cuculus, Columba, Gallus, Tadorna and Anas, to be “nach vorne hin verschmälert …so dass sie ein mehr nadelförmiges oder pfriemenartiges Aussehen darbietet”, thus needle- or bodkin-shaped. The nucleus of the Dendrocopos sperm is longer and more slender than in oscines, and the midpiece differs from that of the Passerida in not extending as a helix along the flagellum; the flagellum therefore differs in having a long free portion. There is no trace of an helical (“undulating”) membrane. Although spermatogenesis in Melanerpes, as in oscines, occurs from a syncytial mass, the components are irregularly arranged and it does not result in precisely aligned bundles of sperm which is diagnostic of oscines (a corvid and six passeridans were examined). The testicular sperm are motile in saline whereas those of oscines are not (Henley et al. 1978). By TEM the sperm are seen to consist of an acrosome, a densely compact nucleus and a midpiece composed of mitochondria of conventional morphology and varying number [about 6 or 7 in transverse sections in the micrograph] surrounding the axoneme (Fig. 8.26A). Although microtubules are present in the syncytium in which the spermatids develop, there is no microtubular sheath, correlating with the absence of an helical membrane. The sperm are also said to differ from those of “sauropsid” sperm, which they generally resemble, and those of oscines in lacking dense fibers peripheral to the axonemal microtubules (Henley et al. 1978) though this absence requires confirmation for all regions of the axoneme. An amorphous sheath appears to be present around the axoneme in a single micrograph (Fig. 8.26A). Fig. 8.26 Contd. ...

transversely through different levels of the components, in the region of the nucleus (N), the mitochondria (M) of the midpiece surrounding the axoneme (A), and in (S) through the tail (principal piece) posterior to the mitochondrial region and showing the amorphous sheath. Note the absence of dense fibers. PM, plasma membrane. From Henley, C., Feduccia, A. and Costello, D. P. 1978. The Condor 80: 41-48, Fig. 7. B-D. Crotophaga ani, Smooth-billed ani. Late spermatids. B. Longitudinal sections of two nuclei surmounted by acrosomes lacking perforatoria. C. Transverse sections of an acrosome vesicle, midpiece and nuclei. D. Detail of midpiece from C. AV, acrosome vesicle; M, mitochondria of midpiece; N, nucleus. Scales are approximate. From micrographs provided by Dr. Sandro Tripepi. E. Longitudinal section of midpiece of elongated spermatid. After Saita et al. (1982). Bolletino de Zoologia 49: 115-123, Fig. 12. A, axoneme; AN, annulus; DC, distal centriole; M, mitochondria.

" $ Reproductive Biology and Phylogeny of Birds Remarks. Henley et al. (1978) appear correct in recognizing the similarity of the woodpecker sperm to those of galliforms and anseriforms but it is not known whether a perforatorium, present in the latter two orders, is present. For a phylogenetic discussion see section 8.11.

8.10.2 Order Strigiformes It is remarkable that the sperm of the Strigiformes have almost entirely been neglected as objects of study. Retzius (1909) gave a drawing of that of Strix aluco, the Tawny owl (Fig. 8.21).

8.10.2.1 Strix aluco The spermatozoon of Strix aluco, illustrated by Retzius (1909) (Fig. 8.21) has, in contrast to Caprimulgus with which is was associated in an enlarged Strigiformes by Sibley and Ahlquist (1990), a morphology typical of some other Coronaves, notably charadriiforms, trogoniforms, and falconiforms, with their button-like acrosomes. The spermatozoon is particularly similar to that illustrated by SEM for Falco (Fig. 8.35), having besides the button-like acrosome, a stout moderately long, cylindrical nucleus, a short midpiece, here with about six tiers of mitochondria, and a moderately long free flagellum.

8.10.3 Order Trogoniformes 8.10.3.1 Trogon collaris Trogon sperm are known only from the light microscope account for Trogon collaris, the Collared trogon, of McFarlane (1963) (Fig. 8.27A). Trogon sperm (Fig. 8.27A) resemble those of the Charadriiformes, exemplified by the gull Larus marinus (Fig. 8.27B) in having a button-like acrosome and straight, cylindrical nucleus and midpiece. The length of the midpiece is less than half that of the nucleus (McFarlane 1963).

8.10.4 Order Cuculiformes The spermatozoon of Cuculus canorus, the European cuckoo, illustrated by Ballowitz (1888) (his Fig. 110) is straight, non-helical, with pointed acrosome much shorter than the stout cylindrical nucleus, midpiece shorter than the nucleus, and a long free flagellum. Spermiogenesis of another cuckoo Crotophaga ani, the smooth-billed ani, has been described ultrastructurally by Saita et al. (1982a, 1982b). Tripepi et al. (1991) have commented on, and illustrated the microtubules of the spermatid. Additional micrographs have been provided by Dr. Tripepi (Fig. 8.26B-D). The acrosome vesicle of the late spermatid forms a cone, with rounded tip and is more than twice as long, at 1.5 mm, as wide (Fig. 15 in Saita et al. 1982b). Less developed acrosomes are shown in Fig. 8.26B. Its rounded base rests in a shallow concavity of the tip of the long cylindrical nucleus (Fig. 8.26B). A perforatorium is absent. In a transverse section the midpiece has five mitochondria encircling the axoneme (Fig. 8.26C, D) and in longitudinal section three or four somewhat elongate mitochondria which appear to be

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Fig. 8.27 Drawings by light microscopy of bird spermatozoa. A-F. Some basic designs. A. Trogon collaris, Trogoniformes. B. Larus marinus, Charadriiformes. C. Somateria mollissima, Anseriformes. D. Claravis pretiosa, Columbiformes. E. Gallus gallus, Galliformes. F. Dendroica coronata, Passeriformes. G-K. Some variations in passerine sperm. G. Deconychura typica, Dendrocalaptidae. H. Tyrannus tyrannus, Tyrannidae. I. Corvus brachyrhynchos, Corvidae. J. Vireo olivaceus, Vireonidae. K. Cardeulis (=Spinus) pinus. Fringillidae. Relabeled after McFarlane, R. W. 1963. Proceedings of the XIII International Ornithological Congress: 91-102, Figs. 1 and 2.

" & Reproductive Biology and Phylogeny of Birds longitudinally conjoined are observed; the midpiece is terminated by a small annulus (Fig. 8.26E). The axoneme in the sections illustrated (Fig. 8.26C, D) appears to be transitional from the distal centriole. The distal centriole penetrates for a short distance into the midpiece (Fig. 8.26E). An helical membrane is absent. The principal piece has an amorphous sheath (Fig. 14, Saita et al. 1982b). Dense fibers are absent from the principal piece but there appears to be a very small density, interpretable as a minute dense fiber, on the outer aspect of each doublet in the anterior region of the midpiece (Fig. 8.26C, D). Tripepi et al. (1991) consider the arrangement of microtubules in the spermatid to be at a third level above the “reptilian” arrangement. There has been a total loss of the longitudinal manchette and only the longitudinal microtubules of the Sertoli cells are present. Phylogenetic remarks. The loss of the perforatorium is a notable development, distinguishing the cuculiform spermatozoon from that of the Galloanserae. The acrosome is also considerably shorter and the distal centriole penetrates for a much shorter distance into the midpiece.

8.10.5 Order Psittaciformes Accounts of spermatozoal ultrastructure exist for Melopsittacus undulatus, Budgerigar, Psittacinae (Humphreys 1975; Samour et al. 1986; Jamieson et al. 1995); Agapornis roseicollis, Peach-faced lovebird, Psittacinae (Jamieson et al. 1995); Platycercus elegans, Crimson Rosella (Jamieson 1999; adult in Fig. 8.3D) and Nymphicus hollandicus, Cockatiel, Cacatuinae (Jamieson et al. 1995). Additional micrographs are included here for N. hollandicus. Retzius (1909) illustrated the sperm of a Psittacus sp. by light microscopy. As shown in the accounts below, psittaciform sperm have the following characteristics: (1) a conical acrosome vesicle; rod-like perforatorium; cylindrical, highly condensed nucleus; proximal and distal centriole embedded in dense material; elongate periaxonemal mitochondrial midpiece, (2) nine dense peripheral axonemal fibers (coarse fibers), (3) they have lost the fibrous sheath around the axoneme which occurs in ratites and lower nonpasserines, (4) mitochondria with linear cristae, lacking intra- (or inter-) mitochondrial dense bodies, (5) restriction of the endonuclear perforatorial canal to the anterior region of the nucleus, (6) a short proximal and distal centriole, and (7) nuclear tip not or only very slightly penetrating the acrosome vesicle. Psittaciforms differ fundamentally from passerines and resemble galliforms, in the elongate nucleus and the much shorter acrosome relative to this, the ratio of length acrosome:length nucleus being 0.2 in Nymphicus hollandicus. The drawing of a Psittacus sperm by Retzius (1909) supports ultrastructural observations for psittaciforms, there being a short pointed acrosome, a shorter rod-like penetration of the anterior end of the nucleus, an elongate though stout nucleus many (ca 13) times the length of the acrosome, together with a

Avian Spermatozoa: Structure and Phylogeny

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short straight midpiece about 0.4 the length of the nucleus, and a long free flagellum (Fig. 8.21).

8.10.5.1

Melopsittacus undulatus

Differentiation of the acrosome in the Budgerigar spermatid has been described ultrastructurally by Humphreys (1975) and some features of the mature spermatozoon by Samour et al. (1986). The following account is taken chiefly from the account of Jamieson et al. (1995). General. The spermatozoon of Melopsittacus undulatus is a filiform cell consisting of a head region containing the nucleus and acrosomal structures, a midpiece, and tail region. Its ultrastructure is summarized, semidiagrammatically, in Fig. 8.28. The anteriorly tapering cylindrical head was estimated from both light microscopy and transmission electron microscopy to be 13 mm long. The nuclear and acrosomal lengths reported for the Budgerigar by Jamieson et al. (1995) and from the work of Samour et al. (1986) are close to those ascertained for three galliform species (Thurston and Hess 1987). The length of the entire head (acrosome + nucleus) is 8 ± 1.97 mm and its width is 0.5 ± 0.05 mm (n = 100) (Samour et al. 1986). Acrosome complex. The anteriormost region of the head consists of the acrosomal complex, which is composed of a smooth, elongated, conical acrosomal vesicle and a rod-like perforatorium lying free in the subacrosomal space (Figs. 8.28, 8.29A, C, D, G, H). The acrosomal vesicle is 1.9 mm long (n=2) (Jamieson et al. 1995), (1.42 ± 0.16 mm (n = 100) Samour et al. 1986), and terminates anteriorly in a blunt tip. Its width is greatest at its distal end. Here its diameter is 0.5 mm and its walls are ~0.15 mm thick (Fig. 8.29C, D). The perforatorium extends proximally very nearly to the tip of the spermatozoon (as in anseriforms), and distally extends within a cylindrical end nuclear canal another 0.6 mm into the anterior region of the nucleus. At its widest point, the perforatorium measures 0.2 mm (Fig. 8.29C). The contents of the acrosomal vesicle are homogeneous and of moderate electron density. The perforatorium is slightly more electron-dense, but of a less uniform composition, with irregularly spaced, electron-lucent channels penetrating it. Granular material surrounds the perforatorium within the subacrosomal space (Fig. 8.29C, D). The space between the outer membrane of the acrosomal vesicle and the plasma membrane is also occupied by granular material. This gap is ~20 nm wide at the distal end of the acrosomal vesicle. The acrosomal vesicle terminates adjacent to the proximal end of the nucleus. These two structures do not overlap or join directly. It appears that only the p1asma membrane and the shared perforatorium hold them in position. This fragile connection is considered to form a point of mechanical weakness and “decapitated” spermatids and isolated acrosomes are often found in testicular sections (Humphreys 1975). This condition contrasts with other investigated avian orders, thus giving a psittaciform synapomorphy and autapomorphy.

"! Reproductive Biology and Phylogeny of Birds

Fig. 8.28 Melopsittacus undulatus, Budgerigar. Semidiagrammatic representation of the ultrastructure of a spermatozoon from the ductus deferens (drawn by C. Tudge). From Jamieson, B. G. M., Koehler, L. and Todd, B. J. 1995. Anatomical Record 241(4): 461-468, Fig. 3. Reprinted with permission from Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc.

Avian Spermatozoa: Structure and Phylogeny

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Parrot sperm, like those of ratites and other birds, differ from reptiles in reduction of the subacrosomal material (subacrosomal cone) to a negligible amount. The psittaciform restriction of the endonuclear canal, housing the perforatorium, to the anterior region of the nucleus is an apomorphic condition (Jamieson et al. 1995) shared with other non-passerines (e.g. Galloanserae and the White-naped crane, Grus vipio). Nucleus. The nucleus, which is a gently curved cylinder (Fig. 8.29A), has an approximate length of 10 mm, a width of about 0.5 mm at its proximal end, and 0.6 mm at the nuclear-midpiece junction (Fig. 8.29C-F). The endonuclear canal (and perforatorium) occupies only about the proximal 5% of the nuclear length and is 0.2 mm wide (Fig. 8.29C, D). The chromatin is compact and electron-dense, although sporadic, small, electron-lucent areas are interspersed throughout the nucleolus. The surface is rough and the nuclear membrane is in close association with the plasma membrane (Fig. 8.29C, D, H, I). The distal end of the nucleus forms a broad, shallow, concave fossa (Fig. 8.29E, F). Midpiece, centrioles, and axoneme. The cylindrical midpiece region is 5.1 mm long (n =1) and ~ 0.6 mm wide along much of its length. At its anterior end, a basal lamina lines the concave surface of the nuclear base. Associated with this lamina is a band of granular, electron-dense material (Fig. 8.29E). Immediately posterior to the lamina is a 0.3-mm-long proximal centriole, positioned centrally, parallel to the base of the nucleus. Lying perpendicular to the proximal centriole is a 0.4-mm-long distal centriole, occupying only a very small fraction of the midpiece length. Both centrioles display the typical pattern of nine triplets of microtubules arranged in a cylinder, and both are embedded in electron-dense pericentriolar material (Fig. 8.29E, F, J, K). Whereas the center of the proximal centriole appears completely electronlucent, mottled granular material occupies the lumen of the distal centriole (Fig. 8.29J). The axoneme begins at a point ~ 1 mm along the length of the midpiece (Fig. 8.29E, F), continuous with the distal centriole, and extending the remainder of the length of the spermatozoon. It is organized according to the usual “9+2” pattern (Fig. 8.29K-M). The A microtubule of each doublet is completely circular and electron-dense and bears two dynein arms, whereas the lumen of the incomplete subunit B is electron-lucent. In the axoneme of the midpiece, associated peripherally with each doublet and in the same radius is a dense peripheral fiber (coarse fiber). In cross section, these fibers, which are approximately equal in size, are circular with a diameter of approximately 30 nm (Fig. 8.29K, L). A single layer of mitochondria surrounds the centrioles and the axoneme along the length of the midpiece. In longitudinal section, the mitochondria appear elongate elliptical and rather loosely aggregated, particularly in the centriolar region where they tend to a spherical form, with flocculent material present between them. They are apparently arranged longitudinally in a long period spiral (Fig. 8.29B, E, F). In transverse section of the spermatozoon, they

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Reproductive Biology and Phylogeny of Birds

Fig. 8.29 Melopsittacus undulatus, Budgerigar. A. Longitudinal section (LS) of the full length of the nucleus. B. LS of the full length of the midpiece. C and D. LS of the acrosomal complex and the anterior region of the nucleus. E and F. LS of the posterior end of the nucleus and part of the midpiece. Note the nine triplets of the transversely sectioned proximal centriole (E), which lies at right angles to the distal centriole, the latter forming the basal body of the axoneme. G. Transverse section (TS) of the acrosomal region through the perforatorium. H. TS nucleus through the endonuclear canal. I. TS nucleus posterior to the endonuclear canal. J. TS through the distal centriole showing granular central element and surrounding mitochondria. K. TS midpiece at the point of transition between distal centriole and axoneme. L. TS midpiece posterior to the distal centriole showing the 9 + 2 Fig. 8.29 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

"!!

appear more circular, and approximately nine are contained, tightly juxtaposed, around the axoneme (Fig. 8.29J-L). Mitochondrial dimensions are roughly 0.3 ¥ 0.15 ¥ 0.15 mm. The mitochondrial matrix is moderately electrondense and the cristae, although difficult to discern, appear to be linear or slightly curved. The estimate of 28-30 mitochondria in budgerigar sperm made by Samour et al. (1986) may be a severe underestimate as the midpiece has as many as nine mitochondria in transverse section and eight or more in longitudinal section. No annulus is discernible at the distal end of the midpiece, but the beginning of the tailpiece can be identified by a reduction in the diameter of the spermatozoon, to ~0.2 mm (Fig. 8.29B). By seemingly lacking an annulus, parrot sperm differ from those of paleognaths (e.g., Baccetti et al. 1991) and lower non-passerines including the mallard duck (Humphreys 1972), and the turkey, chicken, and Guineafowl (Thurston and Hess 1987; this study). An annulus is basal to amniotes and these non-passerines. Absence is therefore an apomorphic feature of budgerigar sperm but appears to be homoplastic relative to absence in passerines. Despite lacking an annulus, the midpiece-tail junction in parrot sperm is clearly demarcated, because at this point there is a rather abrupt narrowing of the spermatozoon. As a fibrous sheath is absent, only a narrow rim of cytoplasm separates the plasma membrane from the axonemal complex throughout the length of the axoneme behind the midpiece. (Samour et al. 1986) states the principal piece is composed of a sheath surrounding the axial filament complex but in the micrograph to which he refers only the plasma membrane closely unsheathes the axoneme. However, a narrow, 10-nm-wide ring of cytoplasm with granular inclusions separates the doublets from the surrounding plasma membrane (Jamieson et al. 1995) but has not been considered to constitute an amorphous sheath. The length of the tail (excluding the midpiece) has been estimated from light micrographs to be 53-55 mm (n = 5), for an entire sperm length of ~70-71 mm (n = 7) (Jamieson et al. 1995). The total length of the tail, including the midpiece, is given by Samour et al. (1986) as 54.31 ± 5.97 mm (n = 100). Transverse sections of the flagellum reveal the typical “9 + 2” microtubular axoneme, but the peripheral accessory fibers seen in the midpiece region do not exist (Fig. 8.29M). Fig. 8.29 Contd. ...

axoneme with peripheral dense fibers. M. TS of the axoneme behind the midpiece. Scale bars = 1 µm (C-M same scale). av, acrosome vesicle; dc, distal centriole; ec, endonuclear canal; m, mitochondrion; n, nucleus; p, perforatorium; pc, proximal centriole; pd, pericentriolar density; pf, dense peripheral fiber (coarse fiber). From Jamieson, B. G. M., Koehler, L. and Todd, B. J. 1995. Anatomical Record 241(4): 461-468, Fig. 1. Reprinted with permission from Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc.

"!" Reproductive Biology and Phylogeny of Birds Seminal glomera. The features and parameters of the ductal spermatozoa of Melopsittacus undulatus correspond very closely with those previously reported for semen samples by Samour et al. (1986) who conclusively confirmed the presence of seminal glomera in the cloaca of this species. Conventionally considered an exclusively passerine feature (Birkhead et al. 1994), the seminal glomera are situated on either side of the cloaca of male birds and are formed by convolutions of the terminal end of the ductus deferens. These sacs have been likened to the eutherian epididymis in that they serve as sites for sperm storage and appear to play a role in sperm maturation (Middleton 1972). The studies of Samour et al. (1986) and Jamieson et al. (1995) indicate that in M. undulatus no significant changes in sperm morphology occur within the seminal glomera as compared with sperm in the ductus deferens.

8.10.5.2 Nymphicus hollandicus, Platycercus elegans, Agapornis roseicollis The spermatozoa of the Cockatiel (Nymphicus hollandicus) (Figs. 8.30A,E, 8.31AN), Crimson rosella (Platycercus elegans) (Fig. 8.30B), and Peach-faced lovebird (Agapornis roseicollis) (Fig. 8.30C, D) essentially agree with the above account for Melopsittacus undulatus, with some differences in acrosome vesicle size and midpiece length. Thus a hollow conical acrosome vesicle encloses a stoutly rod-like perforatorium around which is a small or negligible amount of subacrosomal granular material. The perforatorium, which is penetrated by pale canals, extends almost to the tip of the acrosome vesicle and posteriorly for a short distance into the anterior end of the nucleus within an endonuclear canal (Fig. 8.31A, B, C). The cylindrical nucleus has a shallow concave basal fossa housing only the anterior region of the proximal centriole. This centriole lies parallel to the base of the nucleus and approximately at right angles to the distal centriole. Both centrioles are embedded in dense material. The axoneme, arising from the distal centriole, again lacks a fibrous sheath or an amorphous sheath. In the midpiece, nine coarse fibers are attached externally to the doublets in the same radii. The axoneme, with its coarse fibers is surrounded by a single layer of mitochondria, few and ovoid in cross section of the sperm but seen in longitudinal section to be slightly elongated and apparently arranged in a long period spiral. No annulus has been demonstrated. The coarse fibers do not extend behind the midpiece. The midpiece is very short in N. hollandicus (~2 mm; n=4) (Fig. 8.31D, G, K), but in A. roseicollis (Fig. 8.30D) Fig. 8.30 Longitudinal section (LS) of the acrosome vesicle, perforatorium and endonuclear canal of the spermatozoon of A. Cockatiel (Nymphicus hollandicus). B. Crimson Rosella (Platycercus elegans). C Peach-faced lovebird (Agapornis roseicollis). D. LS of the distal end of the elongate midpiece of A. roseicollis. E. N. hollandicus. Adult female. Both sexes have the crest; the only crested parrot. Photo Barrie Jamieson. av, acrosome vesicle; ec, endonuclear canal; m, mitochondrion, nucleus; p, perforatorium. A, C and D. From Jamieson, B. G. M., Koehler, L. and Todd, B. J. 1995. Anatomical Record 241(4): 461-468, Fig. 2A, C, D. Reprinted with Fig. 8.30 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

"!#

Fig. 8.30 Contd. ...

permission from Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc. B. From Jamieson, B. G. M. 1999. Pp. 303-331. In C. Gagnon (ed). Spermatozoal Phylogeny of the Vertebrata. The Male Gamete. From Basic Science to Clinical Applications, Cache River Press, Vienna, USA, Fig. 13J. A, C, and D are to the same scale.

"!$ Reproductive Biology and Phylogeny of Birds the midpiece has similar dimensions to that of the budgerigar, ~7 mm (n=1). Nuclear and acrosomal dimensions are in the same order as those of M. undulatus. They are: for N. hollandicus, length of acrosome 1.4 mm (n = 3); length of nucleus 6.5 mm (n = 2, width of nucleus proximally 0.5 mm, distally 0.6 mm; for A. roseicollis, length acrosome 1.5 mm (n=2); length nucleus 9.3 mm (n=1); width of nucleus proximally 0.5 mm, distally 0.6 mm (Jamieson et al. 1995; present study). Phylogenetic considerations. The conical acrosome vesicle; rod-like perforatorium; cylindrical, highly condensed nucleus; proximal and distal centriole embedded in dense material; and the elongate periaxonemal mitochondrial midpiece of psittaciform sperm are tetrapod symplesiomorphies. The nine dense peripheral axonemal fibers (coarse fibers) are an amniote synapomorphy; the fibers differ from those of reptiles, and particularly of mammals, in being uniform in size. Loss of the fibrous sheath around the axoneme which is present in paleognaths and lower nonpasserines is an apomorphy known elsewhere in birds only in columbiforms and passerines. Mitochondria with linear cristae, lacking intra- (or inter-) mitochondrial dense bodies are apomorphic relative to basal amniotes (Chelonia, Sphenodon, and Crocodylia). Restriction of the endonuclear perforatorial canal to the anterior region of the nucleus is an apomorphic condition shared with other non-passerines (galliforms and the white-naped crane) and crocodilians. The short distal centriole appears to be a reversal from the elongate distal centriole of crocodiles, ratites, galliforms and anseriforms. The acrosomal-nuclear junction, in which the nucleus abuts on but does not intrude into the acrosome is a synapomorphy of parrot sperm relative to other non-passerines and crocodile. It is also, homoplastically, a Fig. 8.31 Nymphicus hollandicus, Cockatiel. TEM micrographs of sperm. A-H, K. Longitudinal sections. A. Showing the entire length of the nucleus, surmounted by but not intruding into the acrosome vesicle. B. Acrosome vesicle with perforatorium. C. Acrosome vesicle and perforatorium which penetrates a considerable endonuclear canal. D. Centriolar region, showing mitochondria here grouped around the proximal centriole and the base of the nucleus. Note absence of a fibrous sheath around the axoneme. E. Developing acrosome vesicle and perforatorium in a spermatid. F. Proximal and distal centrioles and short midpiece. G. The same, with longer midpiece. H. Proximal and distal centrioles and mitochondria of midpiece also extending along base of nucleus. I. Transverse section (TS) through apex of nucleus, showing perforatorium in endonuclear canal. J. TS nucleus, showing circular profile. K. LS centriolar region and midpiece. L. TS showing small number of mitochondria grouped around axoneme in midpiece; nine uniform dense fibers are attached one to each axonemal doublet. M. TS axoneme with typical 9+2 arrangement. N. TS of an abnormal axoneme with 18 doublets and four singlets. av, acrosome vesicle; dc, distal centriole; df, dense fiber; ec, endonuclear canal; m, mitochondrion, nucleus; p, perforatorium; pc, proximal centriole. F. After Jamieson, B. G. M., Koehler, L. and Todd, B. J. (1995), Anatomical Record 241(4): 461-468, Fig. 2B. Reprinted with permission from WileyLiss, Inc., a subsidiary of John Wiley & Sons, Inc. A-E, G-N. Original.

Avian Spermatozoa: Structure and Phylogeny

Fig. 8.31

"!%

"!& Reproductive Biology and Phylogeny of Birds synapomorphy of passerine sperm but there the acrosome and nucleus are more closely contiguous. The short midpiece of N. hollandicus distinguishes this cacatuine from the four psittacines. The sperm of other cockatoos have not been investigated but this difference may prove to answer in the positive the query as to whether cockatiels are true cockatoos. For further phylogenetic considerations, see section 8.11.

8.10.6

Order Gruiformes

8.10.6.1 Fulica atra, Crex crex For the gruiform Common coot (Fulica atra) (Retzius 1909) (Fig. 8.10) depicts a spermatozoon with a needle-like acrosome much shorter than the elongate nucleus and with a short apparently spiral midpiece with approximately five gyres. In contrast, the spermatozoon of another rallid, Corncrake (Crex crex) (Fig. 8.10), appears to have a button-like acrosome and the midpiece consists of seven tiers of mitochondria grouped around the proximal axoneme but not notably spiral. Both species have a long flagellum. A single genus and species of Rallidae was examined by McFarlane (1963) but he gave no account.

8.10.6.2 Grus vipio The sperm of White-necked crane (Grus vipio) have been examined by light microscopy (LM), SEM and TEM (Asa and Phillips 1987; Phillips et al. 1987). By LM sperm are pleomorphic, with nuclei which are straight or curved or S– shaped or have droplet, spherical or irregular shapes. The acrosome appears as a spherical structure and these motile cells appear to have a simple flagellum. There is no obvious midpiece except for a small phase-dense region, of varying size and density, at the base of the flagellum. Acrosome. By TEM the acrosome (Fig. 8.32A) is described as a spherical moderately electron-dense structure. It is perhaps more accurately described as having the form of a very thick inverted U. The slightly more electron-dense structure extending from the center of the acrosome into the nucleus is clearly a well developed but fairly short perforatorium, the portion of which lying in the short endonuclear canal (anterior nuclear fossa) is approximately equal in length to the portion filling the subacrosomal space; the length of the entire perforatorium is here estimated at about 0.56 µm and the length of the acrosome vesicle 0.57 µm for a width of 0.48 µm. Nucleus. Crane spermatozoa are unique for Aves, so far as is known, in that the degree of condensation varies among spermatozoa, but in no spermatozoon does chromatin become highly compacted. Incomplete condensation of chromatin may be responsible in part for the variety of sperm head shapes (Phillips et al. 1987). From an SEM micrograph, the nucleus appears stout and moderately elongate. Midpiece. The midpiece (Fig. 8.32B) appears by TEM to consist of a cluster of small mitochondria grouped at the base of the nucleus with a length of only about 0.8 µm. It is here considered that a small density seen on each side at its

Avian Spermatozoa: Structure and Phylogeny

"!'

Fig. 8.32 Grus vipio, White-naped crane. A. Longitudinal section (LS) of the acrosome vesicle, perforatorium and short endonuclear canal. B. LS midpiece showing that the irregular mitochondria are located in an enlarged region at the base of the flagellum. C. Sperm chromatin is never highly compacted and the degree of compaction varies between spermatozoa. A distal centriole is seen in TS. D. The variability in size of the nucleus is not always a function of chromatin condensation. A. ¥ 42000. B. ¥ 49300. C. ¥ 16000. D. ¥ 16000. Relabeled after Phillips, D. M., Asa, C. S. and Stover, J. 1987. Journal of Submicroscopic Cytology. 19(3): 489-494, Figs. 3-6.

posterior end may be a small annulus. In a transverse section of the spermatozoon (Fig. 8.32C) mitochondria can be seen to surround the distal centriole. A proximal centriole is not reported or figured. Axoneme. In some transverse sections (Fig. 8.32B) there appears to be a very narrow granular layer between the 9+2 axoneme and its plasma membrane. This is possibly a vestigial amorphous sheath. However, Phillips et al. (1987) state that an amorphous sheath is absent as are outer dense fibers.

"" Reproductive Biology and Phylogeny of Birds Remarks. Phillips et al. (1987) mentions that 11 other species of crane were found to have similarly pleomorphic sperm (Russman and Harrison 1981). Gee and Temple (1978) found no correlation between the percentage of cell types in the ejaculate and fertilizing capacity. Phillips et al. (1987) considered the Grus vipio sperm to be the simplest known non-passerine sperm. It is here considered that this simplicity is probably due to reduction relative to paleognaths and, for instance, galloanserans. A perforatorium similar to that of galloanserans has been retained but the acrosome has lost its pointed form. The midpiece has become reduced and simplified relative to the other groups and the absence or weak development of the amorphous sheath and absence of dense fibers must be considered further reductions. However, the reduction of the midpiece it not constant for gruiforms as it is well developed in Fulica atra and Crex crex (Fig. 8.10). The button-like acrosome in Crex resembles that of Grus in external form and there is perhaps a possibility that the needle like acrosome depicted for Fulica by Retzius (1909) is an exposed perforatorium after loss of the acrosome vesicle. If not, gruiforms must be considered unusual in having both lanceolate and button-like acrosomes.

8.10.7

Order Charadriiformes

Harshman (Chapter 1) cites evidence that monophyly of both Pelecaniformes and Ciconiiformes is falsified by the close relationship among the pelecaniform family Pelecanidae (pelicans) and the two charadriiform families Scopidae (hamerkop) and Balaenicipitidae (shoebill). However, Charadriiformes is monophyletic when turnicids are added to the order. The suborder Charadrii of the order Ciconiiformes is therefore restored to the Charadriiformes in the present account.

8.10.7.1 Constituent families Retzius (1909) illustrated the spermatozoon of the Common guillemot (Uria aalge). It has a button-like acrosome, elongate fusiform nucleus, a midpiece about one third the length of the nucleus and with approximately seven tiers of mitochondria and a short free flagellum (Fig. 8.10). McFarlane (1963) examined the sperm of six families of charadriiforms, with a total of nine genera, by light microscopy (Fig. 8.33), though giving only a fleeting account. He found the underlying feature of the sperm of this order to be a cylindrical shape with a midpiece of slightly small diameter. The very small acrosome vesicle was button-like or subspheroidal (Fig. 8.33). Representative of the four families Recurvirostridae (stilts and avocets), Charadriidae (plovers and lapwings), Laridae (skuas, gulls and terns) and Alcidae (auks and puffins) showed only minor variations of this pattern. This pattern, with small subspheroidal acrosome, was also demonstrated for Larus canus, Common gull (Ballowitz 1888), L. fuscus, Lesser black-backed gull, (Retzius 1909) (Fig. 8.10), and Vanellus vanellus, the Lapwing (Ballowitz 1888; Retzius 1909) (Fig. 8.10).

Avian Spermatozoa: Structure and Phylogeny

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Specimens of the Scolopacidae (sandpipers, phalaropes, and allies), though true charadriiforms (see Fig. 1.2, Harshman, Chapter 1) exhibited a remarkable contrast. In scolopacids, represented by Actitus macularia, Spotted sandpiper (Fig. 8.33E), the spermatozoon has an elongate, spiral form very reminiscent of the passerines. This form was seen in Actitus and Catoptrophorus by McFarlane (1963) and was illustrated in the scolopacids Calidris (=Tringa), the Alpine dunlin, Tringa (=Totanus) ochripus, the Green sandpiper, Scolopax rusticola, the Woodcock, and in a somewhat attenuated form in Philomachus pugnax, the Ruff, by Retzius (1909) (Fig. 8.21). While regarding similarity of scolopacid and passerine sperm as convergence, McFarlane (1963) considered, perhaps not entirely reasonably, that it might indicate a more recent origin for the family than the remainder of the charadriiforms. We may note that convergence is supported by the evidence from DNA hybridization (Sibley and Ahlquist 1990) which indicates that scolopacids do not have passerine affinities but lie in a larid through jacanid clade. Within this clade Scolopacidae are distinct in grouping with Jacanidae and relatives and not with Charadriidae and Laridae which occupy a second subclade. It remains

Fig. 8.33 Variation of spermatozoa among charadriiforms. A. Fratercula arctica, Atlantic puffin. B. Gelochelidon nilotica, Gull-billed Tern. C. Himantopus mexicanus, Black-necked stilt. D. Charadrius wilsonius, Wilson’s plover. E. Actitus macularia, Spotted sandpiper. Relabeled after McFarlane, R. W. 1963. Proceedings of the XIII International Ornithological Congress: 91-102, Fig. 6.

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Reproductive Biology and Phylogeny of Birds

to be investigated what features of the reproductive tract or fertilization biology are correlated with this superficially passerine-like sperm morphology.

8.10.7.2 Jacana jacana Spermiogenesis in Jacana jacana (Jacanidae) has been described by Saita et al. (1983). Although the structure of the mature spermatozoon is not specifically treated the chief features of its morphology are evident (Figs. 8.34). Tripepi et al. (1991) have commented on, and illustrated the microtubules of the spermatid of this species. The spermatozoon or late spermatid has the typical charadriiform morphology of button-like (subspheroidal) acrosome vesicle, elongate cylindrical nucleus, and straight midpiece. As usual the proximal centriole is

Fig. 8.34 Jacana jacana. A. Longitudinal section (LS) of nucleus of advanced spermatid capped by the subspheroidal acrosome vesicle. B. Same. C. Transverse sections (TS) of nuclei and surrounding mitochondria of spermatids. D. LS spermatozoon at junction of midpiece and principal piece. E. TS of spermatozoa, showing circlet of several mitochondria and small dense fibers attached to the doublets of the axoneme. F. TS posterior region of principal piece with fibrous (amorphous) sheath less developed than further anteriorly. an, annulus; av, acrosome vesicle; ax, axoneme; df, dense fiber; fs, fibrous sheath; n, nucleus. Rearranged and relabeled after Saita, A., Longo, O. M. and Tripepi, S. 1983. Accademia Nazionale dei Lincei. (Rendiconti della Classe di Scienze fisiche, matematiche e naturali) 74: 417-430, Figs. 14-19.

Avian Spermatozoa: Structure and Phylogeny

""!

transverse to the long axis of the sperm and the distal centriole, which is slightly longer, forms the basal body in this axis. Further features which are possibly general for charadriiforms are the absence of a perforatorium, the midpiece consisting of several mitochondria arranged around the axoneme and terminating at an annulus, and the presence of an amorphous sheath around the post-mitochondrial axoneme. The arrangement of microtubules in the spermatid is as in Caprimulgus (Tripepi et al. 1991). Remarks. Persistence of the amorphous sheath, seen also, for instance, in galliforms and anseriforms, is a plesiomorphic feature but the rounded acrosome vesicle is clearly apomorphic as it is conical in those orders, in ratites and in crocodiles. The absence of a perforatorial rod is a major distinction from the sperm of the gruiforms as exemplified by Grus vipio, described by Phillips et al. (1987) but whether this is a constant difference between the two orders remains to be ascertained. There is also no evidence that G. vipio sperm possess an amorphous sheath but this also requires confirmation.

8.10.8

Order Falconiformes

Falcons and their near relatives were placed in the infraorder Falconides, within the Suborder Ciconii, of the order Ciconiiformes by Sibley and Ahlquist (1990). However, from analysis of molecular sequences placement within a ciconiiform or charadriiform assemblage does not appear acceptable. There is some morphological evidence for monophyly of Falconiformes but no molecular analyses have so far succeeded in putting all families together. Nor, however, is there strong evidence for relationship of any falconiform family to any non-falconiform family (see Harshman, Chapter 1). The only ultrastructural representation of falconiform sperm appears to be a scanning electron micrograph of spermatozoa of the peregrine falcon by Wagley (Anon. 1980) (Fig. 8.35). Despite re-establishment of falconiforms as a separate order, the morphology of the falcon sperm is consistent with placement of falcons in the Charadriiformes so far as can be judged by our limited knowledge of these. Thus the falcon sperm is very similar to the spermatozoa of the Lapwing (Vanellus) and the Gull (Larus), both charadriiforms, as depicted from light microscopy by Retzius (1912) (Fig. 8.10). Similarities are the short, domed acrosome; the stout cylindrical nucleus many times the length of the acrosome; and the midpiece consisting of a non-helical cluster of mitochondria and much shorter than the nucleus. This structure is consistent with a drawing of the whole sperm by light microscopy by Ballowitz (1888) (his Fig. 114), for the Black kite (Milvus ater). It is, however, different from other charadriiforms such as the Woodcock (Scolopax), the Dunlin (Calidris) and the Green sandpiper (Tringa), in which the acrosome is a tortuous cone, the nucleus is somewhat helical and the midpiece is more elongate (Fig. 8.10).

""" Reproductive Biology and Phylogeny of Birds

Fig. 8.35 Falco peregrinus. Scanning electron micrograph of spermatozoa. Relabeled from Anonymous 1980. Western Release Program-1980. The Peregrine Fund Newsletter. T. J. Cade and P. R. Dague (eds). 8, Fall 1980: 4. Photo L. Wagely.

8.10.9 Order Passeriformes The non-passerine sperm is generally rather plain and elongate. The passerine type has a pronounced spiral to the head region, approaching a conic helix in some forms, an enormous acrosome, often 3-4 times the length of the nucleus. Frequently an helical membrane extends some distance along the axial filament (McFarlane 1963). This succinct definition in fact characterizes the higher, non-corvidan passerines (Passeri). In suboscines and, usually, Corvida the acrosome is shorter than the nucleus and in the suboscines the ‘helical membrane’, at least in the form seen in oscine sperm, is absent.

8.10.9.1 Suborder Tyranni (Suboscines), Parvorder Tyrannida The spermatozoa of the Tyrannidae are very poorly known ultrastructurally. Species that have been investigated are Western kingbird (Tyrannus verticalis, McFarlane 1971 fide Koehler 1995); Eastern Kingbird (T. tyrannus, Feduccia 1979); Eastern wood peewee (Contopus virens, Feduccia 1979) and Great crested flycatcher (Myiarchus crinitus (=M. griseisticta), McFarlane 1971 fide Koehler 1995; Asa and Phillips 1987) but accounts are fragmentary.

Avian Spermatozoa: Structure and Phylogeny

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However, though the account was very brief, McFarlane (1971) examined the sperm of six families and 23 genera of the suborder Tyranni by light microscopy. Feduccia (1979), in his innovative phylogenetic paper on avian ear ossicles, showed that sperm of the suboscines Tyrannus tyrannus and Contopus virens (Fig. 8.36) resemble those of oscines in occurring in the testes in sperm bundles. However, the suboscine sperm differed from those of oscines in important respects indicated below (see Tyrannus tyrannus). Asa and Phillips (1987) added, for the suboscine Myiarchus griseisticta, that there are features resembling both non-passerines and oscines. Unlike most oscines, the acrosome and midpiece are shorter than the nucleus and, according to McFarlane (1963), there are no spiral head membranes or helical tail membranes in the Tyranni. However, an ‘undulating membrane’ of somewhat different construction occurs (see sections 8.10.9.2-5, below). The ratio between head length and tail length is conspicuously lower (1.3-3) than in all oscines (5.5-22) which McFarlane (1963) examined, excepting the Corvidae and the Lanidae (2.2-3), which are closely related (Christidis et al. 1996). The woodhewers (Dendocalaptidae), exemplified by Deconychura typica (Fig. 8.27G) and ovenbirds (Furnariidae) have a simple sperm that forms a spiral which completes only one revolution from the acrosome to the midpiece. A more highly coiled (helical) form occurs in the tyrant flycatchers (Tyrannidae), exemplified by Tyrannus tyrannus (Fig. 8.27H), cotingas (Cotingidae), manakins (Pipridae), and ant-thrushes (Formicariidae)

Fig. 8.36 Contopus virens, Eastern wood peewee. Transverse sections of principal piece of several sperm. Relabeled after Feduccia, A. 1979. Proceedings of the Biological Society of Washington 92(4): 689-696, Fig. 5.

""$ Reproductive Biology and Phylogeny of Birds (McFarlane 1963). Sibley et al. (1988) reduced the Cotingidae and Pipridae to subfamilial rank within the Tyrannidae; an arrangement here observed to agree with sperm structure. However, the spermatologically similar Formicariidae retained familial rank in a separate Parvorder Furnariida. In this parvorder, the Furnariidae, in the superfamily Furnaroidea, contained the Furnariinae and Dendrocalaptinae, both with the simple sperm form mentioned above, and the Formicariidae, with more helical sperm, occupied the separate superfamily Formicaroidea.

8.10.9.2

Tyrannus tyrannus

General morphology. By light microscopy the spermatozoon of Tyrannus tyrannus (Fig. 8.27H) has been shown by McFarlane (1963) to have an helical acrosome, nucleus and midpiece. The sperm is said to be similar to that of corvids but to differ in that in corvids the midpiece is not included in the spiral portion. Although this appears generally to be true, the midpiece in Corvus splendens is, at least developmentally, spiral (Bawa et al. 1990). Acrosome. Although the acrosome is shorter than the nucleus as in nonpasserines [but see also most oscine Corvida] it has lateral projections as in oscines (Asa and Phillips 1987). Nucleus. The nucleus is an helical cylinder as in oscines (Asa and Phillips 1987). Undulating membrane. Asa and Phillips (1987) showed, for Tyrannus tyrannus, that a bundle of singlet microtubules comprising what was considered the oscine-like “helical membrane” is found either completely encircling or clustered on one side of the midpiece and principal piece, the latter condition presumably thought to resemble that in oscines (see also Contopus virens, 8.10.9.5, below). Neck region. The neck region has two centrioles as in non-passerines in contrast with the single, distal centriole of oscines (Asa and Phillips 1987). Mitochondria. In some sagittal sections of the midpiece, mitochondria encircle the distal centriole but in others they appear more clustered on one side, a combination of oscine and non-passerine features (Asa and Phillips 1987). Dense fibers. In Tyrannus tyrannus, small dense fibers are present in the proximal region of the principal piece only (Asa and Phillips 1987).

8.10.9.3 Tyrannus verticalis Koehler (1995) cites some of the unpublished data of McFarlane (1971) for Tyrannus verticalis, the Western kingbird: length sperm 55 µm, length acrosome 2.5 µm, length nucleus14 µm (therefore an acrosome:nucleus ratio of only 0.18, contrasting with oscines), length midpiece 4 µm. The sperm is helical but no helical membrane is reported. There is a shallow postnuclear fossa and accessory dense fibers are present.

Avian Spermatozoa: Structure and Phylogeny

8.10.9.4

""%

Myiarchus crinitus

Koehler (1995) cites some of the unpublished data of McFarlane (1971) for Myiarchus crinitus, Great-crested flycatcher: length sperm 50 µm, length acrosome 2.5 µm, length nucleus 19.5 µm (therefore an acrosome:nucleus ratio of only 0.14, contrasting with oscines), length midpiece 3.3 µm (again much shorter than in Passerida). The sperm is helical; unusual for suboscines an ‘helical membrane’ is reported but is limited to the acrosome and is reduced in size. The mitochondria are spiral but describe only one revolution. As always in passerines, an annulus is absent. There is a shallow postnuclear fossa and accessory dense fibers are present.

8.10.9.5

Contopus virens

Although having an ‘undulating membrane’ as in oscines, the suboscine sperm, exemplified by Contopus virens (Fig. 8.36) and, according to Feduccia, that of Tyrannus tyrannus, differs in the geometry of the microtubules that surround the axoneme; the axoneme is said to be separated from the bundle of microtubules by a double plasma membrane. In addition, the microtubules are not arranged in an helical array with a mitochondrial component bound around the axoneme for a mitochondrial component is said not to be distinguishable. The undulating membrane of light microscopy in both oscine and suboscine sperm is due in oscines to an helically wound tripartite membrane in the latter (lapsus “former”) to an” undulating band” of singlet microtubules that completely surround the axoneme (Feduccia 1979) though here considered to characterize the spermatid.

8.10.10 8.10.10.1

Suborder Passeri (Oscines) Introduction to oscine sperm ultrastructure

This introduction incorporates previous accounts of oscine sperm ultrastructure (see Table 8.4) (notably Humphreys 1972; Asa and Phillips 1987; Henley et al. 1978; Vernon and Woolley 1999), the accounts from light microscopy of McFarlane (1963), and observations, particularly those on Myrmecocichla formicivora and Philetairus socius by Jamieson, Hodgson and Spottiswoode (unpublished). Henley et al. (1978) made light and TEM observations on Turdus migratorius, Pipilo erythophthalmus, Vireo olivaceus, Zonotrichia albicollis, Piranga rubra, Parus bicolor, Cyanocitta cristatus and Melanerpes carolinus but only distinguished between M. carolinus, P. erythophthalmus, C. cristatus and T. migratorius by name. Sperm bundles. Spermatogenesis in oscines (always?) occurs in a syncytial mass. Thus a multinucleate spermatid normally differentiates into uninucleate spermatozoa. In oscines a corollary of this syncytial development is that the sperm in a bundle are very regularly arranged in precise register (Fig. 8.44). In contrast, the sperm produced from the syncytial mass in the non-passerine Melanerpes are not precisely arranged. The testicular sperm of oscines are not motile in 0.9% saline whereas those of the piciform are (Henley et al. 1978).

""& Reproductive Biology and Phylogeny of Birds Head. The acrosome and nucleus constitute the head. This was found to be helical in 18 passeridan species examined by Birkhead et al. (2006) with the exception of the rounded head of the Eurasian bullfinch. The number of head gyres ranged from 2.3 in Song thrush (Turdus philomelos) to 4.6 in Sedge warbler (Acrocephalus schoenobaenus); the corvids, Carrion crow (Corvus corone) and Rook (C. frugilegus) had 3.5 and 3.6 turns. All of the 18 species illustrated by Retzius have helical heads (6 Corvida, Figs. 8.37, 8.38; 12 Passerida, Figs. 8.41, 8.42). Acrosome. The very long, tapering acrosome can be three or four times the length of the albeit short nucleus (McFarlane 1963). Some acrosome:nucleus ratios are given in Table 8.6. These may be compared with ratios for paleognaths and non-passerines in Table 8.5. For references see species cited. It is seen that the acrosome is much shorter than the nucleus in the suboscines and in typical Corvida, though longer than the nucleus in Vireo olivaceus. In contrast, most Passerida have an acrosome:nucleus ratio in excess of 1, reaching 4 in the Summer tananger. The hirundins, other than the Violetgreen swallow, and possibly the blackbird, are exceptional in having a ratio of less than 1. Table 8.5

Ratio of acrosome length : nuclear length in some paleognaths and non-passerines

Taxon Struthioniformes Struthio camelus Dromaius novaehollandiae Galliformes Coturnix japonica Coturnix chinensis Meleagris gallopavo Numida meleagris Piciformes Dendrocopos major Apodiformes Apus apus Strigiformes Strix aluco Psittaciformes Psittacus sp. Melopsittacus undulatus Nymphicus hollandicus Columbiformes Columba livia Charadriiformes Philomachus pugnax Jacana jacana Falconiformes Falco peregrinus

Common name

Acrosome-nucleus ratio (mm)

Ostrich Emu

0.17 0.14

Japanese quail Blue-breasted quail Turkey Guineafowl

0.13 0.3 ca 0.16-0.25 0.14

Great spotted woodpecker

0.09

Common swift

0.03

Tawny owl

0.03

Parrot Budgerigar Cockatiel

0.1 0.19 0.2

Domestic pigeon

0.1

Ruff Jacana

0.04 0.07

Peregrine falcon

0.23

Avian Spermatozoa: Structure and Phylogeny

Table 8.6

""'

Ratio of acrosome length : nuclear length in some suboscines and oscines

Taxon Tyranni (Suboscines) Tyrannus verticalis Myiarchus crinitus Corvida Perisoreus infaustus Corvus frugilegus Vireo olivaceus Muscicapidae-Turdidae Ficedula hypoleuca Myrmecocichla formicivora Sturnus vulgaris Turdus migratorius Turdus philomelos Turdus merula Icteridae Molothrus ater Agelaius phoeniceus Quscalus quiscala Thraupidae Piranga rubra Hirundinidae Tachycineta thalassina Hirundo rustica Riparia riparia Emberizidae Cardinalis cardinalis Passeridae Passer italiae Passer domesticus Ploceidae Philetairus socius Paridae Parus bicolor

Common name

Acrosome-nucleus ratio (mm)

Western kingbird Great crested flycatcher

0.18 0.14

Siberian jay Rook Red-eyed vireo

<0.1 <0.5 1.5

Pied flycatcher Southern ant-eater chat Starling American robin Song thrush Blackbird

1.6 2.6 1.3 2.7 1.5 0.8?

Brown-headed cowbird Red-winged blackbird Common grackle

2 1.6 1.75

Summer tananger

4

Violet-green swallow Barn swallow Sand martin

3 <1 <1

Northern cardinal

1.65

Italian sparrow House sparrow

1.3 1-1.2

Social weaver

1.2

Tufted titmouse

1.2

Of all known oscine sperm, the American robin (Turdus migratorius) examined by Henley et al. (1978), was exceptional in having a hooked acrosome; however, the acrosome crest in the Social weaver (Philetairus socius) (see below) could also be considered hooked. Acrosome keel. An helical ridge on the acrosome is reported for the corvid Corvus splendens (Bawa et al. 1990) and lateral projections or an helical membrane have been reported for the acrosome of the suboscines Tyrannus tyrannus (Asa and Phillips 1987) and Myiarchus crinitus (MacFarlane 1971, Fide Koehler 1995).

"# Reproductive Biology and Phylogeny of Birds The passeridan acrosome had formerly been regarded as an helical structure with or without an encompassing helical keel. The distinctness of this keel from the mitochondrial helix has often been unrecognized, the two, with or without a nuclear keel, being termed the ‘helical membrane’. As shown in this chapter, an helical keel (‘helical membrane’) on the acrosome but not extending onto the nucleus has in fact been described, or is apparent from illustrations, in ultrastructural works for many species: Piranga rubra (Thraupidae); Tachycineta thalassina, the Violet-green swallow (Hirundinidae); Turdus migratorius, American robin (Turdidae) (McFarlane 1971 fide Koehler 1995); Myrmecocichla formicivora, Southern ant-eater chat (Jamieson, Hodgson and Spottiswoode, unpublished); Turdus merula, Blackbird (Turdidae) and Passer italiae, Italian sparrow (Passeridae) (Furieri 1961); Sturnus vulgaris, Starling (Sturnidae) (Vernon and Woolley 1999); Lonchura striata, ‘Loverbird’ (Kondo et al. 1988); Passer domesticus, House sparrow (Passeridae) (Furieri 1961), Passer diffusus, Grey-headed sparrow (Passeridae); Philetairus socius, Social or Sociable weaver; Quelea quelea, Red-billed quelea; Euplectes orix, Red bishop; Ploceus capensis, Cape weaver (Ploceidae) (Jamieson, Hodgson and Spottiswoode, unpublished); and Parus bicolor, Tufted titmouse, (Paridae) (Koehler 1995). Furieri (1961) clearly depicts the acrosomal keel of T. merula and P. italiae as a lateral projection, differing in constitution from what is here called the acrosome core, as also shown for Myrmecocichla formicivora and Philetairus socius (Jamieson, Hodgson and Spottiswoode, unpublished). The bipartite acrosome. The acrosome, in species as far apart phylogenetically as Myrmecocichla formicivora and Philetairus socius is shown to be bipartite in nature in a new interpretation of acrosome structure (Jamieson, Hodgson and Spottiswoode, unpublished). An acrosome core is surmounted by an acrosome crest, like a stock and scion, and the core is invested by a layer which is a posterior extension of the crest, here termed the crest layer or sleeve. The acrosome helix is a lateral extension of the crest layer with or without inclusion of the acrosome core. The crest anterior to the core also shows lateral extensions which constitute a continuation of the acrosome helix. Although the crest is longer in M. formicivora than in P. socius, in both the helix of the crest proper consists of only a single lateral extension or spur whereas the acrosome core enveloped by the crest layer bears three extensions when viewed in longitudinal section. The acrosome keel thus has only one full gyre on the crest and three gyres on the core. It would be of great interest to investigate development during spermiogenesis of the crest and the spur. It is possible that the crest and its layer surrounding the core is the true acrosome vesicle and that the core is subacrosomal or perforatorial material, in which case loss of the perforatorium would not be an apomorphy of passeridans. However, the crest and its posterior layer may merely be a modification of the surface layer of an acrosome vesicle represented by the core. These alternative possibilities would easily be resolved by study of the spermatid. Nucleus. The nucleus of oscine sperm (Figs. 8.9B, 8.27K, 8.37, 8.39, 8.41, 8.55, 8.49, 8.53), twisted into an helical cylinder, is short compared with that of

Avian Spermatozoa: Structure and Phylogeny

"#

non-passerines. Its length is in the order of 3-5 µm (e.g. Koehler 1995) compared with 6.5-10 µm in psittaciforms, ca 13 µm in Struthio, ca 12 µm in Dromaius, 21 µm in Coturnix japonica, 6.4 µm in C. chinensis, 7-9 µm in Turkey, 10-21 µm in Guineafowl and Rooster, i.e. 6-21 µm in galliforms. Neck region. In contrast to the spermatozoa of non-passerines and, so far as is know, suboscines, the neck region in oscines contains only one, the distal, centriole. In all passerine species examined by Asa and Phillips (1987) excepting the White-eyed vireo, Vireo griseus, electron-dense material surrounds the neck anterior to the mitochondria of the midpiece (Fig. 8.9B), as also reported for chaffinch sperm (Furieri 1961, 1962). This dense ring is well developed in Myrmecocichla formicivora but is absent in the ploceid Philetairus socius (Jamieson, Hodgson and Spottiswoode, unpublished). Midpiece. No annulus has been detected in oscine sperm and therefore there is no sharp distinction between midpiece and principal piece (that portion of the axoneme surrounded by dense fibers). The mitochondria do not surround the axoneme in a circlet but wind in a single helical strand along what is defined as the midpiece (Figs. 8.9A, D, 8.40B, 8.46-8.49, 8.51, 8.56). The extent of the mitochondria along the tail is very variable. For instance, in the Whiteeyed vireo, as, we may add, in Corvidae (Fig. 8.37), Lanius (Fig. 8.37), and Oriolus, the mitochondria extend only a short distance along the tail, whereas in Taeniopygia guttata, Zebra finch, the mitochondrion is reported to extend along 46 µm of the 73 µm flagellum (Vernon and Woolley 1999). However, Birkhead, Pellatt et al. (2005) have shown much variation in the length of the midpiece and other sperm components in Zebra finch. They showed that in this species there is an extraordinary degree of inter-male variation in sperm design that is independent of sperm swimming velocity. A quantitative genetic study using data from over 900 zebra finches showed that sperm head, midpiece and flagellum length are heritable, that negative genetic correlations exist between sperm traits, and that significant indirect (maternal) genetic effects exist. It was hypothesized that selection on the zebra finch sperm phenotype may be low because sperm competition is infrequent in this species and that this, in combination with negative genetic correlations and maternal genetic effects, may account for the variation in sperm phenotype between males. In other Passerida, the straightened length of the helix, in mm, is at its shortest at 1.86 in the Eurasian bullfinch (Pyrrhula pyrrhula), and 12.33 in Beavan’s bullfinch (Pyrrhula erythaca) but the range in other species is 53.95 in Song thrush (Turdus philomelos) to 138.48 in Linnet (Carduelis cannabina) (Birkhead et al. 2006). The recorded number of midpiece curves (gyres) in Passerida, excepting the Eurasian bullfinch which lacks a mitochondrial helix, varies from 13.5 in Song thrush) to 29.4 in Moustached warbler (Acrocephalus melanopogon). Thus there is not a total correlation between helix length and numbers of gyres. In contrast only a single gyre is reported for the corvidan Carrion crow (Corvus corone) and Rook (Corvus frugilegus) (Birkhead et al. 2006) as is evident, with

"#

Reproductive Biology and Phylogeny of Birds

small variations in length, from illustrations by Retzius (Fig. 8.37) for these species and Magpie (Pica pica), Jackdaw (C. monedula), Siberian Jay (Perisoreus infaustus), and Red-backed shrike (Lanius collurio). After loss of the microtubular bundle in the excurrent ducts, the “helical membrane” [a term used for an helical external structure of the spermatozoon irrespective of its composition] distal to the acrosome is composed only of the mitochondrial spiral (Asa and Phillips 1987) or may be accompanied by other helical components (see below). The mitochondrial and fibrous helices. The portion of the so-called helical membrane surrounding the axoneme is what is here termed the mitochondrial helix in passeridan species. It has been shown by Jamieson, Hodgson and Spottiswoode (unpublished) that the muscicapid Myrmecocichla formicivora, a second helix, termed the fibrous helix, intertwines with at least the more proximal region of the mitochondrial helix. It is clearly the smaller structure described for Sturnus vulgaris (also a muscicapoid) by Vernon and Woolley (1999) and termed by them ‘(x)’, the larger structure being the single, helical chain of (fused) mitochondria. It was earlier described by Furieri (1961) as a small crest (i.e. keel) in Turdus merula and by Henley et al. (1978), as a fibrous component, in T. migratorius (both also muscicapoids). Thus the fibrous keel is absent at maturity not only in the ploceid Philetairus socius (Jamieson, Hodgson and Spottiswoode, unpublished) but also in the Zebra finch (Taeniopygia (=Poephila) guttata) (Fawcett et al. 1971); and Lonchura striata (Kondo et al. 1988) and is not reported for the many other described passeridan sperm with the exception, requiring confirmation, of Passer italiae where there is said to be a large fiber helically surrounding the anterior region of the axoneme in addition to the mitochondrial helix (Furieri 1961). Loss of microtubular helix. The “helical membrane” has different components in the testis and in the excurrent ducts. In testicular spermatozoa, a single bundle of microtubules winds helically along the nucleus, midpiece and principal piece. The helical bundle is said (Asa and Phillips 1987) never to extend anteriorly beyond the nucleus. However, Bawa et al. (1990) state that in Corvus splendens the microtubules extend around the acrosome, though with no supporting micrograph, other than one (Fig. 8.39E) showing what are here termed the spurs. Contrary to the views of Fawcett et al. (1971) for the nucleus, Bawa et al. (1990) and Asa and Phillips (1987) consider that the microtubular bundle may be involved in disposition of the mitochondrial array. It will be shown below, however, that in the Corvida, the microtubular bundle of the spermatid (microtubular helix of the present author) continues independently posterior of the mitochondria of the short midpiece. After their release into the excurrent ducts, oscine sperm lose their external microtubular helix (Nicander 1970b; Henley et al. 1978), although a portion of it has been reported to remain on the spermatozoa of the American robin (Henley et al. 1978), an observation requiring confirmation.

Avian Spermatozoa: Structure and Phylogeny

"#!

Fig. 8.37 Drawing by light microscopy of corvidan sperm. Pica pica, Magpie; Corvus corone, Carrion Crow. After Retzius, G. 1909. Biologische Untersuchungen, Neue Folge 14(10): 89-122 Taf XXXII, Figs. 1, 12. Corvus monedula, Jackdaw; Lanius collurio; Red-backed shrike. After Retzius, G. 1911. Biologische Untersuchungen, Neue Folge 16: 89-92 Taf XXVII, Figs. 1, 2, 24. Perisoreus infaustus, Siberian jay; Corvus frugilegus, rook. After Retzius, G. 1912. Biologische Untersuchungen, Neue Folge 17: 95-99 Taf XIV, Figs. 21, 22, 30.

Origin of helical coiling. The structure and development of the microtubular helix was elucidated by Fawcett et al. (1971). In the intermediate and later stages of “finch” (Taeniopygia (=Poephila) guttata, Zebra finch, fide Asa and

"#" Reproductive Biology and Phylogeny of Birds Phillips 1987) spermatids (Fig. 8.43B), a hundred or more microtubules occur in a single bundle to which we have referred above which has an helical course around the nucleus. The microtubules are arranged in rows (layers) that alternate with pairs of parallel membranes bounding flat cisternae. The bundle terminates anteriorly at one side of the base of the acrosome where the microtubules end in the concavity of a caplike saccule which seems to be a continuation of the outermost cistern of the microtubule bundle. This saccule, with its microtubules, is here considered to be the equivalent of the spurs seen at the base of the acrosome in Corvus splendens sperm. These authors argue against the view that the microtubules are responsible for the helical shaping the nucleus. On the other hand, they do consider them to be involved in helical shaping of the mitochondria of the midpiece. When the mitochondria first gather around the base of the flagellum in spermiogenesis they are arranged end to end parallel with the axoneme (Fig. 8.43A). Concurrently with the development of the microtubules bundle in the head region and its extension caudad, the mitochondria take on an helical configuration complementary to that of the microtubule bundle (Fig. 8.43B). Thus the axoneme is enveloped by a double helix, one element of which is mitochondrial and the other microtubular. The microtubules disappear as the spermatid matures, as noted above, leaving a single helical mitochondrial sheath with the same pitch as that of the helical nucleus (Fig. 8.43C). These authors suggest that the microtubular helix, though a transient organelle that participates in spermatid elongation, may determine the configuration of the mitochondrial sheath by imposing upon it the same pitch as that of the nucleus. The microtubules extend only around the base of the acrosome vesicle and it is considered that the acrosome acquires is helical form without their participation. Its helical form seems to be acquired as a result of asymmetry of the contents of the acrosome vesicle much as coiling of the nucleus is attributed to properties of the chromatin (Fawcett et al. 1971). Tripartite helix. Henley et al. (1978) also demonstrated the tripartite nature of the “undulating membrane” [helical membrane here termed the tripartite helix] in oscine sperm but they showed that, in addition to the basic three components of axoneme, mitochondrial sheath [mitochondrial helix] and microtubular helix, there was [at least in Turdus] a fibrous component [fibrous helix] associated with the mitochondrial sheath. The four components are shown for Turdus migratorius, the American robin, in Fig. 8.45. They examined eight oscine species, though only detailing T. migratorius and Cyanocitta cristatus, and comparing these with the piciform Melanerpes. However, their summary statement that the eight oscines possessed the tripartite structure, including the helically wound strand of mitochondria, conflicts with their suggestion that an “undulating membrane” was absent from the included corvid, Cyanocitta, and the condition for the corvidan Vireo olivaceus was not described. At maturity the tripartite helix is reduced to only the mitochondrial helix, as shown by Fawcett et al. (1971; Fig. 8.43C), with or without other components described here.

Avian Spermatozoa: Structure and Phylogeny

"##

Granular body and helix. A major addition to our knowledge of the helical components of the passerine sperm has been the demonstration by Tripepi and Perrotta (1991) in several species of the Passeroidea and a certhioid of a ‘granular body’ (GB) which consists of many diffuse granules of medium size in the early spermatid and becomes an helical structure, here termed the granular helix (GH), investing the distal [and only] centriole and the proximal region of the axoneme. Their view that the GB is constant in, and distinctive of, the Passeroidea is not here confirmed as it has not been demonstrated, for instance, in the estrildid Taeniopygia guttata (Fig. 8.43) or the ploceid Philetairus socius (Fig. 8.53) and is present in the Treecreeper (Certhia brachydactyla) now in the Certhioidea. However, in T. guttata (Fig. 8.43) the mitochondrial helix can be seen to be embedded in a granular matrix which it is here considered may be the homologue of the GB. Earlier, Humphreys (1972) had very briefly referred to a ‘granular structure’ in Canary sperm. Koehler (1995) referred to a spiral granular mass or granular body in the anterior region of the midpiece of some passerine sperm in which it forms a spiral mass located anterior to the mitochondrion, as observed in Passer domesticus, with the anterior end of the mitochondrion being a few micrometres caudal to the nucleus. This GB was also mentioned for Starling (Sturnidae), Brownheaded cowbird (Icteridae), and Common Grackle (Icteridae)but not for Great crested flycatcher (Tyrannidae), Cardinal (Emberizidae), or Red-winged blackbird (Icteridae). Further distally, at varying levels in the midpiece proper, the GH is substituted by the mitochondrial sheath [mitochondrial helix], as mentioned. It and the mitochondrial helix are invested by the transient microtubular helix which is lost in the mature spermatozoon so that the fore part of the midpiece spiral, as see in the Treecreeper (Fig. 8.56E), contains only the GH. The length of the GH varies among species but it is generally shorter than the mitochondrial helix. This pattern of spermiogenesis is said by Tripepi and Perrotta (1991) to be characterized by presence of the GB, acrosome: nuclear ratio >1, and sperm length > 100 mm. It was reported for the Wren (Troglodytes troglodytes) Troglodytidae (Fig. 8.51); Cirl bunting (Emberiza cirlus) Emberizidae (Fig. 8.56); House sparrow (Passer domesticus), now in the Passeridae; Chaffinch (Fringilla coelebs) and Greenfinch (Carduelis (=Chloris) chloris) Fringillidae, all of which are Passeroidea; and Treecreeper (Certhia brachydactyla) Certhiidae, Certhioidea. In contrast, in the Corvida, Crow (Corvus corone), Magpie (Pica pica) and Jay (Garrulus glandarius) Corvidae, Red-backed shrike (Lanius collurio) Laniidae, and the sylvioids Nuthatch (Sitta europaea) Sittidae and House martin (Delichon urbica) Hirundinidae, a GB is absent, acrosome: nuclear ratio <1, and sperm length <100 mm. However, although these claims were made for the families represented, it has been noted above that the sperm of the Violet-green swallow (Tachycineta thalassina) has a length of 285 µm.

"#$ Reproductive Biology and Phylogeny of Birds Function of helical coiling. The function of helical coiling of the sperm and of the ‘helical tail membrane’ is uncertain. Helical coiling was considered by McFarlane (1963) to intensify the rotary forward movement of the spermatozoon. He reasonably considered that the helical tail membrane (mitochondrial helix) might be instrumental in energy transfer to distal portions of the axial filament comparable to columbiform sperm, also greatly elongated, in which the midpiece extends far down the tail. Passerine sperm typically spin rapidly while seeming to remain virtually straight. Because there is no evidence for an helical wave on these flagella, other possible means have been considered whereby rotation about the local flagellar axis (self-spin) might be achieved. Sometimes, while maintaining their spinning motion, they adopt a fixed curvature and it is considered that this must be an instance of bend-transfer circumferentially around the axonemal cylinder, though the mechanism is obscure (Vernon and Woolley 1999). Axoneme. The axoneme has the typical 9+2 pattern of microtubules but in contrast to non-passerines, the outer dense fibers associated with the doublets are very prominent as shown for Thryothorus ludovicanus, Carolina wren (Fig. 8.9D), Grallina leucocephala, Magpie lark (Fig. 8.40B) Turdus migratorius, American robin (Fig. 8.45). Myrmecocichla formicivora, Southern ant-eater chat (Fig. 8.48N, O, P); Sturnus vulgaris, Starling (Fig. 8.49D, E); Philetairus socius, Social weaver (Fig. 8.53J, K); Emberiza cirlus, Cirl bunting (Fig. 8.56D) and Tree creeper (Sitta europaea) (Fig. 8.56E). They are uniform in shape at a given level of transverse section, in contrast with those of mammalian sperm, but may vary in form posteriad, for instance from comma-shaped to subcircular, as in Sturnus vulgaris. In the suboscine Tyrannus tyrannus the dense fibers are small (Asa and Phillips 1987). A systematic account of passerine sperm follows.

8.10.11 Parvorder Corvida 8.10.11.1 Taxa investigated Although several species of the Corvida have been examined for spermatozoal ultrastructure, data provided are scanty and the light microscope accounts, and particularly the illustrations, of Retzius therefore assume special importance. Species which he investigated are: Hooded crow (Corvus corone); Magpie (Pica pica) (Retzius 1909); Jackdaw (Corvus monedula) (Retzius 1911); Corvus frugilegus (Rook); and Siberian jay (Perisoreus infaustus) (Retzius 1912) (Figs. 8.37, 8.38). The earlier paper of Ballowitz (1888) also provides useful information on the Corvida: Rook, Red-backed shrike (Lanius collurio), and Golden oriole (Oriolus oriolus (=galbula)). Some data for Carrion crow and Rook are given by Birkhead et al. (2006) (Table 8.7). Brief ultrastructural accounts of spermatozoa or late spermatids exist for Corvidae, Cyanocitta cristata, Blue jay (Henley et al. 1978); Corvus splendens, Crow (Bawa et al. 1990); Grallinidae, Grallina cyanoleuca, Magpie lark

Avian Spermatozoa: Structure and Phylogeny

"#%

Table 8.7 Species examined by light and scanning electron microscopy for pairwise comparison. Slightly modified from Birkhead et al. (2006). Auk (In press). Measurements in µm. Species

Corvidae Carrion Crow Rook Turdidae Blackbird Song Thrush Sylviidae Moustached Warbler Sedge Warbler

Straightened Length Total length Number Number helix length flagellum sperm of of (µm) (µm) midpiece head (gyres) (gyres)

Corvus corone Corvus frugilegus

3.10 3.71

47.32 48.06

59.02 59.78

1.00 1.00

3.50 3.60

Turdus merula Turdus philomelos

55.75 53.95

70.82 74.12

82.82 85.22

14.00 13.50

2.50 2.30

84.34

92.04

105.03

29.40

4.20

70.21

76.62

90.01

23.40

4.60

96.76 77.23 58.16 70.98

98.01 79.53 65.73 73.64

116.13 93.48 77.17 86.19

25.30 19.20 14.00 18.60

4.20 3.70 2.50 3.00

57.27 63.67

67.01 86.52

80.32 101.30

15.60 14.40

4.00 3.60

138.47 111.50 12.33

146.70 119.44 35.48

162.16 132.79 49.11

25.80 23.90 0.00

2.10 2.70 2.60

1.86

38.78

43.96

0.00

0.00

131.46 107.63

138.68 116.90

153.63 130.97

28.60 22.80

2.80 2.80

72.77 64.53

87.51 76.87

101.56 90.13

15.00 13.90

3.00 3.10

Acrocephalus melanopogon Acrocephalus schoenobaenus Chiffchaff Phylloscopus collybita Willow Warbler Phylloscopus trochilus Sylvia atricapilla Blackcap Lesser Sylvia curruca Whitethroat Paridae Parus montanus WillowTit Parus maior Great Tit Fringillidae Carduelis cannabina Linnet Carduelis carduelis Goldfinch Pyrrhula erythaca Beavan’s Bullfinch Pyrrhula pyrrhula Eurasian Bullfinch Emberizidae Emberiza calandra Corn Bunting Yellow Hammer Emberiza citrinella Passeridae House Sparrow Passer domesticus Cape Sparrow Passer melanurus

(Jamieson 1995, 1999; present study); and Vireonidae, Vireo olivaceus, Red-eyed vireo, McFarlane 1971 fide Koehler (1995); Henley et al. 1978) and V. griseus, White-eyed vireo (Asa and Phillips 1987). Data for species of Corvidae and of other oscine families examined by Birkhead et al. (2006) are given in Table 8. 7. The Corvida are of particular interest as they stand at the base of the oscines (Sibley and Ahlquist 1990), or are paraphyletic to a (mostly) monophyletic

"#& Reproductive Biology and Phylogeny of Birds Passerida (see Chapter 1), or are considered by some to contain the Passeriformes. In either view, they may be expected to have spermatozoa displaying a near primitive morphology and ultrastructure for the oscines. From the limited information available it can be seen that the sperm of Corvida retain some characteristics already developed in at least some suboscines: the presence of a microtubular helix in the spermatid and immature spermatozoon, ensheathing at least the nucleus and proximal axoneme (Bawa et al. 1990; Jamieson 1999); and the helical form of the acrosome vesicle and nucleus (Retzius 1909, 1911, 1912; Bawa et al. 1990). They are less derived than and differ from typical Passerida in that the acrosome, although elongate, is shorter than the nucleus (excepting Vireo griseus and V. olivaceus) and the midpiece is shorter than the nucleus (Retzius 1909, 1911, 1912; McFarlane 1963; Bawa et al. 1990), whereas in Passerida at maturity the acrosome is longer than the nucleus, sometimes several times longer (with some exceptions, see Table 8.6), and the mitochondria spiral around a large portion or the greater part of the axoneme (Retzius 1909; McFarlane 1963). The sperm of the Corvidae and Lanidae are also very short in contrast to the extreme elongation which occurs in the higher passerines (McFarlane 1963). The data of Birkhead et al. (2006) (Table 8.7) for Carrion crow and Rook confirm these characteristics. In the Corvida and Lanius collurio, an helical rope winding around a great length of the axoneme in immature sperm, shown in the exquisite drawings of Retzius (1912), here exemplified, albeit from photocopies, by Siberian jay (Fig. 8.38A), and Rook (Fig. 8.38B), is independent of the short midpiece and presumably consists of microtubules only. The microtubular helix in oscines is said by Asa and Phillips (1987) not to extend around the acrosome and the statement of Bawa et al. (1990) that is does in Corvus splendens requires confirmation. These points will be further discussed in the accounts for examined species, below.

8.10.11.2 Corvus splendens Bawa et al. (1990) limited his account of the spermatozoon of Corvus splendens (Fig. 8.39) to a consideration of the microtubular helix. He concluded that the microtubules play a key role in effecting not only the elongation of the nucleus but are also responsible for endowing the corkscrew-shaped appearance of the acrosome and midpiece. This view was contrary to that of Fawcett et al. (1971) who deduced that intrinsic forces of chromatin are solely responsible for shaping of the nucleus though invoking microtubules for acrosomal shaping. The observation of Bawa et al. (1990) that the acrosome and proximal flagellum acquired an helical form after extension of the microtubular helix along them does add weight to the view of the causative effect of microtubules on spermatozoal form. The helix surrounding the acrosome, nucleus, midpiece and proximal axoneme illustrated for corvids by Retzius (1912) (Fig. 8.38) is here considered to be microtubular. The plate provided by Bawa et al. (1990) allows some further observations.

Avian Spermatozoa: Structure and Phylogeny

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Fig. 8.38 Late spermatids of Corvida. A. Perisoreus infaustus, Siberian jay. B. Corvus frugilegus, Rook. Both showing that in the Corvidae before maturity the microtubular helix extends independently far posterior of the midpiece mitochondria, as interpreted and labeled in the present study. After Retzius, G. (1912). Biologische Untersuchungen, Neue Folge 17: 95-99, Taf XIV, Figs. 24 and 31.

Acrosome. The acrosome is helical and bears a distinct spiral ridge (acrosome keel) (Fig. 8.39B, E). Two spurs visible at its base are presumably the anterior limits of the regressing microtubular helix. It is shorter than the nucleus. Nucleus. The moniliform condition of the nucleus as seen by SEM (Fig. 8.39E) and the alternating nodes seen in longitudinal section (Fig. 8.39B) are consistent with an helical form. The microtubular helix invests the helical

"$ Reproductive Biology and Phylogeny of Birds

Fig. 8.39 Corvus splendens. A. Spermatid microtubular (MT) helix extending from the nucleus (N) into the postnuclear region in the vicinity of mitochondria (M). B. Spermatid, showing electron-dense nucleus (N). Microtubules (MT) are restricted to the to the grooved region (asterisks) whereas ridges (arrows) are deficient of microtubules. Microtubules ensheath at least the base of the acrosome. C. Freezefracture replica of spermatid. Microtubules (MT) are disposed in an helical bundle. D. Microtubules of the spermatid (MT) envelope and depress the midpiece which spirals around the axoneme (AX). The mitochondrion (M) tentatively recognized here was not labeled by Bawa. E. SEM of a spermatozoon in the duct. Relabeled after Bawa, S. R., Kaur, R. and Pabst, M. 1990. Proceedings of the XIIth International Congress for Electron Microscopy. Electron Microscopy, vol 3. Biological Sciences: 52-53, Figs. 1-5.

Avian Spermatozoa: Structure and Phylogeny

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nucleus, apparently with some transient mitochondria. From analogy with a finch (Fawcett et al. 1971) it is probable that these microtubules are lost when the spermatozoon reaches the distal region of the male duct. Midpiece. In the spermatid the mitochondria spiraling around the proximal axoneme, and constituting with it the midpiece, are ensheathed by the microtubular helix. The tripartite arrangement of axoneme, mitochondria and helical microtubules is typical of developing oscine sperm.

8.10.11.3

Grallina leucocephala

Knowledge of the sperm of Grallina leucocephala, also a member of the Corvida, is limited to two micrographs of transverse sections of late spermatids (Jamieson 1999, and present study) (Fig. 8.40A, B). They give no indication of the length of the midpiece (short in Corvida) but show clearly the typical tripartite oscine arrangement in it: microtubular bundle, mitochondrion and axoneme, in centripetal sequence. Nine uniform outer dense fibers, smaller than those of Passerida, embrace the doublets of the axoneme. Presence of a helix of densely packed microtubules investing the axoneme, as in Grallina cyanoleuca, was stated by Jamieson (1999) to be a synapomorphy of passerines. This remains the case but it should be noted that the microtubules do not persist in the mature spermatozoon.

8.10.11.4

Lanius collurio

The spermatozoon of Lanius collurio, Red-backed shrike (Ballowitz 1888; Retzius 1911) (Fig. 8.37), known only by light microscopy, closely resembles that of Corvidae.

8.10.11.5

Oriolus oriolus

The spermatozoon of Golden oriole (Oriolus oriolus (=galbula)), described optically by Ballowitz (1888) (his Fig. 63) is also closely similar to that of Corvidae.

8.10.11.6

Vireo

Limited ultrastructural information is available for the corvidans Vireo olivaceus, the Red-eyed vireo (McFarlane 1971 fide Koehler 1995; Henley et al. 1978) and Vireo griseus, the White-eyed vireo (Asa and Phillips 1987). Although included in the study by Henley et al. (1978) the sperm was not specifically described. General morphology. A brief light microscope account for Vireo olivaceus is given by McFarlane (1963) (Fig. 8.27J). He states that a ribbon-like membrane extends from the nucleus to the acrosome and another helical membrane extends a short distance down the tail. It is not stated whether the latter membrane is an extension of the midpiece, and this is not clear in the illustration but it seems likely that it is a purely microtubular helix of an incompletely mature spermatozoon as in the Corvidae. Koehler (1995), drawing from the unpublished thesis of McFarlane (1971), gives the following apparently ultrastructural data for V. olivaceus. Sperm

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Reproductive Biology and Phylogeny of Birds

Fig. 8.40 Grallina leucocephala, Magpie lark. Late spermatids in the testis. A. Transverse section (TS) showing a section of the microtubular helix on one side of Fig. 8.40 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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length 80 µm, acrosome length 8.0 µm, nuclear length 5.5 µm (giving a ratio of 1.5 more characteristic of Passerida), midpiece length 10 µm, shape helical, ‘helical membrane’ [helical acrosome keel] restricted to the acrosome, anterior and posterior nuclear fossae absent, accessory dense (axonemal) fibers present’, one helical mitochondrion present.

8.10.12 8.10.12.1

Parvorder Passerida Introduction

Distinctive features of the Passerida are the persistence of a mitochondrial helix extending for long distances around the axoneme and elongation of the acrosome so that it is (with some exceptions, see Table 8.6) longer than the nucleus. The latter feature is also seen in the corvid Vireo olivaceus. The passeridan sperm type (as it is here termed), with helical, pointed acrosome, helical nucleus, shorter than the acrosome, and mitochondrial helix extending far along the tail is well demonstrated in the drawings of Ballowitz (1888) and particularly of Retzius (1909) (Figs. 8.41, 8.42) and McFarlane (1963) (Figs. 8.27K, 8.50) (though the mitochondrial nature of the posterior helix could not be demonstrated at the time) in the following families and species: Muscicapidae: Muscicapa striata (=Muscicapa grisola), Spotted flycatcher (Ballowitz 1888); Turdus philomelos (= musicus), Song thrush (Fig. 8.41); Luscinia luscinia (=Aedon luscinia), Thrush nightingale; Ficedula hypoleuca (=Muscicapa atricapilla), Pied flycatcher (Retzius 1909) (Fig. 8.42). Sturnidae: Sturnus vulgaris, Starling (Retzius 1909) (Fig. 8.42). Sylviidae: Phylloscopus sibilatrix (=Phylloscopus sibilator), Wood warbler (Retzius 1909) (Fig. 8.42). Hirundinidae: Hirundo rustica, Barn swallow (Ballowitz 1888; McFarlane 1963); Chelidon urbica, House martin (Ballowitz 1888); Petrochelidon pyrrhonota, Cliff swallow; Riparia riparia, Bank swallow; Iridoprocne bicolor, Tree swallow (McFarlane 1963) (Fig. 8.50). Alaudidae: Alauda arvensis, Skylark (Retzius 1909) (Fig. 8.42). Fringillidae: Fringilla coelebs, Chaffinch (Ballowitz 1888; Retzius 1909) (Fig. 8.41); Carduelis spinus (=Chrysomitris spinus) (Retzius 1909) (Fig. 8.41), Cardeulis (=Spinus) pinus, Pine siskin (McFarlane 1963); Carduelis chloris

Fig. 8.40 Contd. ...

the condensing nucleus. B. TS of four spermatids. 1) through the nucleus and ensheathing microtubular helix. 2) and 3) TS through a continuous (?) midpiece mitochondrion invested on one side by the microtubular helix and in turn ensheathing the axoneme. The nine prominent outer dense fibers of the axoneme are well developed and are uniform in size and shape. 4) TS of the anterior end of the midpiece through the (distal) centriole. Original.

"$" Reproductive Biology and Phylogeny of Birds

Fig. 8.41 Spermatozoa of Passerida by light microscopy. Turdus philomelos, Songthrush; Fringilla coelebs, Chaffinch; Passer domesticus, House sparrow, also showing sperm bundle; Carduelis chloris, Greenfinch; Carduelis spinus, Siskin. After Retzius, G. 1909. Biologische Untersuchungen, Neue Folge 14(10): 89-122 Taf. XXXIII Fig. 1, XXXIV Fig. 3, XXXV Figs. 1, 9, XXXVI Figs. 1, 12.

Avian Spermatozoa: Structure and Phylogeny

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Sturnus

Fig. 8.42 Spermatozoa of Passerida by light microscopy, continued: Alauda arvensis, Skylark Phylloscopus sibilatrix, Wood warbler Anthus spinoletta, Rock pipit; Ficedula hypoleuca, Pied flycatcher Emberiza citrinella, Yellowhammer; Sturnus vulgaris, Starling; Luscinia luscinia, Nightingale. After Retzius, G. 1909. Biologische Untersuchungen, Neue Folge 14(10): 89-122 Tafel XXXVII, Figs. 1, 3, 5, 7, 9, 13, 15, 17, 19, 20.

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Fig. 8.43 Three successive stages in differentiation of the mitochondrial sheath of the finch sperm tail. (The example in A has an abnormal double flagellum but the relations are the same as in a normal spermatid.) From Fawcett, D. W., Anderson, W. A. and Phillips, D. M. 1971. Developmental Biology 26: 220-251, Figs. 30-32. With permission from Elsevier.

Avian Spermatozoa: Structure and Phylogeny

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%$(=Chloris chloris), Greenfinch (Fig. 8.41); Emberiza citrinella, Yellowhammer (Retzius 1909) (Fig. 8.42). Passeridae: Passer domesticus, House sparrow (Figs. 8.41, 8.52C); Anthus spinoletta (=Anthus obscurus), Rock pipit (Retzius 1909) (Fig. 8.42). Additional families for which the passeridan sperm type is implied or stated by McFarlane (1963) are: Paridae (1 gen., 4 spp.); Sittidae (1 gen., 2 spp.); Troglodytidae (2 gen., 2 spp.); Mimidae (3 gen., 3 spp.); Bombycillidae (1 gen., 1 sp.); Coerebidae (1 gen., 1 sp.); Parulidae (13 gen., 27 spp.); Ploceidae (1 gen., 1 sp.); Icteridae (8 gen., 8 spp.); and Thraupidae (7 gen., 8 spp.) Passeridan families for which additional details are available will now be considered.

8.10.12.2

Muscicapoidea, Muscicapidae (and Turdidae)

Descriptions of muscicapid sperm by light microscopy are: Muscicapa striata (=Muscicapa grisola) (Ballowitz 1888); Turdus philomelos (= musicus), Song thrush (Fig. 8.41); Luscinia luscinia (=Aedon luscinia), Thrush nightingale; Ficedula hypoleuca (=Muscicapa atricapilla), Pied flycatcher (Retzius 1909) (Fig. 8.42). Harshman (Chapter 1) reviews strong evidence that Turdidae and Muscicapidae are sister groups. They are classical passeridan sperm with the exception that the acrosome is not always longer than the nucleus; some acrosome:nucleus ratios, including ultrastructural accounts, are estimated here as about 2.7 in Turdus migratorius, 2.7 in Myrmecocichla formicivora, 1.6 in Ficedula hypoleuca and 1.5 in Turdus philomelos but only 0.8 in T. merula. The amplitude of the nuclear helix is unusually large in Muscicapa striata (=Muscicapa grisola) (Ballowitz 1888). Ultrastructural accounts of muscicapid sperm are limited to a TEM whole mount showing the anterior end of the sperm of Turdus grayi, Clay-colored robin (McFarlane 1963); a detailed account for T. merula, Blackbird (Furieri 1961); brief reference to T. migratorius, American robin (Henley et al. 1978); and details given here for the Southern ant-eater chat, Myrmecocichla formicivora.

8.10.12.3

Turdus migratorius

The sperm bundles of oscines, as exemplified by Turdus migratorius, have been discussed above. A micrograph of sperm bundle is illustrated in Fig. 8.44 and a section through a bundle in Fig. 8.45. Negative staining of the helical membrane is illustrated in Fig. 8.46 (Henley et al. 1978). Some data are given by McFarlane (1971 fide Koehler 1995). Dimensions are: length spermatozoon 70 µm; acrosome 6.7 µm; nucleus 2.5 µm; midpiece 46 µm. The spermatozoon is helical but only the acrosome bears the so called helical membrane. There are no anterior or posterior nuclear fossae. Dense fibers are present and the midpiece is helical and large. It is here considered that the helical membrane restricted to the acrosome is the same structure as the small crest (here termed the helical acrosome keel)

"$& Reproductive Biology and Phylogeny of Birds

Fig. 8.44 Turdus migratorius, the American robin. Bright-field light micrograph of a single feulgen-stained sperm bundle. The sperm nuclei are still embedded in a common cytoplasmic mass and the remaining parts of the spermatozoa are in sharp register with one another. Relabeled after Henley, C., Feduccia, A. and Costello, D. P. 1979. The Condor 80: 41-48, Fig. 2.

on the acrosome of Turdus merula and Myrmecocichla formicivora (see below). The large amplitude of the helical midpiece is presumably due to presence of a fibrous keel as demonstrated in the latter species by Furieri (1961).

8.10.12.4

Turdus merula

General morphology. Furieri (1961) provides useful diagrammatic sketches of the whole mature spermatozoon accompanied by corresponding cross sections (Fig. 8.47A). The spermatozoon has a length of 85 µm, of which the conical acrosome occupies 4 µm and the cylindrical nucleus 5 µm. Acrosome. The acrosome is solid and somewhat electron-dense, contained in the plasma membrane. It is circular in cross section but bears a small crest

Avian Spermatozoa: Structure and Phylogeny

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Fig. 8.45 Turdus migratorius, American robin. TEM of a transverse section through a bundle of spermatozoa at a level posterior to the region of the nuclei. Each spermatozoon is invested in a plasma membrane (PM) and has dense fibers (DF) surrounding a central axoneme (A). There is a single mitochondrion (M) in each and the plane of section through the sheath of singlet microtubules (SI MT) is almost exactly the same for each. Part of the residue of the cytoplasmic mass (CM) in which the bundle originated is peripheral to and between the spermatozoa. There is a fourth component (IV), roughly triangular in section, of unknown nature [probably the fibrous helix] next to the mitochondrion. From Henley, C., Feduccia, A. and Costello, D. P. 1979. The Condor 80: 41-48, Fig. 5.

[helical acrosome keel], 0.1-0.2 µm high and a little wider at its base. It is clear from Furieri’s drawing that this crest does not continue onto the nucleus and therefore that the fibrous keel (as it is here termed) on the midpiece is a separate entity. In longitudinal section the acrosome is a solid, helical cone. Superficially the acrosome and nucleus are gently curved, the acrosome convexly, and the nucleus concavely. Nucleus. Following the acrosome, the nucleus is an helical cylinder, the chromatin of which is strongly electron-dense but contains vacuoles. The two ends of the nucleus are concave, slightly at the apex, more strongly caudally. Axoneme and appurtenances. The tail is inserted into the basal nuclear fossa. On one side of the flagellum is the commencement of an helical structure [mitochondrial and fibrous helix] that is wrapped around the greater part of the flagellum while on the other side there is an elongated formation, parallel to the flagellum and containing fine amorphous trabeculae, the nature of

"% Reproductive Biology and Phylogeny of Birds

Fig. 8.46 Turdus migratorius, American robin. TEM of negatively stained spermatozoa. A. The investing plasma membrane has been digested away revealing the three components: straight axoneme (A), helical strand of mitochondria (M) and helical array of singlet microtubules (MT). Artifacts (B). B. The same three components in a slightly different orientation with respect to one another. Note the complex interweaving of the microtubules at the arrow. Relabeled after Henley, C., Feduccia, A. and Costello, D. P. 1979. The Condor 80: 41-48, Fig. 6.

which is unclear. The axoneme has the usual 9+2 arrangement and each of the nine doublets is reinforced peripherally by a large dense fiber. These fibers are assumed to be responsible for limiting the flexibility of the axoneme. The helix consists of two parallel superimposed structures. consisting of a mitochondrion overlain by a large cytoplasmic fiber [fibrous helix] of similar appearance but larger which extends far along the flagellum. The mitochondrion embraces about half the circumference of the flagellum. The cytoplasmic fiber has the form of an isosceles triangle in transverse section and occupies about two thirds of the width of the mitochondrion on which it is superimposed. The cristae of the mitochondrion are digitiform and orientated orthogonally relative to the long axis of the flagellum, disposed in quincunx (see further details in Furieri 1961).

Avian Spermatozoa: Structure and Phylogeny

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Fig. 8.47 Diagram of the whole spermatozoon with corresponding transverse sections. A. Turdus merula, blackbird. B. Passer italiae, Italian sparrow. a, acrosome; b, nucleus; c, transverse section (TS) through the tail, showing the flagellum, 9 dense fibers, the helical crest formed, basally, by a mitochondrion to which, in T. merula, is appended a large triangular cytoplasmic fiber [fibrous helix]; d, TS of the tail at the end of the helical crest which here consists only of the mitochondrion; e, TS of the endpiece. After Furieri, P. 1961. Archivio Zoologico Italiano (Napoli) 46: 123-147, Figs. 32 and 33.

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Reproductive Biology and Phylogeny of Birds

Further posteriorly, the keel formed by the dual helix progressively diminishes until the flagellum is naked. The peripheral dense fibers of the axoneme also diminish and disappear. Remarks. It is apparent that of the three components of the helical membrane identified by Henley et al. (1978) in oscine sperm, only the fibrous keel and the mitochondrial helix persist in the mature Turdus merula spermatozoon, the microtubules (presumed to be present in the spermatid) being lost.

8.10.12.5

Muscicapidae, Myrmecocichla formicivora

A study of the spermatozoon of Myrmecocichla formicivora, the Southern anteater chat, has clarified hitherto imperfectly understood aspects and revealed previously unrecognized features of passerine spermatozoon ultrastructure (Jamieson, Hodgson and Spottiswoode, unpublished). This filiform spermatozoon demonstrated par excellence the passeridan features of an acrosome longer than the nucleus and great prolongation of the midpiece as a single mitochondrion wound helically around much of the length of the axoneme. A new finding is the bipartite nature of the acrosome. Acrosome. The acrosome is an helical structure 2.7 times the length of the nucleus which it surmounts (Fig. 8.48L). It is inserted into the nucleus, at the acrosomal-nuclear junction (Fig. 8.48F, G, H), in an asymmetrical fossa. The acrosome is bipartite. Its distal, longer portion, consists of an electron-dense gently helical column of three gyres, here termed the acrosome core, ca 5 µm long, which bears a prominent keel seen in profile as three prominent spurs. The dense core is drawn out towards the keel as can be seen in longitudinal (Fig. 8.48F, G, L) and in transverse section (Fig. 8.48K). The proximal (anterior) portion, here termed the acrosome crest, consists of a narrower spirally angular electron-pale shaft, 3 µm long, tapering to a narrow tip (Fig. 8.48L). Its substance is also drawn out towards its angular projections, as seen in longitudinal (Fig. 8.48F, L) and transverse (Fig. 8.48J) section. Although the acrosome keel is limited to the region of the acrosome core, it consists of electron-pale material which is continuous with that of the acrosome crest and must be considered to be part of the crest. The acrosome crest abuts on the core at an oblique very slightly concave junction (Fig. 8.48F, L). The morphological and developmental implications of the bipartite constitution are explored above in section 8.10.10.1. Nucleus. The nucleus, like the acrosome core, is strongly electron-dense. It forms a very slightly sinuous stout cylinder (Fig. 8.48L), 2.9 µm long, of subcircular transverse section (Fig. 8.48H). Its greatest width, 0.9 µm, is shortly below the acrosomal nuclear junction. Its base closely abuts the pericentriolar dense ring (Fig. 8.48A, B, F, I, L) against which it forms a slight convexity or shallow, slightly protuberant double concavity. Centriolar complex. There is no proximal centriole. The distal centriole is surrounded by and fused with a dense ring outside which is a ring formed by the proximal end of the mitochondrion of the midpiece (Fig. 8.48D). At least

Avian Spermatozoa: Structure and Phylogeny

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some of the centriolar microtubules have been shown to form triplets (Fig. 8.48M). It is short and appears to be penetrated by the two central axonemal singlets or material continuous with these. At its junction with the axoneme, the dense ring has given way to 9 large dense fibers, but it is still encircled by the mitochondrial ring (Fig. 8.48N). Helical components of the axonemal region. Longitudinal sections of the base of the nucleus and adjacent centriolar complex and midpiece reveal two components spiraled around the axoneme: a very elongate mitochondrion which proximally forms the continuous mitochondrial ring and a strongly electron dense component here termed the fibrous helix. Dense fibers are also seen in glancing longitudinal profiles encircling the axoneme (Fig. 8.48A, B, F, I). Further distally, for the greater length of the axoneme, the only helical component is the mitochondrial helix (Fig. 8.48C). Mitochondria. As noted, a mitochondrial ring encircles the junction of the distal centriole and the axoneme, the latter with its large dense fibers (Fig. 8.48N). This mitochondrial ring is continuous with the single, extremely elongate mitochondrion which spirals along the axoneme for the greater part of the length of the latter. The course of this mitochondrial helix is seen in longitudinal section in Fig. 8.48A, B, C, F, I. In transverse section the mitochondrion is seen to accompany the portion of the axoneme which has large dense fibers where initially it lies external to the helical fiber (Fig. 8.48O) and more distally, in the absence of the helical fiber, is in direct contact with the dense fibers (Fig. 8.48P). Fibrous helix. A well developed electron-dense helix intervenes between the mitochondrial helix and the axoneme in the proximal region of the latter, as seen in longitudinal (Fig. 8.48A, B, I) and transverse section in which its crescentic form is seen (Fig. 8.48O). Axoneme. The axoneme has the conventional 9+2 arrangement of microtubules. For much of its length each doublet is accompanied by a dense fiber which is circular in cross section except for a small prolongation which joins each A microtubule near the junction of the latter with the B subtubule (Fig. 8.48O, P). The dense fibers greatly reduce in size distally (Fig. 8.48Q). Judging from the small number of transverse sections, the endpiece of flagellum is short, lacking dense fibers and mitochondrial helix (Fig. 8.48R). The extreme posterior end of endpiece has a disrupted arrangement of doublets and singlets (Fig. 8.48S). Remarks. Examination of the sperm of Myrmecocichla formicivora (with the ploceid Philetairus socius) allowed a new interpretation of the structure of the passeridan acrosome and clarification of the structure of the so-called helical membrane (see section 8.10.10.1 above).

8.10.12.6

Sturnidae, Sturnus vulgaris

The spermatozoon of a further muscicapoid, Sturnus vulgaris has been briefly characterized by Koehler (1995) and Vernon and Woolley (1999). Koehler

"%" Reproductive Biology and Phylogeny of Birds

Fig. 8.48 Myrmecocichla formicivora. TEM of spermatozoon. A, B, F, I. Longitudinal section (LS) of the base of the nucleus and adjacent centriolar complex, midpiece and anterior axoneme. Note two components spiraled around the axoneme: a very elongate mitochondrion which proximally forms a continuous ring and a strongly electron dense component here termed the fibrous helix. Dense fibers are also seen encircling the axoneme. C. LS of the longest portion of the flagellum around Fig. 8.48 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

"%#

gives a sperm length of 50 µm, acrosome length 2.5 µm and nuclear length 3.0 µm; the sperm is helical, has an helical membrane, and has granular material at the midpiece. The passeridan form of the sperm is well illustrated by Retzius (1909) for Sturnus (Fig. 8.42) but it will be noted that a departure from the usual passeridan condition is the relatively short acrosome, though longer relative to the nucleus than in non-passerines. Although Koehler gives its absolute length as shorter than the nucleus in the drawing by Retzius it appears that the acrosome:nucleus ratio in Sturnus is about 1.3. Other characteristics of passeridan sperm which it displays are the helical forms of the sperm and the spiral and very extensive midpiece. The brief account of Vernon and Woolley (1999) is particularly useful as it correlates external morphology as revealed by SEM with transverse sections by TEM. The acrosome, nucleus and the single chain of (fused) mitochondria all contribute to giving the proximal part of the cell an helically grooved surface (resembling a carpenter’s auger bit). The helical groove is sinistral, with approximately four gyres on the sperm head and 14 gyres on the midpiece, where the mean pitch is 3.3 µm (Fig. 8.49A-C). In the neck region, and for two gyres distally an extra helical structure is present (Fig. 8.49B,D). The sperm head is 10.3 µm long. The flagellum is 73.4 µm long, with the mitochondrion wound around the proximal 46 µm. There are nine uniform dense fibers around the axoneme (Fig. 8.49D, E). These fibers diminish distally, where they become round instead of comma-shaped in transverse Fig. 8.48 Contd. ...

which spirals the mitochondrial helix. D. Transverse section (TS) of the distal (and only) centriole, surrounded by a dense ring and the mitochondrial ring. E. LS of the nucleus surmounted by the acrosome. The acrosome is bipartite, consisting of a long, electron-dense acrosome core, which bears a prominent keel, and a distal spirally angular acrosome crest. The acrosome core fits into an oblique fossa at the tip of the nucleus, more clearly seen in G. H. TS nucleus. J. TS acrosome crest. K. TS acrosome core through helical keel. L. LS of the entire length of the acrosome crest and core and of the short nucleus, followed by a glancing section of the mitochondrial circlet. M. Detail of D, showing the triplets of the distal centriole. N. TS mitochondrial ring encircling junction of distal centriole and axoneme with large dense fibers. O. TS near proximal end of axoneme, showing 9+2 pattern with 9 dense fibers, crescentic section of fibrous helix and, external to this, the mitochondrion. P. TS axoneme with dense fibers and section of the mitochondrial helix. Q. TS distal region of the flagellum with no mitochondrial helix and reduced dense fibers. R. TS endpiece of flagellum lacking dense fibers and mitochondrial helix. S. Extreme posterior end of endpiece with disrupted arrangement of doublets and singlets. acr, acrosome crest; ac, acrosome core; ak, acrosome keel; anj, acrosome-nuclear junction; ax, axoneme; dc, distal (only) centriole; df, dense fiber; dr, dense ring around centriole; fh, fibrous helix; m, mitochondrion; mh, mitochondrial helix; n, nucleus. From Jamieson, Hodgson and Spottiswoode, unpublished.

"%$ Reproductive Biology and Phylogeny of Birds

Fig. 8.49 Sturnus vulgaris, Starling. A. SEM of the sperm head (h) and proximal flagellum. The surface has the form of a sinistrally helical keel that is continuous from the acrosome, through the nucleus and into the midpiece. Bar = 1 µm. B. From the neck and into the proximal two gyres of the midpiece, the mitochondrial helix (m) is accompanied by another helical structure (x), as shown also in D. Bar = 1 µm. C. The remainder of the midpiece; the helical keel is now simply mitochondrial as shown also in E. D. TEM of a transverse section (TS) through the proximal midpiece, corresponding to B. There is a mitochondrial helix (M) and another helix (x) of unknown origin. The axoneme is surrounded by nine accessory dense fibers. Bar = 0.25 µm. E. TS of more distal midpiece, corresponding to C. Bar = 0.25 µm. From Vernon, G. G. and Woolley, D. M. 1999. Cell Motility and the Cytoskeleton 42: 149-161, Figs. 16 and 17.

section; they are absent from the distal 7-8 µm of the axoneme. Freeze-etch replicas reveal that the outer dynein arms are typical for vertebrate spermatozoa (Vernon and Woolley 1999). Although the spermatozoon is

Avian Spermatozoa: Structure and Phylogeny

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described as grooved it is clear that the grooving is the complement of two, or more distally one, rope-like helical structures investing the axoneme. The larger of these is the mitochondrial helix and the smaller is a structure of unknown origin (x in Fig. 8.49B and D). This smaller helix is clearly the homologue of the fibrous keel seen in, for instance, Turdus merula (Fig. 8.47A) and described here for Myrmecocichla formicivora (Fig. 8.48). It is possible that it is derived from the transient microtubular helix characteristic of passeridan spermatids.

8.10.12.7

Sylvioidea, Hirundinidae, Petrochelidon, Hirundo, Riparia, Iridoprocne and Sylviidae

McFarlane (1963) used the Hirundinidae as an example of quantitative variation in sperm morphology within a family, for the four genera Petrochelidon, Hirundo, Riparia and Iridoprocne (Fig. 8.50). While some overlap occurs in comparative lengths of the head and midpiece (their combined length), the total lengths of the sperm of the four genera are completely different. The exceedingly great length of the helical element around the axoneme, leaving only a small fraction naked as the endpiece, is shown for Hirundo rustica by Ballowitz (1888) (his Fig. 96).

Fig. 8.50 Variation of sperm in the family Hirundinidae. A. Petrochelidon pyrrhonota. B. Hirundo rustica. C. Riparia riparia. D. Iridoprocne bicolor. E. Full view of spermatozoon of Iridoprocne bicolor; an helical membrane, not apparent in the drawing, extends to within 6 µm of the tail. From McFarlane, R. W. 1963. Proceedings of the XIII International Ornithological Congress: 91-102, Fig. 3.

"%& Reproductive Biology and Phylogeny of Birds Koehler (1995) cites the unpublished TEM description of the sperm of Tachycineta thalassina, the Violet-green swallow, by McFarlane (1971). It reaches the great length of 285 µm. The acrosome is 13.5 µm long and the nucleus is much shorter, at 4.5 µm. The sperm is helical but the helical membrane [probably merely an helical keel] is said to be limited to the acrosome. There is an anterior nuclear fossa, 0.7 µm deep, and a posterior fossa is absent. Only a distal centriole is present. One or two mitochondria are present of which the long one is helical but the length of the midpiece is not given. Dense peripheral fibers are present at the anterior end of the axoneme. As in all investigated passeridans, a perforatorium and an annulus are absent. Dimensions and other morphological data are given by Birkhead et al. (2006) (Table 8.7) for the sylviids Moustached Warbler (Acrocephalus melanopogon); Sedge Warbler (Acrocephalus schoenobaenus); Chiffchaff (Phylloscopus collybita); Willow Warbler (Phylloscopus trochilus); Blackcap (Sylvia atricapilla) and Lesser Whitethroat (Sylvia curruca).

8.10.12.8

Certhioidea: Troglodytidae and Certhiidae

Troglodytidae. Aspects of the ultrastructure of the spermatid of the Wren (Troglodytes troglodytes) have been briefly described by Tripepi and Perrotta (1991) (Fig. 8.51A, B). In this important paper these authors recognize, for the Wren and several other species, a ‘granular body’ (GB) the nature and development of which have been discussed above in section 8.10.10.1. In the advanced spermatid (Fig. 8.51B), the helix is formed internally by the GB, which coils around the proximal axoneme and its dense fibers, and externally by the transient microtubular bundle. In the remainder of the midpiece, the GB is substituted by the mitochondrial sheath [mitochondrial helix]. TEM sections of the sperm of the Carolina wren (Thryothorus ludovicanus) have been illustrated by Asa and Phillips (1987) (Fig. 8.9B, D). The micrographs show the acosome with a keel, the spiral nucleus, and the 9 + 2 axoneme with large dense fibers external to which is the mitochondrial helix. Electron-dense material of the neck possibly represents a granular helix. Certhiidae. For the Treecreeper (Certhia brachydactyla) Tripepi and Perrotta (1991) have demonstrated the presence of a granular helix persisting after regression of the microtubular helix (Fig. 8.56E). The granular helix may, therefore, be characteristic of the Certhiidae in which the subfamily Troglodytinae is sometimes included.

8.10.12.9

Passeroidea, Thraupidae, Piranga rubra

The spermatozoon of Piranga rubra, Summer tananger, is 170 µm long, with an acrosome 12 µm long, nucleus much shorter, at 3 µm, and a very long, 146 µm midpiece (McFarlane 1971, fide Koehler 1995). The spermatozoon is helical but the ‘helical membrane’ (here considered to be only the helical keel) is restricted to the acrosome. Anterior and posterior nuclear fossae are absent.

Avian Spermatozoa: Structure and Phylogeny

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Fig. 8.51 Wren (Troglodytes troglodytes). A. Early spiralized spermatid. The granular body encircles the proximal part of the axoneme. Mitochondria are accumulating in a spiral array. B. Elongating spermatid. The granular body has become a granular helix, distal to which is the mitochondrial helix. Both of these are invested by the transient microtubular helix. A, relabeled after Tripepi, S. and Perrotta, E. (1991). Pp. 1021-1023. In Baccetti B. (ed.), Comparative Spermatology 20 Years After. Serono Symposia Publications, vol. 75. Raven Press, Rome. Fig. 2. B, micrograph courtesy of Dr. Sandro Tripepi.

8.10.12.10

Passeroidea, Passeridae, Passer domesticus

Passeridae are often reduced to subfamilial rank within the Ploceidae but are here retained. Nevertheless, they lie within a ploceid clade (see Chapter 1, Fig. 2.2B). Koehler (1995) tabulates features of the spermatozoon of Passer domesticus, the House sparrow, derived from SEM and TEM examinations and gives an SEM of the acrosome (Fig. 8.52C). A detailed account of spermiogenesis in this species is given by Góes and Dolder (2002) which is summarized in Chapter 7 of this volume. It does not, however, recognize the bipartite nature of the passerine acrosome revealed in the present chapter for Myrmecocichla and Philetairus. Birkhead et al. (2006) present a scanning electron micrograph of the head of a House sparrow sperm, noting the helical head and pointed acrosome and the mitochondrial helix (from Table 8.7, having 15 curves).

"& Reproductive Biology and Phylogeny of Birds The spermatozoon is said to be 77 µm long (but contrast 101.56 µm, Birkhead et al. 2006, see Table 8.7), with an acrosomal length of 6-7 µm, a nuclear length of 6 µm and a midpiece length of 55 µm. There is a small anterior nuclear fossa, 0.7-1.08 µm deep; a posterior nuclear fossa is absent. The acrosome has an electron dense core surrounded by a more electron lucent region but lacks a perforatorium. It is said that the helical membrane (here considered a keel) is limited to the acrosome. This spiral keel is illustrated by SEM (Fig. 8.52C), here considered equivalent to that of Passer italiae. However, from the length of midpiece given and the illustration of Retzius (1909) (Fig. 8.41) it is clear that the midpiece forms a helix extending far along the axoneme, though with a smaller amplitude than in most passerines. A distal but no proximal centriole is present. Accessory [dense] fibers are present in the midpiece (Koehler 1995). The sperm of this species was also examined but not individually described by Humphreys et al. (1972) and Asa and Phillips (1987).

8.10.12.11

Passer italiae

General morphology. Furieri (1961) provides useful diagrammatic sketches of the whole mature spermatozoon of Passer italiae, the Italian sparrow, accompanied by corresponding cross sections (Fig. 8.47B). The acrosome and nucleus are helical and the tail is surrounded by an helical crest (apparently a true helical membrane consisting of the spiral mitochondrion). The spiral nature of the head is more evident than in Turdus merula, the blackbird, but the ‘spiral crest’ of the tail is less evident. The spermatozoon has a length of 110 µm (cf 101.56 µm for P. domesticus, Birkhead et al 2006); the maximum width of the head is 1.5 µm; the length of the acrosome is 9 µm and that of the nucleus 7 µm, thus demonstrating the usual oscine feature of an acrosome:nucleus ratio greater than 1. Acrosome. The acrosome is an elongate gently spiraled cone consisting of homogeneous, strongly electron-dense material. The spiral crest of the acrosome is better developed than in the blackbird and arises from a larger base. It extends perpendicularly to the long axis of the acrosome and differs in constitution from the acrosome. Nucleus. The nucleus resembles that of the blackbird. Flagellum. Unlike the blackbird, the flagellum is not implanted directly in the long axis of the spermatozoon. Whereas the mitochondrial component of the helical crest extends the whole length of the crest in the blackbird, in Passer, the first tract appears to be formed by a large fiber [presumably the granular helix], as in the blackbird, but here preceding the mitochondrion in which the first cristae are evident at the end of the first gyre of the helix. The other difference is the absence in Passer of the large structureless fiber (fibrous keel).

8.10.12.12

Passer diffusus

A scanning electron micrograph of the spermatozoon of the Grey-headed sparrow (Passer diffusus) is given in Figure 8.54A. A well developed helical acrosome keel, and a mitochondrial helix, are seen.

Avian Spermatozoa: Structure and Phylogeny

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Fig. 8.52 Scanning electron micrographs of acrosomes of the sperm of four species of Passerida. Some species have a nearly linear acrosomal axis while others have a spiral shaped axis. All have an helical membrane. A. Quiscalus quiscalus. ¥ 6800. B. Agelaius phoeniceus, Red-winged blackbird. ¥ 8700. C. Passer domesticus, House sparrow. ¥ 9300. D. Cardinalis cardinalis, Cardinal. ¥ 9300. After Koehler, L. D. 1995. Mémoires du Muséum National d’Histoire Naturelle, Paris. 1995; 166: 437-444, Fig. 1.

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Reproductive Biology and Phylogeny of Birds

Fig. 8.53 Social weaver (Philetairus socius), Ploceidae. A. LS of the entire acrosome, nucleus and centriolar region. As in D and I, at least the first gyre of the helix consists of the granular helix. B. Transverse section (TS) of acrosome core through the helical keel. C. Acrosomal-nuclear junction showing anterior nuclear fossa receiving the base of the acrosome core. D. LS base of nucleus and the centriolar region. E. Longitudinal section (LS) of acrosome and anterior nucleus. F. Detail of acrosome crest and anterior acrosome core. G. LS acrosome crest and anterior acrosome core at right angles to F. H. TS nucleus. I. LS base of nucleus, and centriolar and anterior axonemal region surrounded by the mitochondrial helix which appears to commence with a granular helix. J. TS axoneme and mitochondrial helix, showing nine dense fibers associated with the axonemal doublets. K. TS axoneme behind posterior to the mitochondrial helix. L. TS posterior region of axoneme with reduced dense fibers. acr, acrosome crest; ac, acrosome core; ak, acrosome keel; anj, acrosome-nuclear junction; ax, axoneme; dc, distal (only) centriole; df, dense fiber; mh, mitochondrial helix; n, nucleus. Jamieson, Hodgson and Spottiswoode unpublished.

Avian Spermatozoa: Structure and Phylogeny

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8.10.12.13 Ploceidae, Philetairus socius Acrosome. The acrosome of Philetairus socius sperm is a helical structure 1.2 times the length of the nucleus which it surmounts (Fig. 8.53A). It is inserted into the nucleus, at the acrosomal-nuclear junction (Fig. 8.53A, C, E), in an approximately symmetrical V-shaped fossa. The acrosome is bipartite. Its distal, much longer portion, consists of an electron-dense gently helical column of three gyres, the acrosome core, 5.8 µm long, which bears a prominent keel seen in profile as three prominent spurs. Unlike Myrmecocichla formicivora, the dense core is not drawn out towards the keel which is formed solely from the crest layer which invests the core, as seen in longitudinal (Fig. 8.53A, E, G) and transverse section (Fig. 8.53B). The proximal (anterior) portion of the acrosome, the acrosome crest, as seen in longitudinal section, consists of a short electron-pale shaft, 1.3 µm long, tapering to a narrow tip (Fig. 8.53G) and bearing near the level of the core, a single spur representing the keel. However, the crest extends posteriorly as the crest layer or sleeve to the base of the acrosome and bears along its length three spurs representing the continuation of the acrosome keel (seen in cross section in Figure 8.53B). In longitudinal sections approximately at right angles to this (Fig. 8.53A, E, F), the short acrosome crest has the form of an inverted shoe of which the first spur forms the heel. The tip of the acrosome core, fits into a deep asymmetrical fossa formed by the base of the crest (Fig. 8.53A, E, F). Nucleus. The nucleus, like the acrosome core, is strongly electron-dense. It forms a very slightly sinuous stout cylinder (Fig. 8.53A, D, E, G), 6.1 µm long, of subcircular transverse section (Fig. 8.53H). Its greatest width, 0.8 µm, is basal but there is little variation throughout its length. Its base closely abuts the centriolar region (Fig. 8.53A, D). Centriolar complex. There is no proximal centriole. A dense ring surrounding the distal centriole, seen in M. formicivora, has not been found and there is no mitochondrial ring. However, dense convoluted masses are present within the lumen of this centriole and the proximal region of the axoneme (Fig. 8.53I). Helical components of the axonemal region. A longitudinal section of the base of the nucleus and adjacent centriolar complex and midpiece (Fig. 8.53I) reveals a very elongate mitochondrion wound helically around the axoneme. However, at least the first gyre of this helix is amorphous and granular and is presumed to be identifiable as a granular helix. No fibrous helix, seen in M. formicivora, is present. Glancing sections of nine dense fibers are visible between the mitochondrion and the axonemal doublets, as confirmed from transverse sections (Fig. 8.53J). Axoneme. The axoneme has the conventional 9+2 arrangement of microtubules. For much of its length each doublet is accompanied by a dense fiber which is circular in cross section except for a small prolongation which joins each A microtubule near the junction of the latter with the B subtubule, at the level of the mitochondrial helix (Fig. 8.53J) and posterior to this (Fig. 8.53K). The dense fibers greatly reduce in size distally (Fig. 8.53L). Sections of the endpiece have not been obtained (Jamieson, Hodgson and Spottiswoode unpublished).

"&" Reproductive Biology and Phylogeny of Birds

Fig. 8.54 Scanning electron micrographs of Spermatozoa of Passeridae and Ploceidae. A. Grey-headed sparrow (Passer diffusus), Passeridae. B-D. Red-billed quelea (Quelea quelea), Ploceidae. acr, acrosome crest; ac, acrosome core; ak, acrosome keel; anj, acrosome-nuclear junction; ax, axoneme; mh, mitochondrial helix; n, nucleus. Courtesy of A. Hodgson, relabeled.

Other ploceids. The spermatozoa of Red-billed quelea (Quelea qualea) (Fig. 8.54B), Red bishop (Euplectes orix) (Fig. 8.55A-C) and Cape weaver (Ploceus capensis) (Fig. 8.55D, E) are illustrated by scanning electron microscopy. All show an helical acrosome keel and a mitochondrial helix. However, in one specimen of the Red bishop (Fig. 8.55C) the midpiece displays a short, projecting mitochondrion resembling the midpiece of a corvid. The significance of the variant is not clear.

Avian Spermatozoa: Structure and Phylogeny

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Fig. 8.55 Scanning electron micrographs of Spermatozoa of Ploceidae, continued. A, B, C. Red bishop (Euplectes orix). D, E. Cape weaver (Ploceus capensis). acr, acrosome crest; ac, acrosome core; ak, acrosome keel; ax, axoneme; m, mitochondrion; mh, mitochondrial helix; n, nucleus. Courtesy of A. Hodgson, relabeled.

8.10.12.14

Paridae, Parus bicolor, P. major and Cyanistes caeruleus

Koehler (1995) cites the unpublished optical description of the sperm of Parus bicolor, the Tufted titmouse. It is 90 µm long, with an acrosome 7.1 µm long and a shorter nucleus, at 5.8 µm. The sperm is helical but the helical membrane (keel) is said to be limited to the acrosome. There is no anterior

"&$ Reproductive Biology and Phylogeny of Birds nuclear fossa but a slight posterior fossa is present. The midpiece is 50 µm long. Dense fibers, of unspecified location, are present. Birkhead et al. (2006) (Table 8.7) give dimensions and other morphological data for the Great tit (Parus major) and Willow tit (Parus montanus). Comparable measurements for the Blue tit (Cyanistes caeruleus) are straightened helix length 81.80 µm, length flagellum 92.86 µm, total sperm length 109.61 µm, number of midpiece gyres 19.60, number of head gyres 4.00 (T. R. Birkhead, pers. comm.)

8.10.12.15 Icteridae, Icterus galbula, Agelaius phoeniceus, Molothrus ater, and Quiscalus quiscula Spermatozoal ultrastructure in the Icteridae has been briefly treated for Icterus galbula, Northern oriole (Asa and Phillips 1987); Agelaius phoeniceus, Redwing (Asa and Phillips 1987; Koehler 1995) (acrosome, Fig. 8.52B); Molothrus ater, Brown-headed cowbird (Koehler 1995); and Quiscalus quiscula, Grackle (Koehler 1995) (acrosome, Fig. 8.52A). Data for the last three species are given in Table 8.8. Table 8.8

Characters of spermatozoa of the Icteridae (from Koehler 1995)

Species

Molothrus ater

Agelaius phoeniceus

Quiscalus quiscala

Common name Sperm length Acrosome length Nuclear length Ratio acrosome:nucleus Granular body at midpiece Midpiece length Helical shape Helical membrane Anterior nuclear fossa Posterior nuclear fossa Centrioles Accessory dense fibers Mitochondria

Brown-headed cowbird 62 µm 6.0 µm 3.0 µm 2 yes 24 µm yes yes shallow yes

Red-winged blackbird 110 µm 8.0 µm 5.0 µm 1.6

Common grackle 48 µm 7.0 µm 4.0 µm 1.75 yes

yes spiral

yes yes shallow yes distal only yes

These data for icterids, though incomplete, are characteristic of passeridan sperm: sperm helical, acrosome longer than the nucleus (ratio 1.6-2), midpiece spiral and very extensive (in Molothrus ater almost half the length of the axoneme); distal centriole, only, present. The presence of dense peripheral axonemal fibers is shared with non-passerines, though they are larger (size not demonstrated for icterids) in passerines. The passeridan nature of the sperm is confirmed by Asa and Phillips (1987) for Northern oriole and Redwing, in their generalized account of oscine sperm. Allen et al. (1967) gave detailed measurements for the sperm if 12 icterid species or subspecies (Table 8.9). Their data for the species described above by Koehler (1995) differ significantly from those of the latter author. Statistical

14.95 ± .0704 15.73 ± .0491 14.20 ± .688 14.18 ± .0701 15.51 ± .0724 14.75 ± .0543 14.54 ± .0683 16.11 ± .0407 15.97 ± .0982 16.12 ± .0970 15.67 ± .0850 13.74 ± .0763

Sturnella neglecta confluenta Western meadowlark Sturnella magna argutula Southern meadowlark Quiscalus quiscula versicolor Bronzed grackle Dolichonyx oryzivorus Bobolink Icterus bullockii bullockii Bulock’s oriole Icterus spurius Orchard oriole Icterus parisorum Scott’s oriole Agelaius phoeniceus phoeniceus Eastern red-winged blackbird Agelaius phoeniceus mearnsi Florida red-winged blackbird Agelaius phoeniceus nevadensis Grinnell red-winged blackbird Agelaius phoeniceus floridanus Maynard’s red-winged blackbird Cassidix mexicanus major Eastern boat-tailed grackle

Sturnella neglecta confluenta Western meadowlark Sturnella magna argutula Southern meadowlark Quiscalus quiscula versicolor Bronzed grackle Dolichonyx oryzivorus Bobolink Icterus bullockii bullockii Bullock’s oriole Icterus spurius Orchard oriole Icterus parisorum Scott’s oriole Agelaius phoeniceus phoeniceus Eastern red-winged blackbird Agelaius phoeniceus mearnsi Florida red-winged blackbird Agelaius phoeniceus nevadensis Grinnell red-winged blackbird Agelaius phoeniceus floridanus Maynard’s red-winged blackbird Cassidix mexicanus major Eastern boat-tailed grackle

1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

77.94 ± .1939 85.97 ± .2461 90.73 ± .3010 142.02 ± .3832 110.45 ± .2974 105.92 ± .2295 105.93 ± .2337 148.59 ± .4166 146.48 ± .3623 139.64 ± .3327 146.95 ± .3929 109.12 ± .3340

Total length

Sperm head — X t = Std. error

Species and subspecies .0641 .0837 .0896 .0872 .1331 .0408 .0456 .0763 .0761 .0723 .0682 .0566 42.84 ± .2287 49.11 ± .1035 51.96 ± .2873 111.49 ± .2133 70.47 ± .3224 75.07 ± .2030 68.95 ± .1644 106.84 ± .2623 107.85 ± .2326 104.12 ± .2304 111.13 ± .2607 71.45 ± .2320

Principal piece

9.90 ± 10.39 ± 9.55 ± 9.23 ± 10.62 ± 10.48 ± 9.35 ± 10.75 ± 10.48 ± 11.09 ± 10.17 ± 8.80 ±

Acrosome — Xt = Std. error ± ± ± ± ± ± ± ± ± ± ± ±

0.332 .0232 .0194 .0289 .0291 .0291 .0311 .0272 .0253 .0349 .0287 .0270 17.46 ± 18.57 ± 25.02 ± 17.59 ± 23.59 ± 17.75 ± 20.21 ± 24.02 ± 23.24 ± 19.52 ± 18.51 ± 23.92 ±

.1363 .0946 .1569 .1258 .1632 .1289 .0862 .1183 .1233 .1753 .1449 .1184

End piece

1.93 2.00 2.13 2.24 2.24 1.99 2.31 2.13 2.30 2.09 2.35 1.88

Midpiece — X t = Std. error

Table 8.9 Dimensions of spermatozoa of 12 taxa of Icteridae. Relabeled after Allen et al. (1967). Proceedings of the Indiana Academy of Science 77: 434-441,Table 1

Avian Spermatozoa: Structure and Phylogeny

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"&& Reproductive Biology and Phylogeny of Birds analyses showed that the measurements ‘provide criteria for distinguishing between most the twelve blackbird forms studied’. In a line drawing of a typical icterid sperm an helical ‘undulating membrane’ is shown extending throughout the length of the acrosome, nucleus and principal piece. The differentiation of this membrane, in the present chapter, into acrosome keel, nuclear keel and mitochondrial helix was not recognized by light microscopy.

8.10.12.16

Emberizidae, Cardinalis cardinalis and Emberiza spp.

The Cardinal (Cardinalis cardinalis) was included in the oscine study of Henley et al. (1978) but was not specifically referred to. Koehler (1995) has given an SEM of the acrosome (Fig. 8.52D) and the following data: length sperm 76 µm, length acrosome 6.6 µm, length nucleus 4.0 µm, length midpiece 35 µm, sperm helical, helical membrane present. The large acrosome:nucleus ratio of 1.65 and the long midpiece are typical passeridan features. Tripepi and Perrotta (1991) have charted the development of the granular body (GB) in the Cirl bunting (Emberiza cirlus) (Fig. 8.56A-D; see also section 8.10.10.1). The GB is evidenced in the early stages of spermatogenesis (Fig. 8.56A) as the spermatid begins to elongate and the midpiece is forming. Subsequently, when the microtubular helix appears and the spermatid assumes an helical form, the GB constitutes the proximal part of what is here interpreted as an helical band around the axoneme, the granular helix (GH). In the advanced spermatid (Fig. 8.56B, C) the helix is formed internally by the GH, which coils around the axoneme and its dense fibers, and externally by the transient microtubular bundle (Fig. 8.56B, C, D). In the remainder of the midpiece, the GH is substituted by the mitochondrial sheath [mitochondrial helix] (Fig. 8.56B). Tripepi et al. (1991) have commented on, and illustrated, the microtubules of the spermatid of Emberiza cirlus, the Cirl bunting. The arrangement is considered to be at a fourth, and final, level above the “reptilian” arrangement. As in other passeriforms, Sertoli cell microtubules are absent but cytoplasmic microtubules reappear in the spermatid. They are arranged in an helicoidal bundle which is eliminated during the last phase of spermiogenesis. Fig. 8.56 A-D. Cirl bunting (Emberiza cirlus). A. Spermatid at commencement of elongation. The granular body has formed near the base of the uncondensed nucleus. A large acrosome vesicle surmounts the nucleus which is still subspheroidal. B. Elongating spermatid with helical components. The nucleus is condensed and the granular body has become a granular helix around the proximal region of the axoneme and is overlain by the microtubular helix. Further distally, the granular helix gives way to the developing mitochondrial helix. C. Detail of same. D. Two transverse sections of the proximal region of the axoneme, showing the granular helix overlain by the microtubular helix. Dense fibers surround the axoneme. E. Tree creeper (Certhia brachydactyla). Transverse section of the proximal region of the axoneme and dense fibers through the granular helix, after regression of the microtubular helix. A and E. Relabeled after Tripepi, S. and Perrotta, E. (1991). Pp. 1021-1023. In Baccetti B. (ed.), Comparative Spermatology 20 Years After. Serono Symposia Publications, vol. 75. Raven Press, Rome, Figs. 1 and 4. B–D. From micrographs courtesy of Sandro Tripepi.

Avian Spermatozoa: Structure and Phylogeny

Fig. 8.56

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"' Reproductive Biology and Phylogeny of Birds Birkhead et al. (2006) have tabulated data for the Corn bunting (Emberiza calandra) and the Yellow Hammer (E. citrinella) (Table 8.7).

8.10.12.17

Fringillidae

Fringillids are briefly referred to in 8.10.12.1 above where they are shown to have typical passeridan sperm. However, Birkhead et al. (2006) have demonstrated a remarkable and profound departure from passeridan sperm structure in the Eurasian bullfinch (Pyrrhula pyrrhula). The Eurasian bullfinch sperm are relatively short: 46.87 µm ± 3.83 SD (N = 11) (captive-bred males: 47.19 µm ± 4.28 SD (N = 8) and wild males 46.01µm ± 2.75 SD (N = 3)). The sperm have a rounded acrosome and head, and no obvious mitochondrial helix. The sperm of Beavan’s Bullfinch, like those of the Eurasian bullfinch are short (length 49.11µm) but instead of a rounded head they have a more pointed, spiral-shaped head. Both Pyrrhula species have an extremely small midpiece, in contrast to the elongated mitochondrial helix that forms the midpiece in almost all other passerines. A pair-wise study using principle components analysis (PCA) to combine quantitative and qualitative sperm morphology traits, together with a phylogenetic correlation, comparing the sperm morphology of the Eurasian bullfinch and Beavan’s bullfinch Pyrrhula erythaca with nine other pairs of congeneric passerines revealed that the Eurasian bullfinch was a dramatic outlier in terms of its sperm morphology and also more different from the closely related Beavan’s Bullfinch than were any other pair of species. Excluding the Eurasian bullfinch from the analysis showed that most variation in sperm morphology in the other species was attributable to phylogeny (a correlation long espoused by the present author). The Eurasian bullfinch has extremely small testes for its body size, indicating that sperm competition is infrequent in this species; the possibility was discussed that relaxed selection via a lack of sperm competition may have contributed to unusual sperm morphology in this species. The sperm structure in this species resembles that of a spermatid in the early stages of elongation and suggests a suppression of late spermiogenesis resulting in what could be termed spermatozoal neoteny. Fig. 8.57 Lonchura striata domestica Lovebird (Bengalese finch). Late maturation phase of spermatids. A. Longitudinal section (LS), through the head showing the posterior one gyre of the acrosome (A) with the same pitch as the nucleus (N). A spiral keel of the acrosomal ridge is indicated by arrows. MTB, microtubule bundle. B. Oblique section through the posterior region of the acrosome (A), showing the Sertoli cell process (arrow) protruding between the acrosome and the microtubule bundle (MTB). S. Sertoli cell. SER. Smooth endoplasmic reticulum. C. Transverse section (TS) through the microtubule bundle (MTB) on the ridge of the nucleus (N), showing the alternating layers consisting of microtubules and flattened cisternae of endoplasmic reticulum (sER). D. Ts through anterior half of the midpiece, showing the layered structure of the microtubule bundle. MS. Mitochondrial sheath. Fig. 8.57 Contd. ...

Avian Spermatozoa: Structure and Phylogeny

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Fig. 8.57 Contd. ...

SER. Smooth endoplasmic reticulum. E. LS through the posterior half of the midpiece, showing few cisternae of smooth endoplasmic reticulum in the microtubule bundle (MTB). AN. Annulus. MS. Mitochondrial sheath. F. LS through a mature spermatid, showing disappearance of the microtubule bundle. A. Acrosome. MS. Mitochondrial sheath. N. Nucleus. Scales: A. ¥ 14700. B. ¥ 31000. C. ¥ 108400. D. ¥ 62000. E. ¥ 11600. F. ¥ 9000. After Kondo, T., Hasegawa, K. and Uchida, T. 1989. Journal of Ultrastructure and Molecular Structure Research 98: 158-168, Figs. 18-23. With permission from Elsevier.

"'

Reproductive Biology and Phylogeny of Birds

An exception such as the Eurasian bullfinch is not here considered to substantially militate against the value of sperm ultrastructure for taxonomy. One would not argue that we cannot morphologically recognize and classify the Crustacea because Sacculina resembles a fungal mycelium. The statement (Birkhead et al. 2006) that ‘most variation in sperm morphology in the other species was attributable to phylogeny’ supports use of spermatozoal morphology in phylogeny and therefore classification. Sperm morphology is only one tool but a useful complement to other means of determining relationship in addition to its interest per se. In fact the passerine status of the Eurasian bullfinch sperm has since been confirmed by recognition (Birkhead, Giusti, Immler and Jamieson, in preparation) in it of a microtubular helix, a hallmark of passerine sperm, differing from the manchette of any other animal spermatid. Passerines examined by Birkhead et al. (2006) by light and scanning electron microscopy for pairwise analysis of spermatozoal dimensions are listed in Table 8.7. These authors also give the following data on the sperm of

Fig. 8.58 Lonchura striata, Lovebird (Bengalese finch). Scanning electron micrographs of spermatozoon. A. Sperm head and midpiece of the flagellum surrounded by the loosely coiled helical membrane. ¥ 4400, B. Detail of the head. ¥ 34500. After Yasuzumi, G. 1974. International Review of Cytology 37: 53-119, Figs. 10-11. With permission from Elsevier.

Avian Spermatozoa: Structure and Phylogeny

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the Pine Grosbeak (Pinicola enucleator). The sperm are similar to those of other passerines in length (162.7 µm ± 9.63 S.D.; N = 30 sperm, N = 1 male) and resemble typical finch sperm in morphology unlike Pyrrhula.

8.10.12.18

Estrildidae, Lonchura striata

The spermatozoon of Lonchura striata, the ‘Loverbird’, also named the Bengalese finch, Society finch, White-rumped mannikin, or White-rumped munia, is of the classical passeridan type, with helical acrosome, nucleus and midpiece, the midpiece extending far along the axoneme (Yasuzumi and Sugioka 1966; Fawcett et al. 1971; Yasuzumi and Sugioka 1971; Yasuzumi 1974; Kondo et al. 1988) (Figs. 8.57, 8.58). From TEM, the helical membrane (Fawcett et al. 1971; Kondo et al. 1988) (Fig. 8.57C, D) is (axoneme included) of the tripartite type and lacks the additional fibrous keel seen in Turdus. However, the helical membrane on the acrosome is clearly a keel distinct and separate from the helical membrane of the tail, both being absent from the nucleus as here shown for Philetairus socius.

8.10.12.19

Estrildidae, Taeniopygia (=Poephila) guttata

Vernon and Woolley (1999) give some data on the spermatozoon of the Zebra finch (Taeniopygia gutatta). By light microscopy and TEM the sperm are said to be very like those of the starling. The mean head length is 11.3 ± 1.0 µm and flagellar length 64.1 ± 5.7 µm (n=16 from 1 individual). The mean pitch of the mitochondrial helix is 3.13 µm. There is a highly significant negative correlation (r = – 0.85) between the lengths of the mitochondrial and nonmitochondrial regions of the flagellum. The mitochondrial helix is sinistral, as judged from serial sections (Vernon and Woolley 1999).

8.10.12.20

Prunellidae, Prunella collaris

Chiba and Nakamura (2001) give a TEM view of part of the head of the spermatozoon of Prunella collaris, the Alpine accentor, on the wall of a sperm storage tubule (SST) and SEM of the spermatozoon on the utero-vaginal part of the oviduct and a group of sperm on the wall of a SST. The utero-vaginal micrograph well displays the auger-like form of the passerine spermatozoon.

8.11

PHYLOGENETIC SUMMARY OF AVIAN SPERMATOZOA

Introduction. A subset of the spermatozoal characters discussed in this chapter (see Table 8.10) is here used for a preliminary phylogenetic analysis of taxa which have been examined ultrastructurally. Many spermatozoal descriptions are fragmentary and, because of the resulting missing characters, do not support a full parsimony analysis using exhaustive or branch and bound analyses. Resultant consensus trees are highly polytomous. A single nearest neighbor (NJ) tree is therefore presented (Fig. 8.59A). It was obtained using PAUP (Swofford 2001). Although neighbor-joining is essentially phenetic, unambiguous character state changes were plotted on the tree using MacClade (Maddison and Maddison 2000). Some clearly synapomorphic

"'" Reproductive Biology and Phylogeny of Birds characters which were not computed as unambiguous, probably because of missing states in various taxa, are added to the tree in parentheses. The analyses were conducted on 29 avian species (only Corvus and Passer being composite taxa), plus the outgroups Crocodylus (Jamieson et al. 1997) and a composite Chelonia (Healy and Jamieson 1997). The character matrix is available from the author on request. It has been claimed (Johnson 2001, citing Huelsenbeck and Hillis 1993) that the performance of neighbor joining (NJ) in recovering the correct phylogeny is similar to that of parsimony. A portion of a 50% majority rule, maximum parsimony (MR) tree is, nevertheless, shown in Fig. 8.59B. The tree obtained and character transformations indicated can only be taken as heuristic in the broad sense, the chief aim of the analyses being to graphically present the spermatozoal character states of the various taxa. Nevertheless, monophyly is supported for Neognathae, Psittaciformes, Columbiformes, Passeriformes, Passeri (oscines) and Passerida. Spermatozoal evidence rejects basal passerines. A major recent controversy in avian molecular phylogenetics has centered on the conflicting hypotheses that the Passeriformes constitute a basal group, the sister-group of all other birds or the traditional hypothesis that paleognaths (ratites and tinamous) are the sister group of all other birds. The basal passerine hypothesis has derived from analysis of mitochondrial DNA sequences (Mindrell et al. 1997, 1999; Johnson 2001). However, Braun and Kimball (2002) have shown that mtDNA sequences are capable of supporting the traditional classification, with basal paleognaths, depending on the type of analysis, number of sites analyzed and taxon sampling. Phylogenetic analysis of nuclear DNA sequences has consistently supported the traditional hypothesis (Sibley and Ahlquist 1981, 1990; Sibley et al. 1988) Table 8.10

Characters used in phylogenetic analysis of evolution of bird spermatozoa

Characters (All unordered) 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

Acrosome1 shorter 2 equal to 3 longer than nucleus Acrosome 1 conical 2 round tip 3 button-like Perforatorium 1 present 2 absent Endonuclear canal 1 deep 2 short 3 absent Proximal centriole 1 present 2 absent Distal centriole 1 very long 2 shorter 3 short Mitochondria 1 several per TS 2 one per TS 3 small group Midpiece 1 short 2 very long ‘Helical membrane’ 1 present 2 absent Singlets in distal centriole 1 present 2 basal 3 absent Fibrous sheath 1 ribbed 2 amorphous 3 absent Annulus 1 present 2 absent Dense fibers 1 small 2 moderate 3 large Tip granule 1 present 2 absent Nuclear rostrum penetrating acrosome 1 deeply 2 slightly 3 negligibly 4 absent

Avian Spermatozoa: Structure and Phylogeny

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Fig 8.59 A. Neighbor-joining tree of major groups of birds from a PAUP analysis derived from spermatozoal ultrastructure for characters listed in Table 8.10, with Chelonia and Crocodylus johnstoni as the outgroup. Neighbor-joining search settings: ties (if encountered) broken systematically; distance measure = mean character difference. Character changes are those which are unambiguous and were plotted using Macclade. Some deduced synapomorphies which were not plotted as ambiguous have been added in parentheses to the tree. B. Portion of a 50% majority rule most parsimonious tree. Settings used in computation of the tree: heuristic search settings; optimality criterion = parsimony; 15 total characters, all unordered, with equal weight, all parsimony-informative; starting tree(s) obtained via stepwise addition; addition sequence: random; 10 replicates; 1 tree held at each step during stepwise addition; branch-swapping algorithm treebisection-reconnection (TBR); steepest descent option not in effect; Initial ‘MaxTrees’ setting 900 (auto-increased by 100); other parameters at PAUP default settings.

"'$ Reproductive Biology and Phylogeny of Birds and recently Garcia Moreno et al. (2003) have shown that combined mt and nuclear DNA analysis supports the traditional view. A study of a-crystalline sequences also supported the basal position of ratites (Stapel et al. 1984). The NJ tree (Fig. 8.59A), as do intuitive considerations, unequivocally supports a basal position for paleognaths when Chelonia and Crocodylia are used as outgroups. A basal position for passerines is highly unparsimonious and, computationally and intuitively, would involve most implausible pathways for spermatozoal evolution. Are paleognaths paraphyletic? Monophyly of the Palaeognathae has previously been accepted on the basis of spermatozoal studies, as in the molecular ‘Tapestry’ (Sibley et al. 1988; Sibley and Ahlquist 1990), though the position of Tinamou within these has remained unresolved (Asa et al. 1986; Asa and Phillips 1987; Baccetti et al. 1991; Soley 1993). It must be stressed that the spermatozoal characters which appear to unite Palaeognathae are in fact symplesiomorphies: the conical acrosome, ribbed fibrous sheath, elongate centriole, and 9 dense fibers of ratite sperm are all seen in Chelonia and Crocodylia, and do not prove avian or paleognath monophyly, despite the fact that the first three features were considered to unify paleognaths (Baccetti et al. 1991). In the NJ tree monophyly of paleognaths is not supported (Fig. 8.59A). It is not here proposed that paraphyly of paleognaths indicated by spermiocladistics should be considered unquestionable but it does warrant further consideration of paraphyly for the group. The struthioniform spermatozoon is crocodiloid (see section 8.6.1.3), departing from the condition in crocodile sperm in an apparent reduction in size of the outer dense fibers of the axoneme and the deeper, more anterior extension of the nuclear rostrum into the subacrosomal space of the acrosome, neither of which characters computed as unambiguous in the NJ tree. Like the crocodilian (and chelonian) spermatozoon the elongate distal centriole (and the midpiece) is penetrated throughout its length by the central singlets of the axoneme. By deduction, withdrawal of the central singlets from the distal centriole appears to be an apomorphy, but homoplasy, of the Crested tinamou (Eudromia elegans), the Emu (Dromaius novaehollandiae) and the remainder of the birds. However, in the NJ tree withdrawal of singlets from the centriole computes as basic to Neornithes with a, perhaps unlikely, repenetration in the Struthio+Rhea clade. Tinamou has a unique autapomorphy (not computed), the presence of glycogen around the axoneme, but this is phylogenetically uninformative. Emu is distinctive among the paleognaths in loss of the endonuclear canal and of the perforatorium (two probably correlated characters). If the Palaeognathae were truly paraphyletic, the strict application of Hennigian phylogenetics would demand that Rhea+Struthio, Eudromia and Dromaius were each given separate and successively reduced ranks as collectively they would constitute a grade rather than a clade. The author is doubtful, however, that any useful purpose is served by thus distinguishing taxa which have arisen in immediate succession from a basal lineage, here of

Avian Spermatozoa: Structure and Phylogeny

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the Aves, and bear a distinctive morphological and molecular facies. Nevertheless, intervention of Tinamou between the Ostrich and Emu branches (Fig. 8.59A), if borne out by further studies, would be of considerable interest. A possible sister relationship of Emu to the Neognathae is highly questionable as it would require reacquisition of the perforatorium and endonuclear canal in the Galloanserae rather than the more probable retention of these from an ancestor shared with paleognaths. The summary tree for molecular data of Harshman (Chapter 1, Fig. 1.1) indicates monophyly of the Palaeognathae. Galloanserae. Any member of the Galloanserae, consisting of the Galliformes and Anseriformes, could instantly be diagnosed by a combination of features of sperm ultrastructure: acrosome shorter than nucleus: stout perforatorium in a shortened endonuclear canal; both centrioles present; distal centriole long though (unlike paleognaths) not fully penetrating the midpiece; mitochondria in a circlet around the axoneme; midpiece not elongate; ‘helical membrane’ absent; singlets basal in the distal centriole; fibrous sheath amorphous, not ribbed as in paleognaths; annulus present; dense fibers of moderate diameter; nuclear rostrum slightly penetrating the acrosome. However, among these character states there is no distinctive synapomorphy for the group and the NJ tree presents an unresolved assemblage of seven separate galloanseran branches, together with three non-galloanseran clades. Nevertheless, spermatozoal ultrastructure certainly does not contraindicate monophyly of the Galloanserae which is supported by molecular analyses (Harshman, Chapter 1, Fig. 1.1). The Anseriformes and Galliformes can be linked by the synapomorphy slight penetration of the nuclear rostrum into the subacrosomal space and the Galliformes can possibly be separated from the Anseriformes by presence of singlets basally in the distal centriole, both of which are ambiguous characters and do not therefore appear on the NJ tree. The latter (singlet) character is uncertain for the Anseriformes and requires reexamination. It is likely that when more complete spermatozoal data are obtained considerable resolution of the Galloanserae will be obtained. For instance, the presence of a ‘tip granule’ unites Coturnix japonica and Gallus gallus. It is uncertain that its absence from other taxa is real but the thorough examination of Turkey and Guineafowl (Thurston et al. 1982; Thurston and Hess 1987) suggests that it is absent at least in these two species. A notable difference of the spermatozoal analyses from the Tapestry (Sibley et al. 1988) is that the Galloanserae do not form the sister taxon of the ‘Ratitae’. Sibley and Ahlquist (1990), did, however, repudiate this sister taxon relationship in their classification. A sister relationship of Galloanserae and Neoaves is endorsed by Harshman (Chapter 1, Fig. 1.1). Metaves. Three orders of the Metaves have been investigated for spermatozoal ultrastructure: Apodiformes, Caprimulgiformes and Columbiformes. There is no clear spermatozoal support for the grouping Metaves in the computer analyses (Fig. 8.59A).

"'& Reproductive Biology and Phylogeny of Birds Apodiformes. The midpiece-length distal centriole of Apus apus (Fig. 8.22) unless (as seemed most unlikely) a reversal from the ancestral condition retained in crocodiles and paleognaths, could be considered (Jamieson and Tripepi 2006) to place the apodids very basally in the avian phylogenetic tree but the loss of the perforatorium is derived relative to the Galloanserae. The long centriole of A. apus, if not a reversal, is inconsistent with placement of the Apodiformes above (i.e. at a more derived node than) the Psittaciformes from DNA-DNA hybridization by Sibley and Ahlquist (1990). It is not inconsistent, however, with placement in the Metaves (see Harshman, Chapter 1). The centriolar condition might suggest that apodiforms are the plesiomorphic sister taxon of all other Metaves and that they have independently lost the perforatorium. However, the computer analysis (Fig. 8.59A) does not support this position and Apus has a relatively ‘high’ position, lying in a clade which includes Piciformes, Cuculiformes, Charadriiformes, and Columbiformes + Passeriformes at a node above Psittaciformes. It seems necessary to consider a scenario in which a long (midpiece-length) centriole existed at three major nodes: the Palaeognathae-Neognathae node, the Galloanseran-Neoaves node, and though not evident in the present analyses, a Metaves-Coronaves node. The centriole would then become shorter, though still elongate, in the ancestral galloanseran and would be retained in the Neoaves only (so far as is known) in Apus, whether in the Metaves or the Coronaves. Caprimulgiformes. The phylogenetic position of Caprimulgus, as evidenced by spermatozoal ultrastructure (Fig. 8.23), has been partly discussed in section 8.9.2.1 above. The caprimulgid spermatid (and clearly the spermatozoon) is closely similar in structure to that of psittaciforms. In contrast, the Apodidae, placed in the same clade as the Caprimulgidae in the DNA-DNA hybridization analyses of Sibley and Ahlquist (1990), differ from Caprimulgus in the long distal centriole of the spermatozoon (Jamieson and Tripepi 2006). Other members of the unnamed clade of Sibley and Ahlquist (1990) which contains the caprimulgids (Hemiprocnidae through Eurostopodidae) have not been examined for sperm ultrastructure. Although the Trochilidae and (here removed to the Coronaves) the Strigidae in this clade, are listed as having been examined by light microscopy (McFarlane 1963), no data are given. The NJ tree (Fig. 8.59A) places Caprimulgus in the Neognathae but gives no indication of their affinities other than that they do not belong in a columbiform+passerine clade. In the MR tree (Fig. 8.59B), although Caprimulgus lies in the same clade as Apus, this clade also contains members of the Coronaves and therefore the grouping Metaves cannot be considered to be supported. Columbiformes. In the molecular Tapestry (Sibley et al. 1988; Sibley and Ahlquist 1990) the Passeriformes branch off first in the tree dealing with the Passerimorphae and form the sister-group of the Columbiformes + (Gruiformes + ‘Ciconiiformes’), although the classification does not call

Avian Spermatozoa: Structure and Phylogeny

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attention to this order of branching as the Columbiformes is the first order listed in it. In the NJ tree (Fig. 8.59A) columbiforms are the sister of the passerine clade. Association of columbiforms with passerines is marked by the loss of the fibrous or amorphous sheath (a change which we could alternatively envisage as homoplastic). The mutual presence of an elongate midpiece maps as a homoplasy. Homoplastic elongation of the midpiece is acceptable as detailed ultrastructure of the midpiece (not computed) in columbiforms differs from that of passerines. If the Metaves were a valid grouping loss of the amorphous sheath in Columbiformes would be homoplastic with taxa in the Coronaves. Nevertheless, the location of columbiforms by Sibley and Ahlquist (1990) in a clade which would now be attributable to the Coronaves is not contraindicated by spermatozoal ultrastructure. The phylogenetic position of the Columbiformes and the validity of the Metaves are far from settled. The molecular consensus of Harshman (Chapter 1, Fig. 1.1) places Columbiformes in the Metaves far removed from the Passeriformes. Coronaves. Orders of the Coronaves that have been investigated for spermatozoal ultrastructure (see accounts in previous sections) are: Piciformes, Cuculiformes, Psittaciformes, Gruiformes, Charadriiformes, Falconiformes and Passeriformes. As usual in avian studies, the passerines have received the most attention in terms of numbers of taxa investigated. The grouping Coronaves is not supported by sperm ultrastructure. Psittaciformes. In the molecular analyses of Sibley et al. (1988) and Sibley and Ahlquist (1990), the psittaciforms are preceded hierarchically only by the Coliiformes and Cuculiformes at the base of the Passerae. Psittaciforms have a highly unresolved position in the Coronaves in the consensus phylogenies of Harshman (Chapter 1, Fig. 1.2). In the NJ tree psittaciforms lie in the same unresolved assemblage (containing both Metaves and Coronaves) as Caprimulgus. They share with columbiforms a homoplastic reduction in the size of the dense periaxonemal fibers. Psittaciforms show a homoplastic loss of the amorphous sheath, a loss shared with cuculiforms and passeriforms though this character did not map unambiguously on the NJ tree. The distinctive feature, and autapomorphy, of psittaciform sperm compared all other Aves is the condition of the nuclear-acrosomal junction (see section 8.10.5) though, as an autapomorphy, this does not clarify their relationship with other orders. The unique acrosome-nuclear disjunction was not scored in the matrix but has been added to the tree, thus uniting Melopsittacus and Nymphicus. In the portion of a 50% majority rule (MR), maximum parsimony tree shown in Fig. 8.59B Psittaciformes have a sister group relationship with Grus+Falco and successively more basal sister groups are Caprimulgus and Apus. Piciformes. The Piciformes were placed by Sibley et al. (1988) and Sibley and Ahlquist (1990) at the base of the Neoaves (ignoring the questionably placed

# Reproductive Biology and Phylogeny of Birds Turniciformes), i.e. as the sister group of all other Neoaves, a view supported, again from DNA-DNA hybridization, by Bleiweiss et al. (1994). In the NJ tree (Fig. 8.59A) Melanerpes has an unresolved position as part of a polytomy consisting also of a charadriiform (Jacana), a cuculiform (Crotophaga), Columbiformes+Passeriformes. This clade is united by loss of the perforatorium, and endonuclear canal, a feature which could, alternatively, have occurred homoplastically. Features of piciforms differing from oscines and shared with other nonpasserines (asterisks indicate characters that were not computed) are the small acrosome: nucleus ratio, the acrosome being shorter than the elongate nucleus whereas it longer in at least the higher passerines; absence of an helical membrane; distribution of mitochondria in a circlet around the axoneme of the principal piece, as seen in transverse section; the presence (so far as can be deduced from a micrograph, Fig. 8.26) of an amorphous sheath around the principal piece; the stated absence of dense axonemal fibers; and the irregular distribution* of spermatozoa in the testes (a difference from Passerida but possibly not from corvids). On the other hand, there is no mention of presence of a perforatorium seen in Galloanserae and most other non-passerines, nor of the elongate distal centriole seen in paleognaths, Apus and, with some shortening, in Galloanserae. Absence of outer dense axonemal fibers requires confirmation. This uncertainty as to the position of the Piciformes is reflected in the molecular consensus (Harshman, Chapter 1, Fig. 1.2) in which a clade containing the Piciformes has an unresolved position in the Coronaves. Cuculiformes. The sperm of Crotophaga ani as described by Saita et al. (1982b) and in this study, has the following characteristics: acrosome conical, shorter than the nucleus; perforatorium and endonuclear canal absent; short distal centriole, short midpiece with several mitochondria in transverse section, helical membrane absent, central axonemal singlets absent from the centriole, amorphous sheath present; and presence of an annulus. It is thus a distinctly non-passerine sperm. In the molecular consensus of Harshman (Chapter 1, Fig. 1.2) cuculiforms are part of a huge polytomy in the Coronaves. In the NJ tree (Fig. 8.59A) Crotophaga is part of a large polytomy, other members of which are Melanerpes, Jacana, the columbiform+passeriform clade and Apus. Determination of the phylogenetic position of the Cuculiformes thus still requires morphological and molecular clarification. Passerimorphae. The Passerimorphae (sensu Sibley et al. 1988; Sibley and Ahlquist 1990) contains the orders Columbiformes, Gruiformes, the greatly expanded order ‘Ciconiiformes’ and the Passeriformes. Passerimorphae cannot be said to be supported in the spermatozoal analyses. There is, similarly, no equivalent of the Passerimorphae in the consensus phylogenies of Harshman (Chapter 1) as the constituent families (or orders) lie in the Metaves (Columbiformes) and Coronaves (Gruiformes, orders of the former Ciconiiformes and the Passeriformes).

Avian Spermatozoa: Structure and Phylogeny

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Charadriiformes, Gruiformes and Falconiformes. Charadriiformes, subsumed in the Ciconiiformes by Sibley et al. (1988) and Sibley and Ahlquist (1990), are represented for sperm ultrastructure only by Jacana jacana. As shown in section 8.10.7.2, a button-like acrosome characterizes this species and many other charadriiform species examined by light microscopy whereas the Scolopacidae have elongate, spiral sperm. The button-like acrosome fails to group Jacana with Grus and Falco, all of which are ciconiiforms sensu Sibley and Ahlquist, in the NJ and MR trees (Fig. 8.59A, B). Grus and Falco have an unlikely sister-group relationship on the basis of the shared button-like acrosome and midpiece consisting of a small group of mitochondria. It is uncertain how much credence can be placed on this grouping as the possibility exists that this acrosomal form and midpiece morphology are homoplastic. This must indeed be the case in the passeridan Eurasian bullfinch in the sperm of which Birkhead et al. (2006) have demonstrated a button-like acrosome (and, we may add, a falconiform-like external sperm morphology), the only known exception for passerines. Furthermore, Jacana has only an unresolved relationship with Grus + Falco in the MR tree (not illustrated). With regard to the spiral sperm of scolopacids it is pertinent that molecular analyses place them firmly in a charadriiform clade (Paton et al. 2003). Falco has an unresolved position, though also in the Coronaves in the molecular consensus (Harshman, Chapter 1, Fig. 1.2). Suboscines. The sperm of suboscines (Tyranni), represented in the trees only by Tyrannus tyrannus, are poorly known. Nevertheless, they occupy their traditional place at the base of the Passeriformes in the NJ tree (Fig. 8.59A) in a passeriform clade. Monophyly of passeriforms and the basal position in these of suboscines is endorsed by molecular studies (Sibley et al. 1988; Sibley and Ahlquist 1990; Harshman, Chapter 1, Fig. 1.4). In the NJ tree (Fig. 8.59A) the only unambiguous synapomorphy for the Passeriformes is the presence of the ‘helical membrane’ (but see below). Computation of unambiguous changes (not shown) on the MR tree gave none for Tyrannus but grouped all oscines by enlargement of the periaxonemal dense fibers. It is uncertain whether in suboscines the mitochondrial component is unilateral, as it is in Passerida, and there appears to be more than one mitochondrion; the dense outer axonemal fibers have either not been illustrated or, in Tyrannus, are smaller than those of oscines; absence of a nuclear rostrum requires confirmation; and accounts are contradictory as to whether there is an helical membrane, though where described for Tyranni this differs in structure from that of oscines. As noted, an apomorphy (not computed) shared with oscines is the helical nucleus. A plesiomorphic feature not seen in oscines is the (constant?) presence of two centrioles. Passeri (oscines). The Corvida are represented by Corvus splendens but with some reference to the light microscopy of Ballowitz (1888) and Retzius (1911, 1912) and the account of Birkhead et al. (2006). In the NJ tree they were not resolved by any unambiguous character state from Tyrannus. However, they are shown as the plesiomorph sister group of the Passerida in Fig. 8.59A as

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Reproductive Biology and Phylogeny of Birds

one can confidently add two Corvida+Passerida synapomorphies: loss of the proximal centriole and enlargement of the dense fibers. Tyrannus has both centrioles and small dense fibers. These features require confirmation in most suboscines. Corvida have the oscine synapomorphy of a true helical membrane (imperfectly known but at least present as a microtubular bundle in the spermatid) but remain plesiomorphic in the short acrosome and midpiece, relative to the length of the nucleus. However, some exceptions to the typical acrosome: nuclear ratios, below unity in Corvida and above in Passerida are noted in section 8.10.10.1 and Table 8.6. Passerida. Two synapomorphies for the Passerida are well established from the large number of light microscope descriptions reviewed in this chapter and from new information, notably for Myrmecocichla formicivora and Philetairus socius. The most distinctive synapomorphy is the very long spiral midpiece differing from that of the other group with a long midpiece, the columbiforms (and possibly Coturnix japonica), in consisting of a single, spiral mitochondrial strand. The second is the elongation of the acrosome relative to the nucleus, though exceptionally, as recorded in Table 8.6, it remains shorter than the nucleus as in the suboscines and most Corvida. A more discriminating analysis should be attempted when a greater knowledge of sperm ultrastructure in a much larger range of species accrues. For instance, a detail of the mitochondrial helix, the presence of a fibrous helix, might well diagnose the Muscicapoidea and presence of a granular helix may have taxonomic value.

8.12

CONCLUSION

The author has attempted in this chapter to review works on the ultrastructure of avian spermatozoa, with some reference to light microscopical studies, with a view to demonstrating the diversity which exists in sperm structure in birds and phylogenetic patterns which may be discerned. A very preliminary phylogenetic analysis using PAUP has been presented. It is uncertain, when a more complete matrix for all groups is obtained, whether the considerable homoplasy which exists in avian sperm characters will permit the production of highly resolved trees. Nevertheless, the taxonomic and phylogenetic value of sperm characters is indisputable. Thus we have seen that even the highly peculiar, ‘neotenous’ sperm of the Eurasian bullfinch yielded up its identity as a passerine sperm when closely examined. Spermatozoal characters of paleognaths have confirmed crocodiloid and basal amniote relationships. They also allow confident identification of a spermatozoon with its ordinal or subordinal grouping in many cases and increasing discernment will presumably become possible as descriptions are improved. Thus it is already possible to recongize a distinctive sperm type for each of the groups Struthioniformes, Galloanserae, Galliformes, Anseriformes, Psittaciformes, Apodiformes, Columbiformes, Passeriformes, Corvida and Passerida.

Avian Spermatozoa: Structure and Phylogeny

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It is hoped that this compendium will prove a stimulus and aid to other researchers on avian spermatozoal morphology and will provide a background for those interested in sperm function and general biology.

8.13

ACKNOWLEDGMENTS

The author gratefully acknowledges facilities provided by the School of Integrative Biology, University of Queensland, and particularly the assistance, in procedures for electron microscopy, of Lina Daddow and Dr. David Scheltinga, who is also thanked for commenting on the manuscript. Professor Sandro Tripepi and Professor Alan Hodgson kindly provided unpublished micrographs and Dr. Claire Spottiswoode collected material in Africa. Dr. John Harshman provided many insightful comments on the manuscript and aided with cladistic computations. Professor T. R. Birkhead kindly provided some references and commented on the ms. Thanks are also due to the many publishers who have allowed reproduction of illustrations.

8.14

LITERATURE CITED

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Saita, A., Tripepe, S. and Longo, O. M. 1982b. Comparative observations on spermiogenesis II. Nuclear shaping in the absence ofa microtubular manchette in the spermatids of the bird Crotophaga ani (Cuculiformes). Bolletino di Zoologia 49: 115-123. Saita, A., Longo, O. M. and Tripepe, S. 1983. Osservazioni comparative sulla spermiogenesi. III. Aspetti ultrastrutturali della spermiogenesi di Jacana jacana (Charadriformes). Accademia Nazionale dei Lincei. (Rendiconti della Classe di Scienze fisiche, matematiche e naturali) 74: 417-430. Saita, A., Comazzi, M. and Perrotta, E. 1987. Electron microscope study of spermiogenesis in Caiman crocodylus L. Bollettino di Zoologia 4: 307-318. Samour, J. H. The Reproductive Biology of the Budgerigar (Melopsittacus undulatus): Semen Preservation Techniques and Artificial Insemination Procedures. Journal of Avian Medicine and Surgery 16: 39-49. Samour, J. H., Moore, H. D. and Markham, J. A. 1986. Semen collection and spermatozoa characteristics in budgerigars (Melopsittacus undulatus). The Veterinary Record 118: 397-399. Scheltinga, D. M. and Jamieson, B. G. M. 2003a. The Mature Spermatozoon. Pp. 203274. In D. M. Sever (ed). Reproductive Biology and Phylogeny of Urodela. Science Publishers, Inc., Enfield, New Hampshire, U.S.A. Scheltinga, D. M. and Jamieson, B. G. M. 2003b. Spermatogenesis and the Mature Spermatozoon: Form, Function and Phylogenetic Implications. Pp. 119-251. In B. G. M. Jamieson (ed). Reproductive Biology and Phylogeny of Anura, 2, Science Publishers, Inc., Enfield, New Hampshire, U.S.A. Scheltinga, D. M., Wilkinson, M., Jamieson, B. G. M. and Oommen, V. O. 2003. Ultrastructure of the mature spermatozoa of caecilians (Amphibia: Gymnophiona). Journal of Morphology 258: 179-192. Sibley, C. G. and Ahlquist, J. E. 1981. The phylogeny of Ratites birds as indicated by DNA-DNA Hybridization. Pp. 301-335. In G. C. Scudder and J. L. Reveal (ed). Evolution Today, Proceedings of the Second International Congress of Systematic and Evolutionary Biology. Hunt Institute for Botanical Documentation: Pittsburg. Sibley, C. G. and Ahlquist, J. E. 1990. Phylogeny and Classification of Birds: A Study in Molecular Evolution, Yale University Press, New Haven. Sibley, C. G., Ahlquist, J. E. and Monroe, B. L. 1988. A classification of the living birds of the world based on DNA-DNA hybridization studies. The Auk 105: 409-423. Soley, J. T. 1989. Transmission electron microscopy of Ostrich (Struthio camelus) sperm. Electron Microscopy Society of Southern Africa Proceedings 19: 145-146. Soley, J. T. 1993. Ultrastructure of ostrich (Struthio camelus) spermatozoa: I. Transmission electron microscopy. Onderstepoort Journal of Veterinary Research 60(2): 119-130. Soley, J. T. 1994a. Centriole development and formation of the flagellum during spermiogenesis in the ostrich (Struthio camelus). Journal of Anatomy 185: 301-313. Soley, J. T. 1994b. Ostrich sperm ultrastructure—evidence of a close link between the ratites and tinamous. Pp. 5.27-25.28. In N. M. Bradley and J. M. Cummins (eds). Proceedings of the VIIth International Symposium on Spermatology, Cairns, Australia. Murdoch University, Western Australia. Soley, J. T. 1996. Differentiation of the acrosomal complex in ostrich (Struthio camelus) spermatids. Journal of Morphology 227(1): 101-111. Soley, J. and Groenwald, H. B. 1999. Reproduction. Pp. 129-158. In D. C. Deeming (ed). The Ostrich: Biology, Production and Health, CAB International: Cambridge.

# Reproductive Biology and Phylogeny of Birds Soley, J. T. and Roberts, D. C. 1994. Ultrastructure of ostrich (Struthio camelus) spermatozoa: II. Scanning electron microscopy. Onderstepoort Journal of Veterinary Research 61(3): 239-246. Sprando, R. L. and Russell, L. D. 1988. Spermiogenesis in the red-ear turtle (Pseudemys scripta) and the domestic fowl (Gallus domesticus): A study of cytoplasmic events including cell volume changes and cytoplasmic elimination. Journal of Morphology 198: 95-118. Stanley, H. P. 1967. The fine structure of spermatozoa in the lamprey Lampetra planeri. Journal of Ultrastructure Research 19: 84-99. Stapel, S. O., Leunissen, J.A.M., Versteeg, M., Wattel, J. and de Jong, W. W. 1984. Ratitesas oldest offshoot of avian stem—evidence from a-crystalline A sequences. Nature 311: 257-259. Swan, M. A. 1985. Transmission electron microscopy of impaired spermatogenesis in an avian hybrid. Gamete Research 12: 357-371. Swan, M. A. and Christidis, L. 1987. Impaired Spermatogenesis in the Finch Hybrid Lonchura-Castaneothorax by Lonchura-Punctulata Transmission Electron Microscopy and Genetic Analysis. Gamete Research 17(2): 157-172. Swofford, D. L. 2001. Phylogenetic analysis using parsimony, version 4.0, Sinauer Associates, Sunderland, Massachusetts. Temple-Smith, P. D. and Bedford, J. M. 1980. Sperm maturation and the formation of sperm pairs in the epididymis of the opossum, Didelphis virginiana. Journal of Experimental Biology 214: 161-171. Thurston, R. J. and Hess, R. A. 1987. Ultrastructure of Spermatozoa from Domesticated Birds Comparative Study of Turkey Chicken and Guinea Fowl. Scanning Microscopy 1(4): 1829-1838. Thurston, R. J., Hess, R., Hughes, B. L. and Froman, D. P. 1982. Ultrastructure of the guinea fowl (Numidia meleagris) spermatozoon. Poultry Science 61: 1738-1743. Tingari, M. D. 1973. Observations on the fine structures of spermatozoa in the testes and excurrent ducts of the male fowl, Gallus domesticus. Journal of Reproduction and Fertility 34: 255-265. Tripepi, S. and Perrotta, E. 1991. Spermiogenesis and sperm of passerine birds. Pp. 1021-1023. In Baccetti B. (ed.), Comparative Spermatology 20 Years After. Serono Symposia Publications, vol. 75. Raven Press, Rome. Tripepi, S., Saita, A., and Longo, O. 1984. Studio al M.E. sulla spermiogenesi di Apus apus (Uccelli, Apodiformi). Bolletino di Zoologia 51 (suppl) p. 108. Tripepi, S., Tavolaro, P. and F. Rossi 1991. The evolution of microtubular organization during spermiogenesis in birds. Pp. 631-636. In G. Ghiara (ed). Selected Symposia and Monographs U.Z.I, 4, Symposium on the Evolution of Terrestrial Vertebrates. Mucchi, Modena. Tripepi, S. and Perrotta, E. 1992. Spermiogenesis and sperm of passerine birds. Pp. 1021-1023. In B. Baccetti (ed). Comparative Spermatology Twenty Years After, Raven Press, New York. Vernon, G. G. and Woolley, D. M. 1999. Three-dimensional motion of avian spermatozoa. Cell Motily and the Cytoskeleton 42: 149-161. Wagley, L. 1980. SEM of peregrine spermatozoa. Western Release Program - 1980. In T. J. Cade and P. R. Dague (eds). The Peregrine Fund Newsletter. No. 8, Fall 1980: 4. Wen, Z. Q., Zheng, G. M., Song, X. F., Sun, Q. Y. and Chen, D. Y. 1997. Ultrastructure of spermatozoon of Cabot’s tragopan (Tragopan caboti). Acta Zoologica Sinica 43(2): 127-132.

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Woolley, D. M. 1995. The Structure of the Spermatozoon of the Japanese Quail, Coturnix coturnix L. var. japonica. Acta Zoologica (Copenhagen) 76(1): 45-50. Woolley, D. M. and Brammall, A. 1987. Direction of sliding and relative sliding velocities within trypsinized axonemes of Gallus domesticus. Journal of Cell Science 88: 361-371. Xia, L., Clermont, Y., Laill, M. and Buckland, R. B. 1985. Spermiogenesis of the rooster: an electron microscopic study. Poultry Science 64: 201 abstract. Xia, L., Clermont, Y., Lailli, M. and Buckland, R. B. 1986. Evolution of the endoplasmic reticulum during spermogenesis of the rooster: an electron microscope study. The American Journal of Anatomy 177: 301-312. Xia, L., Lalli, M. F., Ansah, G. A. and Buckland, R. B. 1988. Ultrastructure of Fresh and Frozen-Thawed Spermatozoa of High and Low Fertility Lines of Chickens. Poultry Science 67(5): 819-825. Yasuzumi, F. and Yamaguchi, S. 1977. Some aspects of spermiogenesis in the Domestic Pigeon. Okajimas Folia Anatomica Japonica 54: 139-174. Yasuzumi, G. 1974. Electron Microscope Studies on Spermiogenesis in Various Animal Species. International Review of Cytology 37: 53-119. Yasuzumi, G. and Sugioka, T. 1966. Archives of Histology, Japan 27: 259-265. Yasuzumi G and Sugioka, T. 1971. Spermatogenesis in animals as revealed by electron microscopy. XXI. Microkaryosomes and microtubules appearing during spermiogenesis of the lovebird Uroloncha striata var. domestica flower. Zeitschrift für Zellforschung und Mikroskopische Anatomie 114(4): 451-459.

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CHAPTER

9

Testis Size, Sperm Size and Sperm Competition James V. Briskie1 and Robert Montgomerie2

9.1

INTRODUCTION

Almost since the moment animal semen was first viewed under the microscope, it was clear that the shape and size of spermatozoa varied from one species to the next (Leeuwenhoek 1697). Early comparative biologists similarly noted striking differences in the size of the testis across different species (Ray 1678). Although the general features of the male reproductive system are rather conservative across the 10,000 or so extant species of birds, recent work has confirmed large interspecific variation in the sizes of both testes and sperm. For example, the combined mass of the left and right testes of breeding Alpine accentors (Prunella collaris) is 3.0 g and comprises about 8% of adult body mass, while that of the Zebra finch (Taeniopygia guttata) is only 0.05 g or 0.4% of its body mass (Birkhead et al. 1991; Nakamura 1990). Likewise, a single spermatozoon of the Yellow warbler (Dendroica petechia) averages 277 µm in length, while that of the Red-backed shrike (Lanius collurio) is only 43 µm long, a six-fold difference in size (Briskie et al. 1997). Why should testes mass and sperm size differ so much from one species to the next? In this chapter we examine some of the reasons for variation in both testis size and sperm size in birds. A major turning point in our understanding of this variation both within and between species, came with the seminal paper by Parker (1970) in which he introduced the concept of sperm competition. Parker (1970) defined sperm competition as occurring whenever a female mated with more than one male during the course of a single breeding attempt. 1

School of Biological Sciences, University of Canterbury, Private Bag 4800, Christchurch, New Zealand, E-mail: [email protected] 2 Department of Biology, Queen’s University, Kingston, Ontario K7L 3N6, Canada. E-mail: [email protected]

#" Reproductive Biology and Phylogeny of Birds For birds, this might occur when a female engages in an extrapair copulation with a neighboring male, or when her social mating system involves some degree of polyandry. Sperm storage by female birds (Van Drimmelen 1946; Hatch 1983; Briskie and Montgomerie 1993) increases the risk of sperm competition as inseminations even weeks apart can be involved in the fertilization of a single clutch of eggs (Oring et al. 1992; Birkhead 1998a). Thus the intensity of sperm competition varies across species and it soon became clear that differences in the reproductive morphology of birds could be linked to this variation (Møller 1991; Birkhead and Møller 1992a; Briskie and Montgomerie 1992, 1993, 1997). The advent of modern genetic profiling techniques has further invigorated the field as variation in the morphology of reproductive structures was shown to be related to differences in the sexual or genetic (rather than social) mating strategies of males and females, both within and between species (Møller and Briskie 1995; Briskie et al. 1997). Our objective in this chapter is to take an evolutionary approach to the gross morphologies of avian testes and sperm, particularly with respect to variation in size (detailed morphological description of the avian testis is provided in Chapter 2, the process of sperm production in Chapter 7, and sperm ultrastructure in Chapter 8). To do this, we review mainly studies that have used the comparative method (Harvey and Pagel 1991) to investigate adaptive patterns. Thus our approach focuses on deriving explanations for the evolution of the wide diversity of testes and spermatozoa sizes in birds (i.e., the “ultimate” explanations for traits). The quest for such explanations involves searching for potential explanatory variables that correlate with the trait of interest, and then testing (either through experiments or further comparative analyses) whether these explanations can be generalised to other taxa. Despite more than a century of detailed anatomical and physiological work on avian reproduction, modern comparative methods with appropriate statistical rigor have been applied to understanding the evolution of testis and sperm size only during the last two decades, and experimental studies are just beginning (Birkhead et al. 2005). As the application of these approaches to the study of avian reproduction is still in its early stages, we start by describing variation in the number, position and size of the avian testis, and then review the adaptive hypotheses that have been proposed to explain this variation. We also survey variation in sperm size and review the evidence explaining the evolution of this variation. We conclude by outlining gaps that remain in our knowledge and suggesting where future workers can make a contribution.

9.2 9.2.1

EVOLUTION OF THE AVIAN TESTIS Location and Number of Testes

Unlike the male genitalia of most mammals, those of birds are not obvious externally. Instead, the testes of birds are positioned deep inside the body cavity, visible only after dissection and removal of the viscera (Fig. 9.1). No species of bird is known to have their testes in an external scrotum, and there

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Fig. 9.1 Urogenital tract of a male House sparrow during (right) and outside (left) the breeding season, as revealed by dissection. On the right-hand drawing, the right testis is indicated by a dashed line to reveal the location and size of the epididymal duct. Presumably drawn to scale but no scale bars shown in the original. Modified after Witschi, E. 1961. Pp. 115-168. In A. J. Marshall (ed.), Biology and Comparative Physiology of Birds, Vol. II. Academic Press, New York, Fig. 18.

#$ Reproductive Biology and Phylogeny of Birds is no known variation in the position of the testes internally, unlike the situation in mammals (Freeman 1990; Werdelin and Nilsonne 1999). External genitalia could be disadvantageous in flight by reducing streamlining and this may have prevented any adaptive change in testicular positioning in birds. On the other hand, the position of the testes in birds (Fig. 9.1) is similar to that in reptiles and other lower vertebrates, and the testes may have been retained in this ancestral position simply in the absence of any selective pressure to change. The inhibition of sperm production at high body temperatures has often been cited to explain the externalization of the testes in some mammals (Moore 1926; Werdelin and Nilsonne 1999). However, this cannot be a general explanation as birds also maintain high body temperatures, often exceeding that found in mammals (McNab 1966; Prinzinger et al. 1991), and avian testes are not kept cooler than other internal organs (Beaupré et al. 1997). Interestingly, Wolfson (1954) found that the cloacal protuberance of passerine birds (Chapter 9) can store sperm at cooler than body temperatures. He proposed this structure functions in an analogous manner to the externalized testes of mammals, thus improving sperm storage and the later stages of sperm maturation. The absence of a cloacal protuberance in non-passerines appears consistent with their generally lower body temperature than passerines (Lasiewski and Dawson 1967). Unfortunately, little work has been done on sperm storage in relation to temperature in birds since Wolfson’s (1954) pioneering work, and experimental studies are needed to confirm the effects of high temperatures on the storage of sperm in both passerine and non-passerine birds. With few exceptions, male birds have two testes, whereas in females the right ovary and oviduct are usually rudimentary. A few cases each of single and supernumerary testes have been reported but these are generally considered to be developmental abnormalities (e.g., Ferris 1933; Hocking 1992). The only exception to males having two testes appears to be the Centropus coucals, in which the left testis is normally undeveloped or absent altogether (Rand 1933; Chapin 1939; Ligon 1997). Ligon (1997) suggests that the loss of the left testis in the coucals is related to lower testosterone levels in these sex-role reversed species. However, it is difficult to see why hormone titres could not be lowered by reducing the quantity of endocrine tissue in each testis rather than by the loss of an entire gonad with, presumably, a reduction in sperm production. The absence of a left testis in male coucals is also unlikely to be homologous to the loss of the right oviduct in female birds, as different sides of the body are involved (Ligon 1997). The loss of the right oviduct in female birds has been considered an adaptation to the cost of flight (Witschi 1935), but it is not known if flight is particularly costly for coucals or would have been made less costly by the loss of the left testis. Alternatively, if sex-role reversal has led to a reduction in the risk of sperm competition, this may have favored the reduction in testis size. Thus, perhaps the entire loss of one testis simply reduces the costs of maintaining two gonads. Further

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information on the levels of promiscuity in coucals relative to other species with and without sex-role reversal is needed before this idea can be assessed.

9.2.2

Asymmetry in Testis Size

Differences in the size of the left and right testes are widespread among vertebrates (Yu 1998). In birds, the most common pattern is for the left testis to be larger than the right, but there are exceptions, with no difference in size in some species and a larger right testis in others (Friedmann 1927; Lake 1981; Kimball et al. 1997; Birkhead et al. 1998; Merilä and Sheldon 1999). For example, the left testis in the Sedge warbler (Acrocephalus schoenobaenus) averages 0.087 g, and is significantly heavier than the right testis, which weighs only 0.074 g (Birkhead et al. 1997). In contrast, the left (0.326 g) and right (0.328 g) testes of the Tree swallow (Tachycineta bicolor) do not differ significantly in size (Kempenaers et al. 2002). Stanley and Witschi (1940) suggested that the widespread right/left asymmetry in testis development was a by-product of selection for asymmetry in the female reproductive tract, in which only the left side develops. The proximate basis for such an asymmetry may lie in the uneven migration of primordial germ cells to the left and right sides of the body during the development of the embryo (Witschi 1935). As the germ cells migrate to each side of the body before sexual differentiation occurs, the differential attraction of the germ cells to the left side may be so advantageous to females that it may persist despite any disadvantage accrued to the development of the right testis of males (Witschi 1935). However, it is not clear whether the number of primordial germ cells directly affects the size of the mature gonad, especially considering the large seasonal fluctuations in avian testis size that occur through repeated mitosis of the germ cells. Furthermore, the development of the left reproductive tract in female birds is nearly universal whereas there is considerable variation in presence or absence of testis size asymmetry in males, and the magnitude of this asymmetry varies widely both within and among bird species. Thus, in at least some species, testicular asymmetry cannot be a simple by-product of strong selection on female reproductive asymmetry or germ cell number. As a result, a number of authors have sought an adaptive explanation for this pattern. Based on patterns of testicular asymmetry in Barn swallows (Hirundo rustica) and House sparrows (Passer domesticus), Møller (1994) proposed that the smaller right testis plays a compensatory role, developing to a similar size as the normal left testis only when the latter does not develop properly (see also Domm and Juhn 1927). He reasoned that males in poor condition might be more likely to have a malfunctioning left testis and as a result compensate by developing a larger right testis. Thus the degree of testicular asymmetry could be seen as a measure of developmental homeostasis, with the highest quality males sporting the most asymmetrical testes and lower quality males with smaller, more symmetrical testes. An assumption of this hypothesis is that it is more costly to support two similar-sized testes than two testes of

#& Reproductive Biology and Phylogeny of Birds unequal size. It is not clear what these costs might be but they could include increased flight costs (see below). In support of his compensation hypothesis, Møller (1994) found that the degree of asymmetry in testis size varied significantly with male quality as measured by tail streamer length in the swallow, and black bib size in the sparrow. In both species, testis asymmetry was largest in individuals having the most elaborate plumage ornaments, and thus presumed to be of highest quality. A number of subsequent studies across a variety of species have tested Møller’s (1994) compensation hypothesis, but to date no support has been found (Birkhead et al. 1997; Kimball et al. 1997; Merilä and Sheldon 1999; Kempenaers et al. 2002; Graves 2004). Thus, it seems unlikely that this hypothesis can provide a general explanation for testicular asymmetry in birds. Indeed, in the only experimental test of this hypothesis, Birkhead et al. (1998) found the opposite pattern: the difference between the masses of the left and right testes was even greater in Zebra finches whose condition was experimentally reduced. Thus, the patterns shown by Møller (1994) are a curious artefact that deserve further investigation. Despite the general lack of relation between male quality and the degree of testicular asymmetry, a number of studies have found that testicular asymmetry increases with age (Birkhead et al. 1997; Kimball et al. 1997, Merilä and Sheldon 1999). Birkhead et al. (1997) suggested that males with greater testicular asymmetry might survive better, although it is not clear how this would occur unless directional asymmetry was related to male quality (as suggested by Møller 1994) or if testicular asymmetry was more costly to maintain. Graves (2004) found that older Black-throated blue warblers (Dendroica caerulescens) exhibited a higher level of testicular asymmetry than yearling males, as a result of the disproportionate enlargement of the left testis with age. As older males in this species also have larger testes than younger males, variation in the absolute and relative sizes of the left and right testes may be related to variation in mating strategy (Graves 2004; see also Hill 1994; Merilä and Sheldon 1999). For example, in some species, older males appear to be preferred as mates and as extrapair copulation partners (Weatherhead 1984), and engaging in copulations with more than one female may require higher rates of sperm production (Cartar 1985) and thus a greater investment in testis tissue. Despite this, it is not obvious why selection for increased testes size would necessarily favor an increased testicular asymmetry. It is possible that testicular asymmetry functions to reduce the combined testes mass, thereby reducing the costs of flight (as has been suggested for the loss of the right oviduct in females; Witschi 1935). Flightless species, or those that fly only occasionally, might then be expected to show less testicular asymmetry than strongly aerial species (Kimball et al. 1997). Kempenaers et al. (2002) did not find any support for this hypothesis in the highly aerial Tree swallow which, contrary to expectations, had relatively symmetrical testes. However, a wider comparative study is needed to test this idea more thoroughly. It is worth noting that this cost of flight hypothesis might not just

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involve a reduction in total mass, but also a redistribution of mass to increase overall balance along the longitudinal axis of the bird. For example, most of the liver resides on the right side of the body in birds and other organs (e.g., gizzard, spleen) are similarly placed asymmetrically, potentially creating a mass asymmetry between the left and right sides of the body. It is not clear why these organs are positioned asymmetrically in the first place, but a larger left testis might compensate any imbalance that could cause roll during flight. We are not aware that this idea has been tested. Finally, in some passerines, it has been observed that males exhibit a leftside bias when mounting during copulation (Chapters 3 and Volume 6B, Chapter 6). As only the left ovary of the female is functional in most species (Chapter 4), and the left oviduct opens on the left side of the cloaca, it could be advantageous for males to mount from the left if this results in sperm being placed closer to this opening. In turn, this might favor differential investment in sperm production by the two testes, resulting in a left-biased testicular asymmetry if sperm from the left testis and vas deferens is more likely to reach the left oviductal opening. Unfortunately, the dynamics of ejaculation are not known in sufficient detail to test this idea quantitatively. However, as some species have a right-side bias during copulation (e.g., American avocet, Recurvirostra americana, and Black-necked stilt, Himantopus mexicanus; Sordahl 2001), it should be possible to compare patterns of testis asymmetry with copulation behavior to evaluate this hypothesis.

9.2.3

Sperm Production and Testis Size

Although the testes play a role in the production of both hormones and sperm (Chapter 7), the vast majority of tissue in the testes is devoted to the latter function. Thus it is not surprising that larger testes are capable of producing more sperm (Fig. 9.2; Møller 1988, 1989; Schärer et al. 2004). Sperm production rates have been measured in only a few bird species, but rates appear to vary interspecifically (Fig. 9.2). For example, sperm production in the socially monogamous Zebra finch averages 1.9 ¥ 106 sperm/day (Birkhead et al. 1995), while males in the promiscuous White-winged fairy-wren (Malurus leucopterus) produce an average of 646 ¥ 106 sperm/day (Tuttle and Pruett-Jones 2004). Sperm production can also vary among males within a species. Birkhead et al. (1994) found that testes mass in one population of House sparrows ranged from about 0.45 g to 1.0 g, and was correlated with the number of sperm stored in the seminal glomera, suggesting that individuals with larger testes produce more sperm. A similar pattern was found between males with different testis sizes in the Zebra finch (Birkhead et al. 1993a). It should be noted that most data on rates of sperm production by individual males have been collected from domesticated and captive birds, and that a number of different techniques were used. More information from wild birds is needed to confirm that studies of captive birds provide realistic rates of sperm production, and to determine why sperm production rates vary among different males within a population.

#  Reproductive Biology and Phylogeny of Birds

Fig. 9.2 Relation between daily sperm production and testes volume in birds (Model II regression, log(sperm production per day) = 8.11 + 0.69 log (testis volume). r2 = 0.52, n = 9 species. Data from Tuttle, E. M. and Pruett-Jones, S. 2004. Animal Behaviour 68: 541-550, Tables 1 and 4.

Species with large testes not only produce more sperm overall, but apparently also produce more sperm per ejaculate (Møller 1988). Actual ejaculate size (the number of sperm per ejaculate) is known for only a few species, and Møller’s (1988) analyses were based on data from early studies of ejaculate size in domestic birds that used collection methods (e.g., electroejaculation, abdominal massage) likely to provide biased estimates of sperm number. More recently, dummy models fitted with false cloacas have been used to collect ejaculates deposited directly by males, and have provided estimates that are more realistic with respect to natural copulations (Pellatt and Birkhead 1994). For example, ejaculates collected in this manner average 3.89 ¥ 106 sperm in Zebra finches, 12.5 ¥ 106 sperm in Red-winged blackbirds (Agelaius phoeniceus; Westneat et al. 1998), and 124 ¥ 106 in Peafowl (Pavo cristatus; Birkhead and Petrie 1995). In all studies to date, ejaculate size has been found to be highly variable, both among males and within males. In the Peafowl, for example, ejaculates contained from 4.2 ¥ 106 to 248 ¥ 106 sperm (Birkhead and Petrie 1995). Reasons for this variation are not clear, although at least some of it may be adaptive. For example, Nicholls et al. (2001) found that male Bank swallows (Riparia riparia) increased the size of their ejaculates when faced with rival males (i.e., more intense sperm competition). Ejaculate sizes also decline with time since last copulation when successive copulations occur within a short period of time, such that sperm depletion may limit the rate of insemination

Testis Size, Sperm Size and Sperm Competition

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(Birkhead et al. 1995). This in turn should lead to selection favoring increased testis size, and thus increased sperm production, in species with high copulation rates and high levels of promiscuity (Birkhead et al. 1993b).

9.2.4

Seasonal Change in Testis Size

Most bird species undergo cyclical patterns of testicular activity and inactivity (Murton and Westwood 1977), with a period of rapid testicular re-growth or recrudescence in mature males just prior to the start of the breeding season. After breeding, the testes regress and remain inactive until the next breeding season, when they undergo another phase of recrudescence (e.g., Fig. 9.3). The difference in testis size between the non-breeding and breeding seasons can be dramatic (Fig. 9.3). In the House crow (Corvus splendens), for example, the combined mass of both testes in the breeding season is more than 75 times that in the non-breeding season (Dang and Guraya 1978). Increases of 300500 fold between the non-breeding and breeding season have been recorded in the Eurasian tree sparrow (Passer montanus; Lofts and Murton 1973), Common magpie (Pica pica; Erpino 1969), and Brambling (Fringilla montifringilla; Marshall 1961). Changes in testis size over the season are well documented among temperate species in the Northern Hemisphere, but cyclical changes in testis size also occur in tropical environments (Fig. 9.3C, D; Wikelski et al. 2003). Birds living in relatively unpredictable environments, on the other hand, may maintain their testes in a relatively developed state to take advantage of rapid improvement in breeding conditions (Immelmann 1971; Zann et al. 1995). The testes may also remain enlarged for the entire breeding season in species with multiple breeding attempts per season, although sperm production may increase during the periods that coincide with the fertility of their mates (Lofts and Murton 1973). Studies on patterns of testis recrudescence and regression typically focus on a species-specific pattern, but there can also be considerable variation among individuals within a population on the timing of the testicular cycle. For example, Dawson (2003) found that older European starlings (Sturnus vulgaris) initiated testicular recrudescence 3-4 weeks earlier than younger males and did not enter the regression phase until 2 weeks later than younger males. This translated into a 50% longer period of spermatogenic activity in older males. In the Winter wren (Troglodytes troglodytes), males that underwent recrudescence earlier also nested earlier than males that delayed testicular growth (Evans and Goldsmith 2000). Among species breeding year round (e.g., some tropical species), each individual may continue to cycle through periods of recrudescence and regression such that at any given time some males have active spermatogenesis while others do not (Lofts and Murton 1968). In contrast to their wild counterparts, domestic fowl (Gallus g. domesticus) and some domesticated birds have little seasonal change in testis size and are thus capable of producing sperm year round. In the domestic fowl, this pattern has likely been the product of selective breeding to maximize their rate of

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Reproductive Biology and Phylogeny of Birds

Fig. 9.3 Seasonal varition in testis size in some temperate zone (A, B) and tropical (C, D) bird species. A European blackbird (Turdus merula): testis width (mm) of urban birds. Modified after Partecke, J., Van’t Hof, T. and Gwinner, E. 2004. Proceedings of the Royal Society B 271: 1995-2001, Fig. 1(i). B House sparrow: testis mass (mg); shaded zones indicate three successive broods in this rural population. Modified after Hegner, R. E. and Wingfield, J. C. 1990. Pp 123-135. In J. Pinowski and D. Summers-Smith (eds), Granivorous Birds in the Agricultural Landscape. INTECOL, Warsaw. C Red-billed quelea (Quelea quelea): maximum testis diameter (mm); birds maintained on constant 12-h photoperiod over two years. Modified after Lofts, B. and Murton, R. K. 1973. Pp. 1-107 In D. S. Farner, J. R King and K. C. Parkes (eds). Avian Biology, volume 3. Academic Press, New York, Fig. 7. D Red-vented bulbul (Pycnonotus cafer): testis volume (mm3); birds maintained in captivity on natural light cycle. Modified after Lal, P. and Thapliyal, J. P. 1982. General and Comparative Endocrinology 48: 98-103, Fig. 1.

reproduction (Sossinka 1982). Wyndham et al. (1981) suggested that the abundance of food in captivity might also explain why domesticated Budgerigars (Melopsittacus undulatus) undergo reduced periods of testicular regression relative to their wild progenitors. The initiation of testicular recrudescence appears to be under photoperiodic control, at least for temperate-breeding passerines, although it can be modified by temperature, weather, food supply, and behavioral interactions (Lofts and Murton 1968; Lal and Thapliyal 1982; Follett 1984). The histological and physiological changes that take place during the

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testicular cycle are well documented in a variety of bird species (Chapters 2 and 7).

9.2.5

Body Size and Testis Size

Within species, larger males tend to have larger testes (Fig. 9.4A; Rising 1987; Møller and Erritzøe 1988; Merilä and Sheldon 1999; Evans and Goldsmith 2000; Brown and Brown 2003), though body size often accounts for a small proportion of variation in testis size. In 48 populations across the range of Savannah sparrows (Passerculus sandwichensis), for example, only 7% of the variation in testis size among individuals (n = 1157) was explained by variation in body mass, though 29% of the variation in mean testis mass among populations was explained by mean body mass (Fig. 9.4A; Rising 1987). Older males are sometimes larger than young males, either due to growth or differential mortality, but variation in age alone cannot always explain the positive relation between testis size and body size. For example, older male Black-headed grosbeaks (Pheucticus melanocephalus) had relatively larger testes in an analysis controlling for body size (Hill 1994). The greater relative size of testes in larger males may thus be related to a high cost of testicular recrudescence and maintenance that only males in good condition are able to bear. Testis size is also positively correlated with body size across species (Møller 1991; Pitcher et al. 2005). For example, in a sample of 1010 bird species, Pitcher et al. (2005) found that almost half of the inter-specific variation in testis mass was explained by body mass (Fig. 9.4B). In a similar study, Møller (1991) noted that the slope of the regression of testes mass on body mass was significantly less than one (see also Fig. 9.4B). Thus larger-bodied species have relatively smaller testes than smaller species, though the reasons for this are unknown. Because of the strong effect of body mass on testis size, most studies of testis size variation have controlled for body size either by examining residual testis size, or by entering body mass as an independent variable in multiple regression analyses.

9.2.6

Age-related Changes in Testis Size

Birds of many species mature and initiate reproduction during the first breeding season after they hatch. However, among bird species for which data are available from the breeding season, the testes are generally larger in adult birds (2 years and older) than among males breeding for the first time (Wright and Wright 1944; Davis 1958; Selander and Hauser 1965; Foster 1987; Deviche et al. 2000). In Cliff swallows (Petrochelidon pyrrhonota), testes size was smallest in birds aged 1 and 2 years but did not vary with age in males 3 years old or more (Fig. 9.5; Brown and Brown 2003). As mentioned above, the size of the left and right testes may also change differentially with age, such that the degree of testicular asymmetry differs between younger and older males. In the Black-headed grosbeak, males breeding at the end of their first year do not develop adult breeding plumage and thus their smaller testes may be

# " Reproductive Biology and Phylogeny of Birds

Fig. 9.4 Relation between testes size and body size in birds; model II regressions are plotted. A Intraspecific variation among 48 populations (n = 9-45 males per population) of Savannah sparrows [log(testis volume) = –0.81 + 1.25 log(femur length), r2 = 0.29]; single testis volume calculated from length and width measured on freshly-killed specimens. Data from Rising, J. D. 1987. Wilson Bulletin 99: 6372, Table 1. B Interspecific variation among 1010 bird species [log(testes mass) = –2.04 + 0.89 log(body mass), r2 = 0.47]. Testes mass calculated as the combined mass of both testes averaged over at least five breeding males per species; estimated from linear measurements, mainly from museum specimens. Redrawn from data in Pitcher, T. E., Dunn, P. O. and Whittingham, L. A. 2005. Journal of Evolutionary Biology 18: 557-567, Fig. 1.

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Fig. 9.5 Age-related changes in testes size in the Cliff swallow. Testes size is measured as residuals from the regression of the combined testes volume (calculated from length and width measurements) on a multivariate measure of body size. Numbers near data points are sample sizes. Modified after Brown, C. R. and Brown, M. B. 2003. Behavioral Ecology 14: 569-575, Fig. 1.

part of a more general, adaptive delay in reproduction (Hill 1994). As rates of sperm production are related to testis size (Fig. 9.2), younger males would not produce as much sperm as older males. Young males in the lek-breeding Sharp-tailed grouse (Tympanuchus phasianellus) also have smaller testes than older males and are more likely to occupy peripheral territories on the edge of a lek, while older males dominate the central territories (Tsuji et al. 1992). Nitchuk and Evans (1978) confirmed that peripheral males in this species do produce less sperm than central males, but it is not clear that females would obtain fewer sperm by mating with younger males, nor if this would reduce the female’s fertility relative to mating with older males. Although some studies of parentage have found that younger males were more likely to be cuckolded (e.g., Wetton et al. 1995), this could be due to female choice or the failure of young males to protect mates from extrapair copulation attempts, rather than fewer sperm being inseminated by younger males. In some polygynous species (e.g., Graves 2004), younger males are also less likely to attract multiple mates and thus may have smaller testes as an adaptation to the lower number of sperm required to fertilize the ova of a single female (Cartar 1985).

9.2.7

Geographic Variation in Testis Size

Until recently, geographic variation in testis size within species has either been ignored, or has been assumed to be small relative to that observed among species (Pitcher and Stutchbury 1998). For some species the latter appears to

# $ Reproductive Biology and Phylogeny of Birds be true. For example, Graves (2004) found no geographic variation in testis size among 25 populations of the Black-throated blue warbler across its entire North American range. However, for most other species examined to date, testis size appears to vary with geographic location, with the most common pattern an increase in testis size with latitude (Fig. 9.6A-C; Pitcher and Stutchbury 1998; Merilä and Sheldon 1999). Such an increase in testis size with latitude has been suggested to result from either more intense sperm competition with latitude (Pitcher and Stutchbury 1998; Merilä and Sheldon 1999) or the shortened breeding season at high latitudes, which may require high rates of daily sperm production (Kenagy and Trombulak 1986). There are few intraspecific data available on the concomitant levels of sperm competition, breeding seasonality and testis size that would be needed to test either of these ideas. The opposite pattern, seen in Savannah sparrows (Fig. 9.6D), is harder to explain but may be influenced by the larger number of broods and thus longer breeding season at lower latitudes (Rising 1987).

Fig. 9.6 Relation between relative testis size (RTS) and latitude in different bird species. A House finch (Carpodacus mexicanus): 8 populations. B Red-eyed vireo: 19 populations. C Greenfinch (Carduelis chloris): 8 populations. Modified after Merilä, J. and Sheldon, B. C. 1999. Behavioral Ecology and Sociobiology 45: 115123, Fig. 2. D Savannah sparrow: 24 populations. Modified after Pitcher, T. E. and Stutchbury, B. J. M. 1998. Canadian Journal of Zoology 76: 618-622, Fig. 1a, c, d. In A, B and D, RTS was measured as the ratio (¥ 103) of testis mass (calculated from length and width) to body mass; in C, RTS is testis length in mm, correcting for age and body size. Model II regression lines shown to illustrate trends; all relations were significant in the original analyses.

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Broader patterns of geographic variation in testis size have also been examined among species. In a comparative study analyzing data from a variety of temperate and tropical species, Stutchbury and Morton (1995) found that testis size was generally smaller in a small sample of Neotropical birds. Such a pattern might be expected if sperm competition is less intense in tropical environments. Further work is needed to determine whether the intensity of sperm competition varies geographically. In a much larger sample of species, Pitcher et al. (2005) confirmed that testis size varies across geographic regions, with species living on the Eurasian and North American landmasses having larger relative testis size than species in South America and Australasia. It is unclear, however, why residence in a particular continental region should influence testis size. It seems more likely that such geographic differences are due to confounding variables that vary between the regions (e.g., differences in taxonomic representation, climate, body size, mating systems, etc.).

9.2.8

Testis Size and Mating System

Even when differences in season, age, latitude and body size are taken into account, testis size varies dramatically among bird species. Harcourt et al. (1981) suggested a simple explanation for this pattern based on sperm competition theory (Parker 1970), and found that primate species in which females copulate with more than one male had relatively larger testes than monogamous species. As the rate of sperm production increases with testis size (Fig. 9.2), Harcourt et al. (1981) argued that an evolutionary increase in testes mass is favored whenever a male’s sperm competes with the sperm of other males to fertilize a female’s ova. The advantage of producing more sperm in such situations has been likened to a lottery in that the more sperm produced, the greater the probability of securing fertilizations; increased sperm numbers could function either to dilute or displace the sperm of rival males. Subsequent studies have confirmed positive associations between testis size and the intensity of sperm competition across a range of taxa, including birds, in general (Fig. 9.7A; Birkhead and Møller 1998), and waterfowl (order Anseriformes), in particular (Fig. 9.7B: Coker et al. 2002). Testis size is particularly large in species with polygynandrous mating systems such as Dunnock (Prunella modularis; Birkhead et al. 1991), Smith’s longspur (Calcarius pictus; Briskie 1993), and Superb fairy-wren (Malurus cyaneus; Mulder and Cockburn 1993). Combined testes mass in these species varies from 4-8% of body mass, compared to only about 1% for birds in general (Pitcher et al. 2005). In polygynandrous species, both males and females regularly mate with multiple partners during the span of a single breeding attempt. Although large testes may partly be an adaptation to avoid sperm depletion (see below; Cartar 1985), the high levels of polyandry by females in these species also means that sperm competition is particularly intense. Thus, species in which females have highly promiscuous mating behavior tend to have larger testes than species with single-male mating

# & Reproductive Biology and Phylogeny of Birds

Fig. 9.7 Relative testes mass of bird species with different social mating systems. Social mating systems are shown from left to right in order of increasing expected intensity of sperm competition; relative testes mass measured as residuals from the regression of combined testes mass on body mass; number of species is shown at the base of each bar. A Social mating systems in 1002 bird species (MON = monogamy, CPR = cooperative breeding, MPG = monogamy with some males polygynous, PGY = polygyny, LEK = lekking, PAN = polyandry). Modified after Pitcher, T. E., Dunn, P. O. and Whittingham, L. A. 2005. Journal of Evolutionary Biology 18: 557-567, Fig. 4b. B Social mating systems in 29 waterfowl species (MON = monogamous, IFC = infrequent forced extrapair copulations, FFC = frequent forced extrapair copulations, PRM = promiscuous). Modified after Coker, C. R., McKinney, F., Hays, H., Briggs, S. V. and Cheng, K. M. 2002. Auk 119: 403413, Fig. 4A.

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systems (Fig. 9.7A; Møller 1991; Birkhead and Møller 1992a; Rising 1996; Pitcher et al. 2005). This result is further supported by analyses using levels of extrapair parentage from studies of genetic paternity in birds. Thus species with high proportions of extrapair young in their nests have significantly larger testes than expected for their body size (Fig. 9.8; Møller and Briskie 1995; Garamszegi et al. 2005). However, using data from 34 species, Garamszegi et al. (2005) found that residual testes size alone was not a significant predictor of the rate of extrapair paternity (Fig. 9.8) and suggested that testosterone production might mediate this relation. Using path analysis they compared statistically three models for the interaction between these two variables on the evolution of testes size, as follows. First, they suggested that extrapair paternity (EPP) and residual peak testosterone level (RPT) might affect each other and both directly influence the evolution of testis size. Second, they proposed that testis size might evolve first, under the influence of intense sperm competition, resulting directly in higher levels of EPP and a secondary, indirect effect on RPT. Finally, they suggested that testis size might evolve first under sperm competition, but then have a primary effect on RTP and a secondary one on EPP. Their analysis provided most support for the second model, explaining 28% of the variation in the data. Species that nest in colonies or at high density have also been suggested to experience higher levels of sperm competition than solitarily nesting species

Fig. 9.8 Rate of extrapair paternity in relation to relative testes size in 34 bird species. Rate of extrapair paternity is measured as the percent of offspring sired by extrapair males; relative testes size is measured as the residuals from the regression of combined testes mass on body mass, both log-transformed; model II regression is plotted to show trend. Data from Garamszegi, L. Z., Eens, M., Hurtrez-Bousses, S. and Møller, A. P. 2005. Hormones and Behaviour 47: 389-409, Table 1.

#! Reproductive Biology and Phylogeny of Birds (Gladstone 1979; Møller 1991; Birkhead and Møller 1992a). This may occur either because of the proximity of potential extrapair copulation partners or due to increased opportunities for multiple copulation partners in those species in which one member of the pair must stay and defend the nest site, thereby preventing males from guarding their mates. As expected, colonial species have significantly larger testes than solitarily nesting species (Møller 1991; Pitcher et al. 2005). Moreover, even within a species, there is evidence that the increased risk of sperm competition in colonial nesters can favor increased testis size. Brown and Brown (2003) found that Cliff swallow males nesting in larger colonies had testes masses almost three times that of males nesting in small colonies or solitarily. Although this pattern could be a facultative response to the perceived risk of sperm competition (i.e., males invest more in sperm production when nesting in big colonies), Brown and Brown (2003) suggested there may also be a genetic component to the difference as the preference for individuals to nest in different-sized colonies was heritable. Although testis size correlates with the intensity of sperm competition, an alternative hypothesis was proposed by Cartar (1985) based on the frequency of matings that a male obtains (the fertilization frequency hypothesis). He suggested that selection should favor larger testis size in species in which males pair and copulate with more than one female, simply because they require more sperm to fertilize the additional ova. Under this hypothesis, males in polygynous species are expected to have relatively larger testes than males in a monogamous species, even in the absence of sperm competition. In support of this hypothesis, Cartar (1985) found that polygynous waders (family Scolopacidae) had larger testes than monogamous species. It is now clear that sperm competition is widespread in birds (Birkhead and Møller 1992a; Birkhead 1998a,c): studies of mating behavior in a number of species have shown high levels of mate infidelity, and the use of DNA profiling techniques has demonstrated that extrapair paternity is common (Griffith et al. 2002). Thus, Cartar’s (1985) results were likely confounded by differences in the levels of sperm competition between socially monogamous and socially polygynous waders, though there are few data currently available to test this idea. Recently, in a large comparative study with data from 934 bird species, Pitcher et al. (2005) found that testis size increased with clutch size, suggesting that species with larger clutches actually require larger testes (Fig. 9.9). Large testes might be required to produce enough sperm to fertilize a greater number of ova and avoid sperm depletion during the fertilization period. The fitness costs of sperm depletion could thus favor increased testis size not only in species in which males require more sperm to fertilize a larger number of females (as originally proposed by Cartar 1985), but also in situations where each female has more eggs to fertilize. In addition, sperm depletion has been suggested to play a role favoring increased testis size in species in which breeding is restricted to a few days or weeks, such as at high latitudes (Kenagy

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Fig. 9.9 Relation between residual testes mass (RTM; calculated as in Fig. 9.7) and clutch size (CS) in 934 bird species. Model II regression line is shown (RTM = –0.81 + 0.22 CS, r2 = 0.02, P <0.0001). Redrawn from data in Pitcher, T. E., Dunn, P. O. and Whittingham, L. A. 2005. Journal of Evolutionary Biology 18: 557-567, Fig. 4c and appendix.

and Trombulak 1986). The increase in testis size with latitude found in some species (Pitcher and Stutchbury 1998; Merilä and Sheldon 1999) is consistent with this idea, but further work is required to tease apart the influences of variation in the intensity of sperm competition with both shortened breeding seasons and geographic range. Moreover, the relation between clutch size and relative testis size is very weak, with clutch size explaining only about 1% of the variation in testis size. This may suggest that some confounding variable is actually responsible for the reported relation (Fig. 9.9).

9.2.9

Costs of Large Testis Size

Despite the seeming benefits of increased testis size when sperm competition is intense, it is not clear what ultimately limits testis size in birds. In other words, are there costs to larger testes and can these costs explain some of the variation in testis size across species? Studies on the costs associated with relatively large testes have lagged far behind those on the proposed benefits, but a few hypotheses have been suggested. For example, Zuk et al. (1990) proposed that increased testis size might increase the risk of parasitemia brought on by the immunosuppressive effects of higher androgen levels (see also Folstad and Karter 1992; Salvador et al. 1996; Merilä and Sheldon 1999). However, it is not clear whether increased testis size necessarily brings about an equivalent increase in androgen production (Weatherhead et al. 1993; Wikelski et al. 2003). In theory it should be possible to increase spermatogenic

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Reproductive Biology and Phylogeny of Birds

tissue without increasing endocrine tissue but, across species, larger testes generally result in higher circulating levels of testosterone (Wingfield and Moore 1987; Garamszegi et al. 2005). Brown and Brown (2003) were able to increase testis size experimentally in Cliff swallows by applying insecticidal powder to their nests, thereby reducing their ectoparasite burden. One interpretation of this result is that exposure to parasites elevates levels of plasma corticosterone, which in turn alters the adrenal response to stress, resulting in both a suppression of testosterone and a reduced testes mass (Dunlap and Schall 1995). It is also possible that a reduced parasite load frees up resources for investment in testis tissue that otherwise would be needed for immune responses. Larger testes might also be costly energetically. This could include costs of production and maintenance, as well as the increased flight costs that result from carrying more mass. In most species, the mass of both testes combined is only 1-3% of body mass, so would seem unlikely to result in an appreciable increase in flight costs. However, in species subject to intense sperm competition, with relatively large testes as a result, testes mass can range from 4-8% of body mass (Briskie 1993). Studies that have experimentally attached weights to birds have found significant effects of the extra burden at levels similar to this testes mass (e.g., Wright and Cuthill 1989). The fact that younger males have smaller testes than older males, and that the testes of males of all ages typically undergo a period of regression in the non-breeding season, suggests there is a cost to maintaining enlarged testes. Perhaps studies that experimentally induce earlier or larger recrudescence with gonadstimulating hormones (and thereby result in a male with a larger pair of testes than normal) can be used to examine the presumed costs of large testis size.

9.3 9.3.1

EVOLUTION OF AVIAN SPERM The Avian Spermatozoon

The avian spermatozoon has two main structural parts, the head and the tail. The sperm head consists of an anterior acrosome and a posterior nucleus, while the tail consists of an anterior midpiece (plesiomorphically just posterior to the nucleus) that surrounds an axoneme comprised of an anterior principal piece and a posterior endpiece. In the parvorder Passerida, the midpiece mitochondria typically extend for most of the length of the tail (Chapter 8). Extensive variation in the structure of sperm among different species of birds was described by early workers (Ballowitz 1888; Retzius 1909, 1911), and more recent studies have particularly uncovered extensive interspecific variation in the lengths of sperm and their component parts (McFarlane 1963; 1971; Briskie and Montgomerie 1992; Koehler 1995). The ultrastructure of avian sperm has been described across a wide range of species and is reviewed comprehensively in Chapter 8. The greatest differences in avian sperm morphology are found when comparing passerine (order Passeriformes) and non-passerine birds (all

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remaining extant orders). The spermatozoa of non-passerines are similar to some reptilian sperm and are linear (or slightly undulating) in the head region, while the sperm of passerine birds are coiled in a helical fashion (McFarlane 1963, 1971; Chapter 8). Passerine sperm can vary among species from being helical only in the acrosomal region to having a helical acrosome, nucleus and midpiece (Koehler 1995). At the ultrastructural level, passerine sperm lack the perforatorium, endonuclear canals, distinct centrioles, and annulus found in non-passerines, while they possess a series of nine accessory fibers that is absent or less developed in non-passerines (Asa and Phillips 1987; Koehler 1995). Passerine sperm also have an ‘undulating membrane’ that arises from the apex of the acrosome and is said to extend for most of the length of the sperm (Humphreys 1972; Henley et al. 1978; but see Chapter 8). Species in the orders Charadriiformes and Procellariformes also possess a helical sperm membrane but this is thought to have evolved independently of the similar membrane found in passerines (Koehler 1995). The only exception to the distinctive nature of passerine sperm was recently reported by Birkhead et al. (2006) in the Eurasian bullfinch (Pyrrhula pyrrhula). Unlike other passerines, the sperm of this species has a rounded head, a tiny midpiece, and no obvious, external helical structure. It does, however, retain the passerine hallmark of a microtubular helix (Birkhead, Giusti, Immler and Jamieson, pers. comm.). This sperm structure may be unique to this species, as the congeneric Beavan’s bullfinch (P. erythaca) has sperm closer to that of other passerines (Birkhead et al. 2006). As sperm morphology has been examined microscopically in only about 100 bird species (Chapter 8), it is too early to say if the Eurasian bullfinch is the only exception to the generally conservative structure of passerine sperm.

9.3.2

Variation in Sperm Size

Sperm length is highly variable in birds (McFarlane 1963, 1971; Allen et al. 1968; Humphreys 1972; Briskie and Montgomerie 1992; Briskie et al. 1997; Johnson and Briskie 1999). Total sperm length varies six-fold among passerines species that have been examined, from 42.7 µm in the Red-backed shrike (Briskie et al. 1997) to 292.4 µm in the Reed bunting (Emberiza schoeniclus; Dixon and Birkhead 1997). Among waders (families Scolopacidae, Charadriidae and Jacanidae), sperm length varies from 57.0 µm in the Semipalmated plover (Charadrius semipalmatus) to 133.2 µm in the Ruff (Philomachus pugnax; Johnson and Briskie 1999). Even within a family, sperm length can be quite variable. Allen et al. (1968) measured the sperm lengths of 9 species (and 4 subspecies) of passerines in the family Icteridae (New World blackbirds) and found that sperm length varied from 77.9 µm in the Western meadowlark (Sturnella neglecta) to 148.6 µm in eastern populations of the Red-winged blackbird (A. p. phoeniceus). Although published information on sperm length is available for only about 120 bird species, there are no striking differences between the sperm

#!" Reproductive Biology and Phylogeny of Birds lengths of passerines (maximum 292.4 µm in the Reed bunting) and nonpasserines (maximum ~230 µm in the Japanese quail, Coturnix japonica; Woolley 1995). However, more information on a wider variety of species is needed to determine how sperm size varies with taxonomic affiliation and to what degree such phylogenetic effects might constrain evolutionary changes in sperm size. Across species, the relative size of each region of the spermatozoon can also vary. For example, the size of the midpiece varies from only 3.3 µm in the Great crested flycatcher (Myiarchus crinitus; Koehler 1995) to 161.4 µm in the Japanese quail; Woolley 1995). Among 17 species of passerines studied by Briskie and Montgomerie (1992), tail length varied from 36 µm in the Least flycatcher (Empidonax minimus) to 262 µm in the Yellow warbler. In this sample of birds, differences in the length of the tail explained more than 99% of the variation in total sperm length (Briskie and Montgomerie 1992). In contrast, the size of the nucleus among passerines varies only from 2.5 µm in the American robin (Turdus migratorius) to 18.5 µm in the Great crested flycatcher (Koehler 1995), and the size of the acrosome only from 2.5 µm in the European starling to 13.5 µm in the Violet-green swallow (Tachycineta thalassina; Koehler 1995). Even though there is relatively little interspecific variation in the size of the nucleus and acrosome, the lengths of the various components of the spermatozoon appear to be correlated across species (Johnson and Briskie 1999). For example, Birkhead et al. (2005) found that flagellum length was positively correlated with midpiece length in a sample of 29 passerine species. As the midpiece extends for most of the length of the flagellum in most passerines, this result is perhaps not unexpected (see Chapter 8), though it does suggest that genetic correlation plays a role in sperm design (Birkhead et al. 2005). Most comparative studies of sperm size variation have assumed that there is little variation within a species. Although interspecific variation is usually much larger than variation within a species (Briskie and Montgomerie 1992; Briskie et al. 1997), recent work has revealed that variation in sperm length (and the size of each component part) can also vary significantly among males of a single species (Tuttle et al. 1996; Morrow and Gage 2001a; Birkhead et al. 2005). Thus, in the Zebra finch, variation in sperm flagellum length and midpiece length between males was even greater than variation in an external morphological trait such as tarsus length (Birkhead et al. 2005). Total sperm length in this species varied from 45 to 78 µm (Birkhead et al. 2005) but, within an individual male, sperm length was highly consistent, both within an ejaculate and over time (Birkhead and Fletcher 1995). Intraspecific variation in the size of sperm and its component parts are heritable and negatively correlated in Zebra finches (Birkhead et al. 2005), confirming that this variation is not simply due to environmental effects.

9.3.3

Sexual Selection and Sperm Morphology

To date, understanding variation in sperm size between species has focused mostly on identifying the likely benefits to increased sperm length as an

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adaptation to variation in the female reproductive tract. As all birds have internal fertilization, it is the female’s reproductive tract that forms the arena in which sperm must function and in which selection will favor changes in sperm size. Two factors that have attracted the most attention in this regard are the relations between sperm length and the forces of sperm competition (Birkhead and Møller 1992a) and the effects of cryptic female choice (Eberhard 1996). Given a limited budget for expenditure on gamete production, males subject to intense sperm competition are expected to increase the number of sperm produced (i.e., ejaculate size, sperm concentration, copulation frequency) at the expense of sperm size (Parker 1982). In other words, males face a trade-off between the number and size of sperm, and sperm competition is expected to favor males that produce large numbers of small sperm in order to dilute or displace the ejaculates of rival males. However, this expected pattern has generally not been supported in birds and most other groups of animals (Gomendio and Roldan 1991; Briskie and Montgomerie 1992; Gage 1994; Briskie et al. 1997; Johnson and Briskie 1999). Instead, bird species subject to intense levels of sperm competition generally have longer sperm (Fig. 9.10; Briskie et al. 1997). Species with large testes also generally produce longer sperm (Johnson and Briskie 1999), as would be expected if testis size is a reasonable index of the intensity of sperm competition, as outlined earlier in this chapter. This suggests that males do not directly trade-off sperm size for sperm number but instead invest proportionately more into gamete

Fig. 9.10 Sperm length in relation to the rate of extrapair paternity in 21 species of passerines. Model II regression line is shown (sperm length = 52.4 + 1.74 EPP rate, r2 = 0.31, P = 0.01). The relation remains significant when controlling for phylogeny. Modified after Briskie, J. V., Montgomerie, R. and Birkhead, T. R. 1997. Evolution 51: 937-945, Fig. 2.

#!$ Reproductive Biology and Phylogeny of Birds production (via increased testis and sperm size) in species subject to more intense sperm competition. Early studies of sperm size variation proposed that increased sperm length was an adaptation that resulted in faster swimming sperm (Gomendio and Roldan 1991; Briskie and Montgomerie 1992). Among a small sample of domesticated mammal species, Gomendio and Roldan (1991) found a positive relation between sperm length and swimming speed, and they suggested that increased levels of sperm competition would favor increased swimming speeds in the race to fertilize a female’s ova. Thus longer flagella would make sperm swim faster and generate more thrust (Katz et al. 1989; Katz and Drobnis 1990). However, birds differ from mammals in that sperm are typically stored by the female in specialized sperm storage tubules (SSTs) before they move up the oviduct to fertilize an ovum in the infundibulum (Hatch 1983; Shugart 1988; Briskie and Montgomerie 1993; Chapters 6 and 10). This led Briskie and Montgomerie (1992) to propose that increased sperm length in birds might be an adaptation to increase swimming speed in order to reach SSTs before the sperm of rival males. Indeed, wader species with more promiscuous mating systems had longer midpieces (the site of energy stores) than species with more monogamous mating systems, suggesting that increased resources were devoted to locomotion (Johnson and Briskie 1999). Birkhead et al. (2005), however, found no relation between sperm length and sperm velocity among a sample of 105 Zebra finch males with flagellum lengths varying from about 37 to 68 µm, and midpieces varying from 15 to 48 µm. Measures of sperm swimming speed across a wide range of species are needed to resolve this issue but it seems unlikely that increased sperm length is simply a function of the benefits accrued by greater swimming speed. The storage of sperm in a female’s SSTs for days or even weeks before fertilization (Birkhead and Møller 1992b) suggests that variation in sperm length might be related to variation in sperm storage. Both the number and size of SSTs vary widely across bird species, and sperm length is positively correlated with SST length (Fig. 9.11A; Briskie and Montgomerie 1992, 1993; Briskie et al. 1997). In most species studied to date, the length of a single SST is about twice that of the sperm that they store (Fig. 9.11A). A few species have sperm only slightly shorter than an SST (e.g., Red-eyed vireo, Vireo olivaceus), while others have SSTs that are more than 3 times as long as the sperm (e.g., American robin; Briskie and Montgomerie 1993). A similar correspondence between the size of sperm and the structures that females use to store sperm has also been observed in other taxa (e.g., Dybas and Dybas 1981; Presgraves et al. 1999). Variation among species in the ratio of SST length to sperm length, and the fact that SSTs are usually about twice as long as a single sperm, suggests that SSTs have not simply evolved to store sperm (Briskie and Montgomerie 1992). In some species, sperm appear to form layers within each SST, and it has been suggested this might be a mechanism by which females could control paternity through the differential storage of sperm from different males

Testis Size, Sperm Size and Sperm Competition

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Fig. 9.11 Sperm length in relation to A length and B number of sperm storage tubules (SSTs) in females of 20 passerine bird species. Solid lines are model II regressions; dashed line in (A) indicates SSTs that are twice as long as a single sperm. Both relations are significant with and without controlling for phylogeny. Modified after Briskie, J. V. and Montgomerie, R. 1992. Proceedings of the Royal Society of London B 247: 89-95, Fig. 4a, b.

(Compton et al. 1978; Briskie and Montgomerie 1993). More recent work suggests that such sperm stratification is unlikely to play a role in control of paternity (Birkhead 1998a,c). Nonetheless, differences between species in the lengths of their sperm and the lengths of SSTs (Fig. 9.11A) suggest that the way in which sperm are used by females may vary interspecifically. Using species in which genetic profiling techniques were used to estimate levels of promiscuity, Briskie et al. (1997) found that sperm length was correlated

#!& Reproductive Biology and Phylogeny of Birds positively with levels of extrapair paternity (Fig. 9.10). However, a path analysis revealed that the relation between sperm length and extrapair paternity arose only indirectly through the positive relation between the length of the SSTs and extrapair paternity (Briskie et al. 1997). In other words, SST length may evolve in response to mating with multiple males, and males in turn, respond by evolving longer sperm. Why increased length of SSTs would be advantageous to females with promiscuous mating systems is not clear. One possibility is that increased SST size incites competition between the sperm of rival males, and thus selects for males that produce sperm that are successful competitors (Keller and Reeve 1995). This might then select for males that have longer sperm, if such sperm are better are competing for access to SSTs, or can displace rival sperm from SSTs. Like SST size, SST number also varies across species, ranging from 325 per female in the Blackpoll warbler (Dendroica striata; Briskie 1996) to over 24,000 per female in the Domestic turkey (Meleagris gallopavo; Goodrich-Smith and Marquez 1978). In an early study of SST variation, Briskie and Montgomerie (1992) found that sperm length was inversely correlated with SST number (Fig. 9.11B). Such a pattern was explained by long sperm being advantageous, perhaps via increased swimming speed, in those species in which a lower number of SSTs increased competition for access to limited storage sites (Briskie and Montgomerie 1992). However, subsequent work failed to confirm a link between sperm length and SST number, and it is now thought this relation is due to an inverse relation between SST number and SST size, which instead reflects the necessity of decreasing SST number to accommodate larger SSTs in the limited size of the uterovaginal junction where SSTs reside (Briskie et al. 1997). An alternative view to long sperm evolving as an adaptation to sperm competition was suggested by Eberhard (1996), who proposed that postcopulatory female choice (or ‘cryptic female choice’) could explain the elaborate structures associated with mating, including sperm morphology. Under this scenario, females may prefer males with long sperm for many of the same reasons they might choose males with bright plumage or long tails feathers during the precopulatory phases of mate choice (Keller and Reeve 1995). Whether such cryptic mate choice occurs, and whether it might lead to directional selection on sperm length in birds is still unresolved, although there is evidence that female domestic fowl can preferentially eject sperm from subordinate sires (Pizzari and Birkhead 2000). Although there is no evidence at present that female birds selectively favor longer sperm, Briskie et al. (1997) suggested that by increasing the length of their SSTs, females might increase the probability that the sperm from two or more males mix in the SSTs. This would incite greater sperm competition and indirectly favor longer sperm if increased sperm length is advantageous in situations of sperm competition (see above). Studying the processes of differential sperm use in birds poses technical problems that will not be easy to overcome, nor will it be easy to separate the

Testis Size, Sperm Size and Sperm Competition

#!'

effects of mate choice from sperm competition (Birkhead 1998b). At present it is not possible to determine whether the correlation observed between increased sperm length and high levels of promiscuity among different species of birds is the result of sperm competition or cryptic female choice (or both). However, the patterns observed to date do suggest that sexual selection plays a key role in shaping sperm morphology in birds. The challenge is now to determine how changes in sperm morphology, such as increased length, have evolved to overcome both the barriers they encounter in the female reproductive tract and the competition they face from sperm of rival males.

9.3.4

Costs of Large Sperm Size

Compared to the production of a single egg, the costs (energetic or otherwise) of producing a single spermatozoon would appear trivial. This has fostered the view that sperm are ‘cheap’ relative to the costs of producing eggs. Of course, males do not produce a single sperm for each ovum produced by a female, and it is now clear that sperm production can limit ejaculate size and, ultimately, male reproductive success (Nakatsuru and Kramer 1982; Birkhead 1991). If sperm are sometimes limiting for males, then long sperm may be more costly to produce than short sperm. Thus the differences between costs and benefits of long sperm should vary within and among species in ways that might explain the extensive variation in the sperm length of birds. One obvious cost of increased sperm length is the greater energetic and material costs of producing a larger structure. The energetic costs to birds of producing sperm are unknown, but it is logical that larger sperm must be more costly to produce. Birkhead et al. (1998) found that male Zebra finches on experimentally reduced diets developed significantly smaller testes and produced fewer sperm, but did not produce smaller sperm than control males. However, spermatozoa in this species are relatively short, and further tests of this type are needed to determine if sperm size is limited by resource availability in species with long sperm. Indeed, sperm size might not be influenced much by environment, requiring selection experiments to examine the influence of resource availability on the evolution of sperm size (e.g., Morrow and Gage 2001b). Long sperm may also be costly if they have reduced longevity due the increased demands on energy stores when propelling a long flagellum (Stockley et al. 1997). Fish species with longer sperm have reduced periods of swimming activity of the sperm, suggesting just such a trade-off between sperm size and sperm longevity (Stockley et al. 1997), and therefore a potential cost to production of long sperm. Although we do know that sperm can be stored in a female’s SSTs for periods up to several weeks (Birkhead and Møller 1992b), the factors that influence the duration of sperm storage (and thus sperm longevity) in birds have not yet been studied. If increased sperm length in birds reduces their longevity then an inverse relation between the duration of sperm storage and sperm size would be expected.

#" Reproductive Biology and Phylogeny of Birds

9.3.5

Other Explanations for Diversity in Sperm Morphology

At present, variation in sperm size in birds seems likely to be a product of sexual selection. However, few studies of sperm size evolution in birds have been undertaken, and it is possible that other factors may explain some of the variation in sperm morphology. For example, sperm size in butterflies increases with body size across species (Gage 1994) and it is possible that some variation in sperm length in birds could be explained by a similar allometric effect. No significant relation between sperm length and body size was found among waders (Johnson and Briskie 1999), but sperm length is negatively related to body size among passerines (Fig. 9.12; Briskie and Montgomerie 1992). Both of these analyses were based on small sample sizes so further work is needed before the influence of allometry on sperm length can be determined. Sperm length might respond to selection on female gametes. If gamete size is determined by a similar proximate process in both sexes, then selection for increased ovum size might result in a correlated response in sperm size (Halliday and Arnold 1987). Johnson and Briskie (1999) examined this linkage disequilibrium hypothesis in a comparative study of sperm size in waders, but found no significant relation between egg volume and sperm length. There was also no relation between sperm length and egg size in a comparative study of passerines (Briskie and Montgomerie 1992). Sperm size might also be influenced by genome size. As genome size varies across species, those species with larger (and presumably heavier) genomes

Fig. 9.12 Relation between sperm length (SL) and body mass (BM) in 20 passerine bird species (both variables log-transformed). Model II regression is shown (logSL = 2.4 –1.1 logBM, r2 = 0.19, P = 0.05). Modified after Briskie, J. V. and Montgomerie, R. 1992. Proceedings of the Royal Society of London B 247: 89-95, Fig. 4a, b.

Testis Size, Sperm Size and Sperm Competition

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might require longer tails and midpieces to carry the greater load. For example, the size of the enucleated red blood cells in birds is known to be positively related to genome size (Gregory 2001), suggesting that a similar pattern might be found with respect to sperm size. Gage (1998) tested this idea across a wide range of mammal species but found no significant relation between sperm length and nucleus size. Though genome size has not been compared to sperm size in birds, genome size varies only two-fold among bird species (Gregory 2001) and thus seems unlikely to explain the more than 6-fold variation in sperm length (Briskie and Montgomerie 1992). Most modern comparative studies have used statistical methods to control for phylogenetic effects (Harvey and Pagel 1991) on the assumption that at least some of the observed variation in sperm morphology may be due to phylogeny and may thus not be an adaptation to current selective pressures. In other words, two species may have similar sperm length as a consequence of inheriting this trait from a common ancestor. Although phylogeny clearly plays a major role in determining some of the variation in sperm structure (e.g., compare passerine and non-passerine sperm; McFarlane 1963; Birkhead et al. 2006; Chapter 8), sperm length varies so much between even closely related species that phylogeny does not appear to have much influence on sperm size. Artificial selection experiments in insects confirm the evolvability of sperm length (Morrow and Gage 2001b) and sperm length in birds has been shown to be heritable (Birkhead et al. 2005), so there is clearly potential for selection to favor changes in sperm size in response to some of the factors described above. However, the degree to which phylogeny limits adaptive changes in sperm length are poorly understood and there is a need for a broad scale comparative study of sperm morphology to assess the degree to which sperm size in birds depends on their phylogenetic history.

9.4

SCOPE FOR FUTURE RESEARCH

Despite a long and productive tradition among anatomists, physiologists and histologists, the study of avian reproductive morphology has only recently attracted the attention of evolutionary biologists and field-based ornithologists. Much of the work done to date has been highly descriptive and based on correlation analyses, rather than focussing on experiments explicitly designed to distinguish among potential causes and control confounding variables. Comparative analyses of testis and sperm morphology, in particular, have mainly been limited to the analysis of variation in size. Little of the intricate detail of sperm morphology and variation among species revealed by histologists has been examined from an evolutionary perspective (but see Chapter 8). Much could be learnt from the application of evolutionary theory to the ultrastructure of the reproductive systems of birds. Mitochondrial size and number, and tail structure, for example, are known to differ among bird species (e.g., Henley et al. 1978; Soley 1993) and it would be interesting to determine whether variation in the intensity of sperm competition can explain

#"

Reproductive Biology and Phylogeny of Birds

this variation. Intense sperm competition appears to have led to convergent evolution among unrelated species in relative testis size and absolute sperm length, but whether similar patterns will be found at a finer structural level remains to be seen. The patterns of testis size variation and sperm morphology reviewed here might give the impression that there is little new to learn by examining and studying the reproductive anatomy of additional species. However, sperm size in birds has been measured in only about 120 species worldwide, and the sperm of relatively few species has been examined in finer detail at the microscopic level (Chapter 8). Moreover, most of the species whose sperm size is known are North American and European birds, and almost nothing is known about variation in sperm morphology in the majority of avian families (Chapter 8). Even among common bird species, we lack basic information. Recently, for example, Birkhead et al. (2006) discovered that the structure of Eurasian bullfinch sperm is unlike that of any other passerine, though the reasons for this remain obscure. One can only wonder what sort of variation might be found in the >9000 other species of birds whose sperm have yet to be examined. The collection of sperm from birds is easy and non-destructive, and can be mastered with a little practise (Wolfson 1952; Immler and Birkhead 2005). Thus, systematic surveys of sperm morphology can be undertaken during field studies of birds and we encourage field ornithologists to consider semen collection as one of their routine procedures. More data on interspecific variation in testis size is available than on sperm size, but there is still a strong bias to species in temperate areas of the northern hemisphere. For a small set of species, testis mass has been measured directly, but most of the available data on testis mass has been taken from labels of museum skins, in which length (and sometimes width) of one or both testes were measured by the collector (Pitcher et al. 2005). These measurements are then used to estimate testes mass, assuming that testis shape, density, and asymmetry is the same for all species (Pitcher et al. 2005; but see Kempenaers et al. 2002). The collection of birds from the wild has long gone out of favor, but non-destructive techniques are available to measure testis dimensions (e.g., laparotomy; Risser 1971), and recent developments in the use of ultrasound and magnetic resonance imaging (Dietz et al. 1999; Czisch et al. 2001) may provide less invasive ways of estimating testis size. Where permitted by law, birds salvaged from road kills, building collisions, and extreme weather events can also provide considerable information on testis mass and asymmetry without the need for destructive sampling (e.g., Brown and Brown 2003). The development of sperm competition theory in the 1970s provided a sound framework (Parker 1970) that later workers used to guide their studies. There is no doubt that this approach has been fruitful and that sperm competition is likely the major factor (aside from body size) that explains much of the variation in testis size and sperm size in birds (Birkhead 1998a, c).

Testis Size, Sperm Size and Sperm Competition

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However, it is too early to rule out other potential explanations for this variation, and it is perhaps simplistic to conclude that variation in the intensity of sperm competition alone can account for all of the variation in testis and sperm size. Recent studies that have found high levels of promiscuity in species with both small testes (e.g., Moustached warbler, Acrocephalus melanopogon; Schulze-Hagen et al. 1995; Blomqvist et al. 2005) and short sperm (e.g., Malurus fairy-wrens; Tuttle and Pruett-Jones 2004), provide striking anomalies that do not fit the patterns expected from broad-scale comparisons with respect to sperm competition. These examples suggest that we are missing the full picture. Likewise, the reasons for testicular asymmetry, and why it varies with age, are almost a complete mystery. More data, not only on male reproductive morphology in a greater variety of species, but also on factors that are likely to be important (e.g., mating frequency, levels of infidelity, breeding season duration, patterns of sperm storage by females, sperm longevity, etc.), are now needed to separate the causal factors and determine how they interact. Such data will also provide a firm basis for future work as the field develops from one of description and correlation, to one in which controlled experiments are used to test the robustness of proposed causal pathways and elucidate the mechanisms responsible for the observed patterns.

9.5

ACKNOWLEDGMENTS

We are particularly grateful to Meghan Goodchild, Kristen Scott, Jason Clarke and Christina Cliffe for help with library work; to Trevor Pitcher for supplying us with his data on testis size; to Tim Birkhead and Barrie Jamieson for useful insights; and to the University of Canterbury (to J. V. B.) and the Natural Sciences and Engineering Research Council of Canada (to R. M.) for supporting our research on avian reproduction. During manuscript preparation R. M. was supported by a Killam Research Fellowship, and the Queen’s University Research Chairs program.

9.6

LITERATURE CITED

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#"$ Reproductive Biology and Phylogeny of Birds Dybas, L. K. and Dybas, H. S. 1981. Coadaptation and taxonomic differentiation of sperm and spermathecae in featherwing beetles. Evolution 35: 168-174. Eberhard, W. G. 1996. Female Control: Sexual Selection by Cryptic Female Choice. Princeton University Press, Princeton, NJ. 501 pp. Erpino, M. J. 1969. Seasonal cycle of reproductive physiology in the Black-billed magpie. Condor 71: 267–279. Evans, M. R. and Goldsmith, A. R. 2000. Male wrens with large testes breed early. Animal Behaviour 60: 101-105. Ferris, H. B. 1933. Triple testes in a Snowy plover. Condor 35: 124. Follett, B. K. 1984. Birds. Pp. 283-350. In G. E. Lamming (ed.), Marshall’s Physiology of Reproduction. Vol. 1. Reproductive Cycles of Vertebrates. Churchill Livingstone, London. Folstad, I. and Karter, A. J. 1992. Parasites, bright males, and the immunocompetence handicap. American Naturalist 139: 603-622. Foster, M. S. 1987. Delayed maturation, neoteny, and social system differences in two manakins of the genus Chiroxiphia. Evolution 41: 547-558. Freeman, S. 1990. The evolution of the scrotum: a new hypothesis. Journal of Theoretical Biology 145: 429-445. Friedmann, H. 1927. Testicular asymmetry and sex ratio in birds. Biological Bulletin of Woods Hole 52: 197-207. Gage, M. J. G. 1994. Associations between body size, mating pattern, testis size and sperm lengths across butterflies. Proceedings of the Royal Society of London B 258: 247-254. Gage, M. J. G. 1998. Mammalian sperm morphometry. Proceedings of the Royal Society of London B 265: 97-103. Garamszegi, L. Z., Eens, M., Hurtrez-Bousses, S. and Møller, A. P. 2005. Testosterone, testes size, and mating success in birds: a comparative study. Hormones and Behaviour 47: 389-409. Gladstone, D. E. 1979. Promiscuity in monogamous colonial birds. American Naturalist 114: 545-557. Gomendio, M. and Roldan, E. R. S. 1991. Sperm competition influences sperm size in mammals. Proceedings of the Royal Society of London B 243: 181-185. Goodrich-Smith, M. A. and Marquez, B. J. 1978. Estimation of the numbers of sperm storage tubules located at the uterovaginal junction in Turkey oviduct. Poultry Science 57: 1139. Graves, G. R. 2004. Testicular volume and asymmetry are age-dependent in Blackthroated blue warblers (Dendroica caerulescens). Auk 121: 473-485. Gregory, T. R. 2001. The bigger the C-value, the larger the cell: genome size and red blood cell size in vertebrates. Blood Cells, Molecules, and Diseases 27: 830-843. Griffith, S. C., Owens, I. P. F. and Thuman, K. A. 2002. Extra pair paternity in birds: a review of interspecific variation and adaptive function. Molecular Ecology 11: 2195-2212. Halliday, T. and Arnold, S. J. 1987. Multiple mating by females: a perspective from quantitative genetics. Animal Behaviour 35: 939-941. Harcourt, A. H., Harvey, P. H., Larson, S. G. and Short, R. V. 1981. Testis weight, body weight and breeding system in primates. Nature 293: 55-57. Harvey, P. H. and Pagel, M. 1991. The Comparative Method in Evolutionary Biology. Oxford University Press, Oxford. 438 pp.

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Hatch, S. A. 1983. Mechanism and ecological significance of sperm storage in the Northern fulmar with reference to its occurrence in other birds. Auk 100: 593-600. Hegner, R. E. and Wingfield, J. C. 1990. Annual cyccle of gonad size, reproductive hormones, and breeding activity of free-living House sparrows (Passer domesticus (L.)) in rural New York. Pp. 123-135. In J. Pinowski and D. and Summers-Smith (eds), Granivorous Birds in the Agricultural Landscape. INTECOL, Warsaw. Henley, C., Feduccia, A. and Costello, D. P. 1978. Oscine spermatozoa: a light- and electron-microscopy study. Condor 80: 41-48. Hill, G. E. 1994. Testis mass and subadult plumage in Black-headed grosbeaks. Condor 96: 626-630. Hocking, P. M. 1992. Bilateral testicular asymmetry and supernumerary testes in the Domestic fowl (Gallus domesticus). British Poultry Science 33: 455-460. Humphreys, P. N. 1972. Brief observations on the semen and spermatozoa of certain passerine and non-passerine birds. Journal of Reproduction and Fertility 29: 327336. Immelmann, K. 1971. Ecological aspects of periodic reproduction. Avian Biology 1: 341-389. Immler, S. and Birkhead, T. R. 2005. A non-invasive method for obtaining spermatozoa from birds. Ibis 147: 827-830. Johnson, D. D. P. and Briskie, J. V. 1999. Sperm competition and sperm length in shorebirds. Condor 101: 848-854. Katz, D. F. and Drobnis, E. Z. 1990. Analysis and interpretation of the forces generated by spermatozoa. Pp. 125-137. In B. D. Bavister, J. Cummins, and E. R. S. Roldan (eds), Fertilization in Mammals. Serono Symposia, Norwell, Massachusetts. Katz, D. F., Drobnis, E. Z. and Overstreet, J. W. 1989. Factors regulating mammalian sperm migration through the female reproductive tract and oocyte vestments. Gamete Research 22: 443-469. Keller, L. and Reeve, H. K. 1995. Why do females mate with multiple males? The sexually selected sperm hypothesis. Advances in the Study of Behavior 25: 291315. Kenagy, G. J. and Trombulak, S. C. 1986. Size and function of mammalian testes in relation to body size. Journal of Mammalogy 67: 1-22. Kempenaers, B., Peer, K., Vermeirssen, E. L. M. and Robertson, R. J. 2002. Testes size and asymmetry in the Tree swallow Tachycineta bicolor: a test of the compensation hypothesis. Avian Science 2: 115-122. Kimball, R. T., Ligon, J. D. and Merola-Zwartjes, M. 1997. Testicular asymmetry and secondary sexual characters in Red junglefowl. Auk 114: 221-228. Koehler, L. D. 1995. Diversity of avian spermatozoa ultrastructure with emphasis on the members of the order Passeriformes. In B. G. M. Jamieson, J. Ausio and J-L. Justin (eds), Advances in Spermatozoal Phylogeny and Taxonomy. Mémoires du Muséum National D’Histoire Naturelle 166: 437-444. Lake, P. E. 1981. Male genital organs. Pp. 1-61. In A. S. King and J. McLelland (eds), Form and Function in Birds. Academic Press, London. Lal, P. and Thapliyal, J. P. 1982. Thyroid-gonad and thyroid-body weight relationship in the Red-vented bulbul, Molpastes cafer. General and Comparative Endocrinology 48: 98-103. Lasiewski, R. C. and Dawson, W. R. 1967. A re-examination of the relation between standard metabolic rate and body weight in birds. Condor 69: 13-23.

#"& Reproductive Biology and Phylogeny of Birds Leeuwenhoek, A. van. 1697. Continuatio Arcanorum Naturae Detectorum. H. Kroonevelt, Delphis Batavorum, 568 pp. Ligon, J. D. 1997. A single functional testis as a unique proximate mechanism promoting sex-role reversal in coucals. Auk 114: 800-801. Lofts, B. and Murton, R. K. 1968. Photoperiodic and physiological adaptations regulating avian breeding cycles and their ecological significance. Journal of Zoology, London 155: 327-394. Lofts, B. and Murton, R. K. 1973. Reproduction in birds. Pp. 1-107. In D. S. Farner, J. R King and K. C. Parkes (eds), Avian Biology, volume 3. Academic Press, New York. Marshall, A. J. 1961. Reproduction. Pp 169-213. In A. J. Marshall (ed.), Biology and Comparative Physiology of Birds, Vol. II. Academic Press, New York. Merilä, J. and Sheldon, B. C. 1999. Testis size variation in the Greenfinch Carduelis chloris: relevance for some recent models of sexual selection. Behavioral Ecology and Sociobiology 45: 115-123. McFarlane, R. W. 1963. The taxonomic significance of avian sperm. Proceedings of the 13th International Ornithological Congress: 91-102. McFarlane, R. W. 1971. The ultrastructure and phylogenetic significance of avian spermatozoa. Ph.D. thesis, University of Florida, USA. McNab, B. K. 1966. An analysis of body temperature of birds. Condor 68: 47-55. Møller, A. P. 1988. Testes size, ejaculate quality and sperm competition in birds. Biological Journal of the Linnean Society 33: 273-283. Møller, A. P. 1989. Ejaculate quality, testes size and sperm production in mammals. Functional Ecology 3: 91-96. Møller, A. P. 1991. Sperm competition, sperm depletion, paternal care, and relative testis size in birds. American Naturalist 137: 882-906. Møller, A. P. 1994. Directional selection on directional asymmetry: testes size and secondary sexual characters in birds. Proceedings of the Royal Society of London B 258: 147-151. Møller, A. P. and Briskie, J. V. 1995. Extra-pair paternity, sperm competition and the evolution of testis size in birds. Behavioral Ecology and Sociobiology 36: 357-365. Møller, A. P. and Erritzøe, J. 1988. Badge, body and testes size in House sparrows Passer domesticus. Ornis Scandivanica 19: 72-73. Moore, C. R. 1926. The biology of the mammalian testis and scrotum. Quarterly Review of Biology 1: 4-50. Morrow, E. H. and Gage, M. J. G. 2001a. Sperm competition experiments between lines of crickets producing different sperm lengths. Proceedings of the Royal Society of London B 268: 2281-2286. Morrow, E. H. and Gage, M. J. G. 2001b. Artificial selection and heritability of sperm length in Gryllus bimaculatus. Heredity 87: 356-362. Mulder, R. A. and Cockburn, A. 1993. Sperm competition and the reproductive anatomy of male Superb fairy-wrens. Auk 110: 588-593. Murton, R. K. and Westwood, N. J. 1977. Avian Breeding Cycles. Oxford University Press, Oxford. Nakamura, M. 1990. Cloacal protuberance and copulatory behavior of the Alpine accentor (Prunella collaris). Auk 107: 284-295. Nakatsuru, K. and Kramer, D. L. 1982. Is sperm cheap? Limited male fertility and female choice in the Lemon tetra (Pisces: Characidae). Science 216: 753-755. Nicholls, E. H., Burke, T. and Birkhead, T. R. 2001. Ejaculate allocation by male Sand martins. Proceedings of the Royal Society of London B 268: 1265-1270.

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Nitchuk, W. M. and Evans, R. M. 1978. A volumetric analysis of Sharp-tailed grouse sperm in relation to dancing ground size and organization. Wilson Bulletin 90: 460-462. Oring, L. W., Fleischer, R. C., Reed, M. C. and Marsden, K. E. 1992. Cuckoldry through stored sperm in the sequentially polyandrous Spotted sandpiper. Nature 359: 631-633. Parker, G. 1970. Sperm competition and its evolutionary consequences in insects. Biological Reviews of the Cambridge Philosophical Society 45: 525-567. Parker, G. 1982.Why are there so many tiny sperm? Sperm competition and the maintenance of two sexes. Journal of Theoretical Biology 96: 281-294. Partecke, J., Van’t Hof, T. and Gwinner, E. 2004. Differences in the timing of reproduction between urban and forest European blackbirds (Turdus merula): result of phenotypic flexibility or genetic differences. Proceedings of the Royal Society B 271: 1995-2001. Pellatt, E. J. and Birkhead, T. R. 1994. Ejaculate size in Zebra finches Taeniopygia guttata and a method for obtaining ejaculates from passerine birds. Ibis 136: 97-106. Pitcher, T. E. and Stutchbury, B. J. M. 1998. Latitudinal variation in testis size in six species of North American songbirds. Canadian Journal of Zoology 76: 618-622. Pitcher, T. E., Dunn, P. O. and Whittingham, L. A. 2005. Sperm competition and the evolution of testes size in birds. Journal of Evolutionary Biology 18: 557-567. Pizzari, T. and Birkhead. T. R. 2000. Female feral fowl eject sperm of subdominant males. Nature 405: 787-789. Presgraves, D. C., Baker, R. H. and Wilkinson, G. S. 1999. Coevolution of sperm and female reproductive tract morphology in Stalk-eyed flies. Proceedings of the Royal Society of London B 266: 1041-1047. Prinzinger, R., Pressmar, A. and Schleucher, E. 1991. Body-temperature in birds. Comparative Biochemistry and Physiology A 99: 499-506. Rand, A. L. 1933. Testicular asymmetry in the Madagascar coucal. Auk 50: 219-220. Ray, J. 1678. The Ornithology of Francis Willughby. John Martyn, London. Retzius, G. 1909. Die spermien der vogel. Biologische Untersuchungen 14: 89-122. Retzius, G. 1911. Spermien ostafrikanischer tiere. Biologische Untersuchungen 16: 8288. Rising, J. D. 1987. Geographic variation in testis size in Savannah sparrows (Passerculus sandwichensis). Wilson Bulletin 99: 63-72. Rising, J. D. 1996. Relationship between testis size and mating systems in American sparrows (Emberizinae). Auk 113: 224-228. Risser, A. C. 1971. A technique for performing laparotomy on small birds. Condor 73: 376-379. Salvador, A., Veiga, J. P., Martin, J., Lopez, P., Abelenda, M. and Puerta, M. 1996. The cost of producing a sexual signal: testosterone increases the susceptibility of male lizards to ectoparasitic infection. Behavioral Ecology 7: 145-150. Schärer, L., Ladurner, P. and Rieger, R. M. 2004. Bigger testes do work more: experimental evidence that testis size reflects testicular cell proliferation activity in the marine invertebrate, the free-living flatworm Macrostomum sp. Behavioral Ecology and Sociobiology 56: 420-425. Schulze-Hagen, K., Leisler, B., Birkhead, T. R. and Dyrcz, A. 1995. Prolonged copulation, sperm reserves and sperm competition in the Aquatic warbler Acrocephalus paludicola. Ibis 137: 85-91. Selander, R. K. and Hauser, R. J. 1965. Gonadal and behavioral cycles in the Greattailed grackle. Condor 67: 157-182.

## Reproductive Biology and Phylogeny of Birds Shugart, G. W. 1988. Uterovaginal sperm-storage glands in sixteen species with comments on morphological differences. Auk 105: 379-384. Soley, J. T. 1993. Ultrastructure of Ostrich (Struthio camelus) spermatozoa. 1. Transmission electron-microscopy. Onderstepoort Journal of Veterinary Research 60: 119-130. Sordahl, T. A. 2001. Copulatory behavior of American avocets and Black-necked stilts. Auk 118: 1072-1076. Sossinka, R. 1982. Domestication in birds. Pp. 373-403. In D. S. Farner, J. R King and K. C. Parkes (eds), Avian Biology, volume 6. Academic Press, New York. Stanley, A. J. and Witschi, E. 1940. Germ cell migration in relation to asymmetry in the sex glands of hawks. Anatomical Record 76: 329-342. Stockley, P., Gage, M. J. G., Parker, G. A. and Møller, A. P. 1997. Sperm competition in fish: the evolution of testis size and ejaculate charateristics. American Naturalist 149: 933-954. Stutchbury, B. J. and Morton, E. S. 1995. The effect of breeding synchrony on extrapair mating systems in songbirds. Behaviour 132: 675-690. Tsuji, L. J. S., Kozlovic, D. R. and Sokolowski, M. B. 1992. Territorial position in Sharptailed grouse leks: the probability of fertilization. Condor 94: 1030-1031. Tuttle, E. M. and Pruett-Jones, S. 2004. Estimates of extreme sperm production: morphological and experimental evidence from reproductively promiscuous fairy-wrens (Malurus). Animal Behaviour 68: 541-550. Tuttle, E. M., Pruett-Jones, S. and Webster, M. S. 1996. Cloacal protuberances and extreme sperm production in Australian fairy-wrens. Proceedings of the Royal Society of London B 263: 1359-1364. Van Drimmelen, G. C. 1946. “Spermnests” in the oviduct of the Domestic hen. Journal of the South African Veterinary Medical Association 17: 43-52. Weatherhead, P. J. 1984. Mate choice in avian polygyny: Why do females prefer older males? American Naturalist 123: 873-875. Weatherhead, P. J., Metz, K. J., Bennett, G. F. and Irwin, R. E. 1993. Parasite faunas, testosterone and secondary sexual traits in male Red-winged blackbirds. Behavioral Ecology and Sociobiology 33: 13-23. Werdelin, L. and Nilsonne, A. 1999. The evolution of the scrotum and testicular descent in mammals: a phylogenetic view. Journal of Theoretical Biology 196: 6172. Westneat, D. F., McGraw, L. A., Fraterrigo, J. M., Birkhead, T. R. and Fletcher, F. 1998. Patterns of courtship behavior and ejaculate characteristics in male Redwinged blackbirds. Behavioral Ecology and Sociobiology 43: 161-171. Wetton, J. H., Burke, T., Parkin, D. T. and Cairns, E. 1995. Single-locus DNA fingerprinting reveals that male reproductive success increases with age through extra-pair paternity in the House sparrow (Passer domesticus). Proceedings of the Royal Society of London B 260: 91-98. Wikelski, M., Hau, M., Robinson, W. D. and Wingfield, J. C. 2003. Reproductive seasonality of seven neotropical passerine species. Condor 105: 683-695. Wingfield, J. C. and Moore, M. C. 1987. Hormonal, social, and environmental factors in the reproductive biology of free-living male birds. Pp. 149-175. In D. Crews (ed.), Psychobiology of Reproductive Behavior: An Evolutionary Perspective. Prentice Hall, NJ. Witschi, E. 1935. Origin of asymmetry in the reproductive system of birds. American Journal of Anatomy 56: 119-141.

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Wolfson, A. 1952. The cloacal protuberance—a means for determining breeding condition in live male passerines. Bird-Banding 23: 159-165. Wolfson, A. 1954. Sperm storage at lower than body temperature outside the body cavity in some passerine birds. Science 120: 68-71. Woolley, D. M. 1995. The structure of the spermatozoa of the Japanese quail, Coturnix coturnix L., var. japonica. Acta Zoologica 76: 45-50. Wright, J. and Cuthill, I. 1989. Manipulation of sex differences in parental care. Behavioral Ecology and Sociobiology 25: 171-181. Wright, P. L. and Wright, M. H. 1944. The reproductive cycle of the male Red-winged blackbird. Condor 46: 46-59. Wyndham, E., Hutchison, R. E. and Brockway-Fuller, B. F. 1981. Gonadal condition of non-breeding wild and domesticated Budgerigars Melopsittacus undulatus. Ibis 123: 511-518. Yu, Z.-H. 1998. Asymmetrical testicular weights in mammals, birds, reptiles and amphibia. International Journal of Andrology 21: 53-55. Zann, R. A., Morton, M. S., Jones, K. R. and Burley, N. T. 1995. The timing of breeding by Zebra finches in relation to rainfall in central Australia. Emu 95: 208222. Zuk, M., Johnson, K., Thornhill, R. and Ligon, J. D. 1990. Parasites and male ornaments in free-ranging and captive Red jungle fowl. Behaviour 114: 232-248.

n n

CHAPTER

10

Fertilization Urszula Stepinska1 and Murray R. Bakst2

10.1

INTRODUCTION

Until recently, the cellular and molecular events comprising fertilization in birds were rather poorly understood. However, several recent studies in mammals as well as in birds have considerably contributed to our knowledge of the fertilization process. Mammals exhibit physiological monospermy, that is, only a single sperm enters the ovum. Monospermy is ensured by a variety of mechanisms, which is collectively known as the block to polyspermy. Unlike mammals, the principal feature of fertilization in birds, as well as in many fishes, amphibians and reptiles, is physiological polyspermy, that is, penetration of the ovum by many sperm. It has been suggested that polyspermy occurs in animals that produce large yolky (megalecithal) ova. This review is not a comprehensive account of all aspects of avian fertilization but rather concentrates on the following: oviductal sperm selection, storage and transport; the interaction of gametes; the role of envelope of the ovum; and formation and further fate of pronuclei. The main focus is on the complexities of polyspermic fertilization and the mechanisms preventing excessive pathological polyspermy that ensure normal embryonic development. This chapter also includes some data on assisted reproductive technologies used in birds, such as artificial insemination (AI), in vitro fertilization (IVF) and intracytoplasmic sperm injection (ICSI). Although the latter two methods have yet to find practical application, they can shed some light on at least some mechanisms of avian fertilization in vivo. Most of the available information on avian fertilization relates to the chicken (Gallus gallus), since it is commonly used as an experimental bird. In this chapter, all observations will be derived from research with the chicken unless otherwise noted. 1

Institute of Genetics and Animal Breeding, Polish Academy of Sciences, Jastrzebiec n. Warsaw, 05-552 Poland. E-mail: [email protected] 2 Biotechnology and Germplasm Laboratory, Beltsville Agricultural Research Center, ARS/ USDA, Beltsville, MD 20705 USA. E-mail: [email protected]

##" Reproductive Biology and Phylogeny of Birds

10.2 MORPHOLOGY OF THE OVULATED OVUM As in most other vertebrates, the avian ovum is ovulated with its nucleus at metaphase of the second meiotic division. The voluminous 3-4 cm diameter ovum is primarily comprised of yellow yolk. On its surface is a 3-5 mm diameter germinal disc (GD), composed of white yolk that forms the animal pole. The GD contains the female complement of genetic material as well as the majority of cytoplasmic organelles (Bakst and Howarth 1977a; Perry et al. 1978). At the GD the plasma membrane, the oolemma forms an array of microvillar projections (Fig. 10.1), which are longer and more abundant than that of the vegetal hemisphere where the oolemma is discontinuous and forms few elaborations (Bakst and Howarth 1977a; Bakst 1978). Given the role of physiological polyspermy in the fertilization process, it should not be surprising that cortical granules have not been observed in the ooplasm of avian oocytes (Guraya 1982).

10.2.1 The Inner Perivitelline Layer At the time of ovulation the ovum is enveloped by an acellular glycoprotein coat known as the inner perivitelline layer (IPVL), sometimes erroneously referred to as the inner layer of vitelline membrane in the laid egg (Bellairs et al. 1963; Bakst and Howarth 1977a; Okamura and Nishiyama 1978a; for

Fig. 10.1 Gallus gallus. A scanning electron micrograph (SEM) showing the microvilli-like extensions of the oolemma at the germinal disc (GD). White yolk spheres are observed in the GD. Original.

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nomenclature see Baumel et al. 1979) (Fig. 10.2A, B). The IPVL is considered to be analogous to the egg envelope of other animal species such as the zona pellucida (ZP) of mammals, vitelline envelope of frogs, and the chorion of fishes (see Dumont and Brummett 1985). The IPVL of birds is a threedimensional network of fibers (Bellairs et al. 1963), where spaces between the fibers are filled with a ground substance (Bakst and Howarth 1977a) (Fig. 10.2A, B). Bakst and Howarth (1977a) observed that the part of the IPVL overlying the GD and that covering yolk had an identical structure and exhibited similar variation in thickness (2-4 mm). On the other hand, it has been claimed that the IPVL from the GD region appears thinner, and has smaller and less numerous fibers than that from other regions of the ovum (Bellairs et al. 1963; Perry et al. 1978). In mammals, the ZP is composed of three glycoproteins designated as ZP1, ZP2 and ZP3 (Wassarman 1988), also known as ZPB, ZPA and ZPC, respectively (Harris et al. 1994). Two glycoproteins have been identified in the IPVL of the avian ovum. One displays significant homology to members of the ZP3/ZPC family of mammals and therefore has been named either chicken ZP3 (chkZP3, 32 kDa) (Kido and Doi 1988; Waclawek et al. 1998; Takeuchi et al. 1999) or quail ZP3 (qZP3, 33 kDa) (Mori and Masuda 1993; Kuroki and Mori 1995; Pan et al. 2000). The other IPVL glycoprotein is a homologue of the ZP1/ZPB family, about 95 kDa, and designated chkZP1 (Bausek et al. 2000) or qZP1 (Sasanami et al. 2003). The presence of immunological homologues of chkZP1 and chkZP3, identified by Western blotting with anti-chicken ZP1 and ZP3 antibodies, respectively, has been demonstrated in the IPVL from laid eggs of several avian species including Japanese quail, duck, goose, pheasant and turkey (Stewart et al. 2004). Both glycoproteins showed some differences in molecular mass among the avian species studied: ZP1 from 91 to 106 kDa and ZP3 from 34 to 42 kDa. There are no data indicating the presence of a third glycoprotein in avian IPVL, a homologue of ZP2 found in the mammalian ovum envelope. It is possible that the putative ZP2 homologue is much less abundant in avian IPVL than ZP1 and ZP3, and/or that it is lost during IPVL isolation and solubilization (Bausek et al. 2000). In chicken and quail, glycoprotein ZP3 is synthesized by the granulosa cells surrounding the oocyte during the rapid growth phase of the ovarian follicle (Kuroki and Mori 1995; Waclawek et al. 1998; Takeuchi et al. 1999; Pan et al. 2001; Sasanami et al. 2002). It has been suggested that testosterone stimulates ZP3 production at the gene transcription level in the granulosa cells (Pan et al. 2001). In contrast, avian ZP1 glycoprotein seems to be synthesized under estrogen control in the liver and then transported through the bloodstream to the ovarian follicle (Bausek et al. 2000; Sasanami et al. 2003). Thus, the site of biosynthesis and the hormonal regulation of avian IPVL are different for ZP1 and ZP3. In fish, chorion proteins appear to be synthesized in the liver of spawning females in response to estrogens and are then transported to the ovary by blood circulation (Hamazaki et al. 1989). In Xenopus, there is clear evidence that all egg envelope glycoproteins are

##$ Reproductive Biology and Phylogeny of Birds

Fig. 10.2 Gallus gallus. Initial step of interaction of sperm with inner perivitelline layer (IPVL). A. A differential interference contrast (DIC) micrograph of the IPVL with a single sperm (arrow) highlighted by a fluorescent nuclear stain. Note the fibrous nature of the IPVL. Original. B. A SEM of a sperm with an intact acrosome on the surface of the IPVL. The arrows indicate remnant cytoplasmic processes possibly from the oolemma (top arrow) and the granulosa cells (bottom arrow). Original.

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synthesized by the oocytes (Yamaguchi et al. 1989). This is likewise true for mice (Bleil and Wassarman 1980). However, in other mammals, such as rabbits, humans and pigs both the oocyte and its accompanying granulosa cells contribute to the synthesis of the ZP glycoproteins (Grootenhuis et al. 1996; Kolle et al. 1996). Interestingly, Olszanska et al. (2001) have identified by RT-PCR the presence of ZP3 transcripts in the GD and the ooplasm in quail oocytes. This highlights the possibility of ZP3 synthesis in the avian oocyte in addition to that in the granulosae cells.

10.3

OVIDUCTAL SPERM SELECTION, STORAGE AND TRANSPORT

This section will provide a brief overview of oviductal sperm selection, storage and transport in birds with emphasis on poultry. More detail explanations of the subjects can be found in reviews by Bakst et al. (1994), Wishart and Horrocks (2000) and Birkhead and Moller (1998), and the proceedings of a workshop edited by Hamlett (2003). The left oviduct in birds has three major functions: egg formation; sperm selection, storage and transport; and is the site of fertilization and early embryonic development. Egg formation begins when the ovulated ovum is captured by the fimbriated region of the infundibulum. Whether sperm are present or not, the ovum passes through infundibulum where albumen-like proteins form the attenuated continuous layer and the more extensive outer perivitelline layer (OPVL). This fibrous investment strengthens the PVL complex supporting the megalecithal ovum in its transit through the oviduct. The oviductal ovum progresses through the magnum where the albumen is secreted, the isthmus where the shell membrane is formed, and then the egg resides in the shell gland (uterus) for shell formation prior to its brief transit through the vagina at oviposition. This “ovulatory cycle”, that is, ovulation, egg formation, and oviposition is about 24-25 h in duration and is repeated with each egg in a clutch. If the ovum was fertilized in the infundibulum, the initial cleavage furrows would appear in about 5-7 h after sperm penetrates the ovum, about when the egg was in the isthmus or uterus. Depending on the species, the fertilized, fresh laid egg posseses a blastoderm consisting of 30,000 to 60,000 cells. That being said about egg formation and the site of fertilization and early embryonic development, the main objective of this section is to present our current understanding of oviductal sperm selection, storage and transport. Briefly, in hens that have mated naturally or subjected to AI, a relatively small number of sperm are transported to storage sites at the anterior end of the vagina, a region referred to as the utero-vaginal junction (UVJ). Here sperm enter the sperm storage tubules (SST) which are discrete tubular structures originating as invaginations of the UVJ surface epithelium (Fig. 10.3A, B). Sperm that successfully traverse the vagina and populate the SST are distinct in both mobility characteristics and surface membrane composition.

##& Reproductive Biology and Phylogeny of Birds

Fig. 10.3 Gallus gallus. Sperm in sperm storage tubule (SST). A. Sperm are observed at the orifice of a SST. The arrow indicates the sharp transition between the ciliated epithelium of the uterovaginal junction and the SST epithelium. Original. B. Fluorescing sperm are observed in the basal lumen of a SST viewed by DIC. The outside diameter of the SST is about 40 mm. Original.

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Depending on the species, sperm reside for variable lengths of time in the SST before they are released and ascend to the infundibulum, the site of fertilization and presumptive secondary sperm storage site. Sperm transport is probably a combination of sperm mobility and activity of the ciliated epithelium lining the oviductal mucosa. There has been no significant research done in the factors controlling oviductal sperm transport in birds for many years (see Bakst et al. 1994 and Wishart and Horrocks 2000 for reviews). What we do know that the vast majority of sperm initially transferred into vagina at copulation or AI fail to populate the SST. The known exception to this is when a commercial turkey is inseminated 1014 days after photostimulation but prior to the actual onset of egg production. For reasons that we can only speculate, sperm numbers in the SST are nearly twice as great in the SST when inseminated just prior to the onset of egg production as when the hen was AI initially after the onset of egg production (Brillard and Bakst 1990; Bakst 1994). But even under these circumstances, the maximum number of sperm in the SST remains below 10 million even though the hen was inseminated with 150-350 million sperm. Obviously, the vagina orchestrates an intense sperm selection process. The mechanisms behind such selection remain to be elucidated. Interestingly, if a hen is inseminated within 0.5 h following oviposition, upon manual eversion of the vagina, the inseminator may actually be able to deposit the semen in the uterus, bypassing the vagina and UVJ. While large numbers of sperm are transported to the infundibulum, this results in increased embryo mortality, possibly by pathological polyspermy. The sperm which do reach the UVJ and enter the SST can be viewed as “selected” sperm (see Bakst et al. 1994). Yet, what favors the transport of some sperm and not other sperm to the SST remains unknown. As pointed out by Wishart and Horrocks (2000), sperm transport from the site of deposition in the vagina to the UVJ requires that the sperm be motile and that the sperm plasmalemma possesses certain glycoproteins. Differences in sperm mobility, as measured by the ability of sperm to move progressively through a viscous medium, contribute to the likelihood of sperm success in traversing the vagina (Froman et al. 1999; Birkhead et al. 1999). Likewise there appears to be control by the hen as to which sperm she will accept in the SST or to interact with the hen’s ovum (Pizzari et al. 2002). Elucidating the cellular and molecular basis for successful sperm selection will no doubt lead to improvements in AI technology, specifically, the insemination of lower numbers of sperm at longer intervals between successive inseminations. One must assume that sperm residing in UVJ SST undergo a reversible suppression of certain activities. They may include: 1) reversible suppression of sperm metabolism and motility; 2) stabilization of membrane systems; 3) stabilization of the acrosomal enzymes; and 4) suppression of sperm immunogenicity (Bakst et al. 1994). One needs also to assume that the sperm exiting the SST undergo some type of activation, possible comparable to mammalian capacitation (see 10.4.2 and Chapter 4).

#$ Reproductive Biology and Phylogeny of Birds A long contentious issue is whether sperm release from the SST is associated with a particular stage of the ovulatory cycle, such as immediately following oviposition, or whether they are released slowly over the course of the ovulatory cycle. Recently Froman (2003) suggested that sperm residing in the SST swim against a fluid current generated by the SST epithelium that flows in the direction of the UVJ lumen. Once the velocity of this current exceeds the velocity of the sperm, sperm are swept out of the SST. The localization of aquaporins in the SST epithelium (Zaniboni and Bakst 2004) would appear to add support to Froman’s hypothesis. Interestingly, extensive innervation of the UVJ mucosa surrounding the SST as well as an actinfilament network in apical cytoplasm of the SST epithelial cells suggests that individual SST may be capable of contracting, thereby facilitating the release of resident sperm (Freedman et al. 2001). Table 10.1 contains a hypothetical model describing the fate of sperm after transfer to the vagina. Table 10.1 Hypothetical model for oviductal sperm transport and storage in birds. Based on works by Bakst et al. (1994), Poultry Science Reviews 5: 117-143 and Froman (2003) Biology of Reproduction 69: 248-253. • • • • •

A small percentage of the sperm transferred to the vagina reach the SST SST resident sperm are subjected to a reversible suppression of metabolism and motility, inhibition of enzyme systems, and stabilization of the plasmalemma Resident sperm metabolize lipids and carbohydrates released from SST epithelium As resident sperm mobility decreases, increasing numbers of sperm are swept out of the SST Liberated sperm are transported to the site of fertilization at the infundibulum

It has long been assumed that the infundibulum serves as a secondary sperm storage site in the chicken and turkey oviduct (see Bakst et al. 1994 and Wishart and Horrocks 2000 for reviews). However, upon extensive examination of the infundibular mucosa within 48 h of insemination no more than 1 to 3 sperm were ever located (Bakst 2003). When one considers the benefits of the UVJ SST to reproductive success (Table 10.2) and given the paucity of a sperm in the infundibulum after a normal insemination or copulation, one is led to the conclusion that the infundibulum is not a true sperm storage site analogous to the UVJ SST. Table 10.2 • • • •

Benefits of sperm storage in the UVJ SST Elimination of synchronization of copulation with ovulation Sperm transfer to hen is not necessary for production of fertile eggs over one or more clutches Provides resevoir for “selected” sperm Affords protection to sperm during the daily ovulatory cycle

We have no knowledge of what cross-signaling may exist between resident sperm and the SST or what genes are turned on and off in the presence or

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absence of resident sperm. Long et al. (2003) constructed 2 SAGE (separate serial analysis of gene expression) libraries with SST RNA obtained from turkey hens inseminated with diluted semen or semen extender only. They found that only 1 percent (214) of the putative genes was differentially expressed between SST with or without resident sperm. These data support the idea that differential gene expression occurs in the SST mucosal epithelium within 48 h after AI with semen. This raises the possibility that sperm themselves induce specific gene expression events required for prolonged sperm storage and/or release from the SST.

10.4 10.4.1

INTERACTION OF SPERM WITH THE OVUM Site of Fertilization

In birds, fertilization takes place in the infundibulum, the most anterior segment of the oviduct, within about 15 minutes of ovulation (Olsen and Neher 1948). Sperm must contact the surface of the ovum, specifically the IPVL overlying the GD, before appreciable amounts of infundibular secretory material is deposited around the IPVL. This tertiary investment derived from the infundibular mucosa forms the OPVL, which sperm cannot penetrate. It is the OPVL that prevents pathological polyspermy (Fig. 10.4). The mechanisms responsible for drawing the sperm to the GD remain unknown.

Fig. 10.4 Gallus gallus. The perivitelline layer complex composed of the inner (IPVL) and outer perivitelline layers (OPVL) are observed. The OPVL is formed by infundibular secretory activity and functions as a block to pathological polyspermy. Numerous sperm are seen embedded in the OPVL. After Bakst, M. R. and Howarth, B. H. 1977b. Biology of Reproduction 17: 370-379, Fig.12.

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Reproductive Biology and Phylogeny of Birds

10.4.2 Capacitation In mammals, ejaculated sperm are progressively motile yet do not have an immediate capacity for fertilization. In the case of in vivo fertilization they gain this ability after residing in the female genital tract or, in the case of IVF, after incubation for some time in an appropriate medium. The physiological changes that render the sperm competent to fertilize is referred to as capacitation (see Austin 1952; Yanagimachi 1994). It is generally believed that capacitation of sperm is not essential for fertilization in birds (see Howarth 1984; also Chapter 4). This was primarily based on observations that avian ova could be fertilized in vitro by fresh ejaculated sperm which had not been previously treated with a special capacitation medium (Howarth 1971; Nakanishi et al. 1990; Olszanska et al. 2002). It was demonstrated that the duration of gamete interaction needed for successful IVF in birds is short and does not exceed 10-15 min (Nakanishi et al. 1990; Olszanska et al. 2002). Likewise, initiation of in vitro hydrolysis of IPVL fragments by ejaculated sperm was found to occur within 2.5 min (Robertson et al. 1997). Moreover, it was shown in vivo that avian sperm do not need to reside long in the female reproductive tract to be competent for fertilization, since fertilized eggs could be observed within 15 min after AI into the vagina (Bobr et al. 1964) or infundibulum (Olsen and Neher 1948). While fresh ejaculated sperm can fertilize an ovum in vitro, the sperm that fertilize a daily succession of ova have resided days or weeks in the oviductal SST. Bakst et al. (1994) suggested that sperm residing within the SST are quiescent, with motility, metabolism, enzyme systems reversibly suppressed (“decapacitated” in the mammalian sense). Upon release from the SST, the oviductal milieu may activate sperm (capacitate?) prior to ascent to the site of fertilization.

10.4.3 Sperm-Inner Perivitelline Layer Interaction What can be considered the first step in the process of fertilization is binding of sperm to the IPVL (Fig. 10.5A, B). Sperm binding properties have been observed in chicken and quail IPVL even after its removal from oocytes (Howarth 1990; Kuroki and Mori 1997). Sperm-binding induces the acrosome reaction (AR) (Fig. 10.5B) thereby releasing hydrolytic enzymes that digest a path through the IPVL (Fig. 10.6A,B) (Okamura and Nishiyama 1978a; see Howarth 1984). The ability of IPVL to stimulate an AR was demonstrated in chicken in vitro by incubating sperm with fragments of IPVL isolated from freshly ovulated ova (Koyanagi et al. 1988) or with homogenized IPVL isolated from the PVL complex of laid eggs (Horrocks et al. 2000). Okamura and Nishiyama (1978a) documented the process of the AR and subsequent passage of sperm through the IPVL of the hen. They have shown that when the acrosome region of a sperm contacts the IPVL the outer acrosomal membrane and overlying plasmalemma of the sperm fuse at different points, thereby releasing its complement of hydrolytic enzymes. These hydrolytic enzymes digest small holes in the IPVL,

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Fig. 10.5 Gallus gallus. Interaction of sperm with IPVL. A. A SEM of a sperm making initial contact with IPVL. The arrow highlights remnant cytoplasmic processes on the IPVL surface. After Bakst, M. R. and Howarth, B. H. 1977b. Biology of Reproduction 17: 370-379, Fig. 14. B. A transmission electron micrograph (TEM) of a sperm head interaction with the surface of the IPVL. The plasmalemma overlying the acrosome is highly ruffled and discontinuous in places (arrow) suggesting the beginning of acrosome reaction. Original.

#$" Reproductive Biology and Phylogeny of Birds

Fig. 10.6 Gallus gallus. “Sperm holes” in the IPVL. A. A light micrographs of “sperm holes” (arrows) in the IPVL overlying the GD. These holes are sites where the sperm have hydrolyzed a path through the IPVL to reach the oolemma. Original. B. A TEM shows the hydrolyzed fibers and ground substances composing a portion of a sperm hole in the IPVL. Original.

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approximately 10 mm in diameter, through which the sperm penetrate to reach the oolemma (Figs. 10.6A, B, 10.7A, B) (Bakst and Howarth 1977b; Okamura and Nishiyama 1978a). Interestingly, sperm penetration of the IPVL is seen preferentially at the GD region of the ovum (see 10.4.3.4). Sites of sperm penetration through the IPVL can be observed in uterine eggs (Okamura and Nishiyama 1978a) or after oviposition (Bramwell and Howarth 1992a; Birkhead et al. 1994; Bramwell et al. 1995; Wishart 1997). Similar holes were found after the coincubation of sperm with fragments of the IPVL isolated from the largest preovulatory oocytes (Steele et al. 1994; Kuroki and Mori 1997; Takeuchi et al. 2001), from recently ovulated ova (Howarth 1990; Bramwell and Howarth 1992b; Steele et al. 1994) or from oviposited eggs (Steele et al. 1994; Robertson et al. 1997). It is assumed that a trypsin-like acrosomal enzyme, acrosin, is involved in the hydrolysis of the IPVL. Acrosin has been isolated from chicken (Ho and Meizel 1970; Froman 1990) and turkey (Richardson et al. 1988) sperm. The acrosin activity of turkey sperm is much higher (about 100 times) than that of human sperm and is similar to that of boar sperm (Glogowski et al. 2001). Ho and Meizel (1975) observed in vitro hydrolysis of IPVL of chicken ova by partially purified acrosin. It has been reported that the presence of trypsin inhibitors in the proteins forming the OPVL reduced sperm fertility in vivo (Palmer and Howarth 1973) and inhibited sperm hydrolysis of chicken and quail IPVL in vitro (Howarth and Digby 1973; Kuroki and Mori 1997; Takeuchi et al. 2001).

10.4.3.1

Molecular aspects of sperm-inner perivitelline layer interaction

Our understanding of the role of the ZP glycoproteins and the mechanism of fertilization is quite advanced for the mouse. All ZP proteins (ZP1-3) are structural glycoproteins necessary for the formation of the ZP during mouse oocyte growth (see Wassarman et al. 2004 and Yanagimachi 1994 for reviews). ZP2 and ZP3 form long filaments which are cross linked by ZP1 (see Wassarman et al. 2004). In addition to their structural properties, ZP2 and ZP3 also play roles in gamete interaction. Mouse ZP3 glycoprotein has been identified both as the acrosome-intact sperm receptor (primary sperm receptor) as well as the inducer of AR in the bound sperm (Litscher et al. 1995; see Wassarman et al. 2004 and Yanagimachi 1994 for reviews). Considerable evidence shows that O-linked oligosaccharides of mouse ZP3 account for the sperm receptor activity (Florman and Wassarman 1985; Bleil and Wassarman 1988), whereas the induction of AR reaction in the sperm is mainly dependent on its polypeptide chain (Leyton and Saling 1989). In turn, mouse ZP2 glycoprotein seems to be mainly responsible for the binding of acrosomereacted sperm to the ZP (secondary sperm receptors) (Bleil et al. 1988). Initially, ZP3 glycoprotein was postulated to be the most likely candidate for the sperm receptor also in avian IPVL (Waclawek et al. 1998; Pan et al. 2000). However, the above suggestion was based only on the extensive amino

#$$ Reproductive Biology and Phylogeny of Birds

Fig. 10.7 A. Gallus gallus. After staining the perivitelline layer (PVL) complex overlying the GD, seven sperm holes (arrows show 2 pairs of holes) can be viewed. Fig. 10.7 Contd. ...

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%$acid homology of avian and mammalian ZP3, and not on any experimental data. It has recently been suggested that another major glycoprotein of the IPVL, chkZP1, is responsible for the interaction of sperm with the IPVL. In vitro sperm penetration of the IPVL from the largest ovarian follicles was completely inhibited after pretreatment of the IPVL with monoclonal antibodies directed against chkZP1 (Takeuchi et al. 2001; Bausek et al. 2004). It has also been shown that in vitro sperm interaction with the IPVL leads to proteolytic activity that only affects chkZP1, degrading it into discrete fragments, while chkZP3 remains intact (Bausek et al. 2004). Alternatively, Takeuchi et al. (2001) have reported that sperm degraded IPVL chkZP1 and chkZP3 in vitro. Bausek et al. (2004) showed that chkZP1 and chkZP3 bind specifically to the acrosome of sperm in vitro. Thus, it appears that in birds the interaction between sperm and the IPVL is mediated by both chkZP1 and chkZP3. There is evidence that N-linked oligosaccharides of the IPVL with terminal N-acetylglucosamine residues may play a crucial role in the interaction of avian sperm with IPVL (Robertson et al. 2000). In their study, the in vitro interaction of sperm and IPVL from laid chicken eggs, measured as the number of holes produced in the IPVL, was significantly decreased after treatment of the IPVL with either: 1) N-glycanase (removing N-linked sugars from IPVL); 2) lectins with affinity for N-acetylglucosamine (masking the terminal N-acetylglucosamine in IPVL); or 3) when the sperm:IPVL interaction was carried out in the presence of N-acetylglucosamine (competing with endogenous N-acetylglucosamine). One should realize that the interaction of sperm with IPVL involves both the binding of sperm to the IPVL, the induction of the AR, and then the hydrolysis of the IPVL. In the above study, the overall sperm-IPVL interaction was measured by the number of holes produced in the IPVL, thus it is not possible to determine which stage of the interaction was affected by the experimental manipulation. However, using a stain specific for acrosome, it was revealed that N-linked oligosaccharides of the IPVL glycoproteins can induce the AR in chicken sperm (Horrocks et al. 2000). Of significance, their study revealed that the oligosaccharides need to have terminal N-acetylglucosamine groups and that they are capable of inducing the AR even when detached from their protein anchor. At present, it is not known to which IPVL protein the N-linked oligosaccharides are attached. That the IPVL N-linked oligosaccharides appear to be responsible for the induction of the AR does not exclude the possibility that they may influence the binding of sperm to IPVL and its hydrolysis after the AR.

Fig. 10.7 Contd. ...

Note the texture, lighter color, and doughnut-shape of the PVL overlying the GD. The diameter of the GD is about 3mm. Original. B. A SEM of the underside of the IPVL shows two sperm exiting a sperm hole. The sperm on the right appears to have lost its acrosome but not the one on the left. Complements of B. Howarth, Jr.

#$& Reproductive Biology and Phylogeny of Birds

10.4.3.2

Changes in the inner perivitelline layer between ovulation and oviposition

Steele et al. (1994) found that the number of holes in IPVL produced in vitro by chicken sperm was several times less in IPVL from laid eggs compared with that isolated from the largest follicular and freshly ovulated oocytes. However, it should be mentioned that in case of laid eggs whole PVL, not isolated IPVL, was used for the experiment. Likewise, the number of sperm that could bind in vitro to the IPVL at the surface of the GD was lower in laid eggs than in follicular oocytes (Kuroki and Mori 1997). A simple explanation may be that the IPVL sperm binding site are masked by the OPVL or the sperm are sterically hindered from reaching the IPVL by the components of the OPVL. In addition, the IPVL from laid eggs differs, from that of both follicular oocytes and ovulated ova. Chicken and quail ZP3 glycoproteins from laid eggs had lower molecular weights than the ZP3 glycoproteins isolated from follicular oocytes and ovulated ova (Kuroki and Mori 1995; Waclawek et al. 1998; Pan et al. 2000). Alterations of the quail ZP3 glycoproteins on the oviductal ovum includes removal of N-terminal amino acids (probably due to the action of a protease secreted by the infundibulum) and modifications of N-linked and Olinked oligosaccharide chains (Pan et al. 2000). Even though these changes were observed in virgin hens, the modifications of the ZP3 glycoproteins were suggested to be a block to excessive sperm IPVL penetration before the OPVL formed (Waclawek et al. 1998; Pan et al. 2000). In contrast to the observations of Steele et al. (1994), Robertson et al. (1997) observed that the IPVL from laid chicken eggs, from follicular oocytes and ovulated ova gave similar responses to hydrolysis by sperm in vitro. However, it should be noted that in the above experiment IPVL was used not only in case of follicular oocytes and ovulated ova but also in case of laid eggs, as IPVL was isolated from PVL, whereas Steele et al. (1994) used whole PVL from laid eggs. Robertson et al. (1997) did observe that one of the IPVL glycoproteins, probably ZP3, appeared smaller in laid eggs than from follicular oocytes. Notwithstanding, these authors proposed the use of IPVL from laid eggs instead of IPVL from follicular oocytes or ovulated ova. The former are considerably easier to obtain when performing in vitro assays of sperm-IPVL interaction for predicting the fertilizing ability of semen from different males.

10.4.3.3

Species specificity in sperm-inner perivitelline layer interaction

In mammals, sperm interaction with the ZP is usually described as a speciesspecific event, although cross fertilization may occur between closely related species (see Yanagimachi 1994). The interaction between sperm and IPVL from guinea fowl, turkey and chicken was demonstrated by IPVL hole formation following homologous and heterologous coincubation in vitro (Steele et al. 1994). However, sperm showed a slightly higher specificity towards the homologous IPVL. Stewart et al. (2004) coincubated chicken sperm

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with IPVLs from laid, unfertilized eggs from closely and distantly related avian species and demonstrated that chicken sperm were able to hydrolyze the IPVL of all species examined. The functional cross reactivity between chicken sperm and heterologous IPVL seemed to correlate with the phylogenetic distance between the species. Within the order Galliformes, the interaction of chicken sperm with the IPVL from turkey, quail, peafowl, pheasant and guinea fowl was not significantly different from that with the homologous chicken IPVL. The hydrolytic activity of chicken sperm towards IPVL was strongest within Galliformes, intermediate in Anseriformes (goose and duck) and the weakest in Passeriformes (zebra finch) and Columbiformes (dove). These findings show that, unlike the mammalian ZP, the IPVL does not represent such strong species specific barrier to heterologous sperm penetration. This may be a factor facilitating hybrid production both in wild and domesticated birds after AI.

10.4.3.4

Preferential attraction of sperm to the inner perivitelline layer over the germinal disc

During fertilization in birds sperm bind to and penetrate the IPVL over the entire surface of the ovum, although with a much greater frequency on the IPVL overlying the GD. This is advantageous in ensuring syngamy of the male and female nuclei. Preferential sperm penetration of IPVL overlying GD has been demonstrated in laid eggs of several avian species after natural mating (Bramwell and Howarth 1992a; Birkhead et al. 1994; Steele et al. 1994; Bramwell et al. 1995; Wishart 1997; Wishart and Staines 1999) and in freshly ovulated and then in vitro fertilized chicken ova (Bakst and Howarth 1977b). The number of sperm penetrating the IPVL (number of holes produced by sperm in IPVL observed in fertilized laid eggs) over GD is approximately 20 times greater than in other regions of the egg in both chicken and turkey (Bramwell et al. 1995; Wishart 1997) (Fig. 10.7A). The total number of holes in the IPVL region over the GD, which is about 3 mm in diameter, can reach several hundred. However, because this represents a minor fraction of the area of the whole IPVL, there are approximately 50-fold more holes found evenly distributed in the IPVL outside this region (Wishart 1997; Wishart and Staines 1999). The mechanisms responsible for the preferential hydrolysis of the IPVL overlying the GD remain unknown, despite considerable interest and speculation (Ho and Meizel 1975; Bramwell and Howarth 1992b; Steele et al. 1994; Kuroki and Mori 1997). This may be attributed to different densities of sperm receptors, differential induction of the AR or different susceptibility of various regions of the IPVL to hydrolysis. Kuroki and Mori (1997) have found that sperm receptors are concentrated in the IPVL over the GD disc region of quail oocytes from preovulatory follicles. This was demonstrated in an assay in which hydrolysis of the IPVL by sperm in vitro was inhibited with trypsin inhibitors and the binding of sperm to the IPVL could be observed. An alternative explanation is that the IPVL overlying the GD region may be more

#% Reproductive Biology and Phylogeny of Birds susceptible to hydrolysis since that region of the IPVL of chicken ova is preferentially hydrolyzed in vitro by partially purified cock sperm acrosin (Ho and Meizel 1975). Steele et al. (1994) observed a higher density of holes in the IPVL over GD in fertile laid eggs, but after in vitro coincubation of fragments of the IPVL from ovulated ova with sperm, holes appeared with similar frequency throughout the whole IPVL surface. These data do not support the conclusions of Bramwell and Howarth (1992b) that in vitro sperm preferentially hydrolyze the GD region of IPVL (isolated from ova) and that this region is particularly rich in sperm receptors. However, it should be noted that the above conclusions were based on a single result only in which the concentration of holes in the IPVL over the GD region was only 1.33 times that found in the IPVL from other ovum areas. Although some authors reported that the IPVL appeared thinner in the GD region, Bakst and Howarth (1977a) found the IPVL over the GD and remaining regions of the ovum structurally identical. It was also demonstrated that the electrophoretic protein patterns of IPVL over the GD and non-GD regions were indistinguishable (Steele et al. 1994). It is therefore possible that the preferential hole formation in the GD region is not associated with the IPVL but originates from the GD itself. Bakst (1978) found that the microvillar projections of the oolemma were longer and more abundant at the GD region when compared to the remainder of the oolemma (Figs. 10.1, 10.2B, 10.5A). He suggested that the oolemma projections may protrude through the IPVL and that they were possibly the site of sperm receptors. He later indicated that the remnant processes seen on the surface of the IPVL by scanning electron microscopy could also have originated from the granulosa cells (Bakst 1979) (Fig. 10.2B). Wishart (personal communication) has taken the view that some components in the yellow yolk act to suppress sperm hydrolysis of the IPVL. His suggestion supports the observation of few IPVL sperm holes in intact ova are observed overlying the yellow yolk whereas, if the IPVL is washed free of the yellow yolk, sperm holes are more abundant. It is interesting to note that sperm penetrate the IPVL over GD in such a manner as to form a doughnut-shaped configuration with the center of the GD remaining largely unpenetrated (Fig. 10.7A). This was demonstrated both in vivo, in the IPVL isolated from fertile laid eggs of several avian species (Birkhead et al. 1994; Wishart 1997), as well as in vitro in chicken after incubation of IPVL fragments (isolated from ovum) with sperm (Bramwell and Howarth 1992b). The hypothetical physiological advantage of the central area of IPVL over the GD being less attractive to sperm is unclear at present but it is likely that it may restrict excessive sperm entry into the GD in the vicinity of the female nucleus (see 10.5).

10.4.4

The Outer Perivitelline Layer and its Role in Blocking Excessive Polyspermy

In birds there is no block to polyspermy at the level of IPVL. However, a block to pathological polyspermy is in the form of tertiary investment, the OPVL,

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deposited by the distal infundibulum and proximal magnum around the IPVL (Bellairs et al. 1963; Bakst and Howarth, 1977b; Okamura and Nishiyama 1978a). Ultrastructurally, the OPVL is composed of irregularly deposited concentric arrays of what are assumed to be fibrous proteins. These fibers are densely arranged near the IPVL, then the space between the concretions gradually widens. A single “continuous layer” is observed separating the IPVL from the OPVL. Biochemically, the OPVL is composed of enzymatically active lysozyme (about 60% of dry weight), an insoluble ovomucin complex (Back et al. 1982), and two basic proteins, the 17.5-kDa VMO I (vitelline membrane outer I) (Back et al. 1982) and the 6-kDa VMO II (Kido et al. 1992). Ovomucin appears to form the skeleton of OPVL, but some other proteins, especially lysozyme, are responsible for its integrity (Back et al. 1982). The functions of VMO I and VMO II are not known. Luminal sperm in the upper region of the oviduct become passively trapped between and within the fibers of the OPVL and are incapable of penetrating this investment (Fig. 10.4). Sperm embedded in the OPVL can be observed in fertilized eggs recovered from distal infundibulum and proximal magnum (Bakst and Howarth 1977b) or from laid eggs (Wishart 1987; Wishart 1997). The OPVL is thought to act as a block to excessive pathological polyspermy and to ensure the integrity of the ovum by preventing the rupture of IPVL, weakened by the sperm hydrolytic activity (Bakst and Howarth 1977b; Okamura and Nishiyama 1978a). Under in vitro conditions, preparations of OPVL can inhibit the hydrolysis of IPVL by sperm as well as interfere with the induction of AR by IPVL (see Wishart and Horrocks 2000). Sperm trapped in the OPVL of chicken eggs do not appear to have undergone an AR (Okamura and Nishiyama 1977b) and if acrosin were released, the OPVL contains trypsin inhibitors (see Howarth 1984) that would inhibit hydrolytic activity. Therefore, it is possible that OPVL presents not only a mechanical barrier preventing further sperm penetration into the IPVL but can also inhibit the AR and acrosin activity. Fertile eggs possess about ten times more sperm trapped in the OPVL than there are holes in the IPVL (Wishart 1997). In chicken and turkey eggs sperm trapped in OPVL were evenly distributed over the ovum (Wishart 1987; Wishart 1997). However, in several other avian species the density of sperm in OPVL appeared higher over the GD area than elsewhere (Birkhead et al. 1994). In fertile eggs of the emu and ostrich the number of sperm trapped in OPVL over the GD was very low in the center of GD and then increased rapidly in a band around the center of GD. Here the concentration of sperm was the highest and then gradually declined with increasing distance from the center (Malecki and Martin 2003). Thus, the pattern of distribution of sperm trapped in the GD region of the OPVL of emu and ostrich eggs corresponds with the pattern of holes produced by sperm in the same region of the IPVL that has been observed by some authors in laid eggs of several avian species (Birkhead et al. 1994; Wishart 1997).

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Reproductive Biology and Phylogeny of Birds

In laid eggs of chickens and turkeys there is nearly a linear relationship between the number of holes in the IPVL over the GD, the number of holes in the IPVL over non-GD regions, and the number of sperm trapped in the OPVL (Bramwell et al. 1995; Wishart 1997; Wishart and Staines 1999). The above authors found a logarithmic relationship between these three parameters and the egg fertility status in chicken and turkey and concluded that estimation of any of these parameters could be applied for assessing the mating efficiency of commercial turkey and broiler breeders.

10.4.5 Sperm Penetration into the Ovum Following their passage through the IPVL, one or more sperm interact with the oolemma and are incorporated into the ooplasm (Fovanova 1965; Bekhtina 1966; Okamura and Nishiyama 1978b; Perry 1987; Nakanishi et al. 1990; Waddington et al. 1998). As the studies on the entry of sperm into the avian ovum have been mainly limited to the GD region, the fate of sperm that penetrated the non-GD region remains unknown. Although sperm holes are seen throughout the IPVL, the non-GD oolemma is discontinuous and there is an absence of ooplasm containing organelles. It is assumed that these sperm do not contribute to the fertilization process. It has been suggested that holes in the IPVL in the non-GD region were artifactual and the result of acrosin release from sperm trapped in the OPVL adjacent to the IPVL (Birkhead et al. 1994; Steele et al. 1994). Likewise, Okamura and Nishiyama (1978b) claimed that although sperm may penetrate any part of the IPVL, the entry of sperm into the ovum occurs only within the GD region. Following penetration of IPVL over the GD, the apex of the sperm head which has undergone the AR comes in close contact with the oolemma (Okamura and Nishiyama 1978b). After the surface of the inner acrosomal membrane has come in contact with the oolemma, the two membranes fuse and the sperm passes into ooplasm. Sperm penetration into the ovum by means of a phagocytic process has also been observed in chickens (Okamura and Nishiyama 1978b). However, as the sperm plasmalemma is intact, it is not clear if male pronuclei are formed.

10.4.6 Formation of Pronuclei and Syngamy Sperm nuclei decondense within the GD to form male pronuclei (Okamura and Nishiyama 1978b; Perry 1987). During this process there is a disintegration of the nuclear envelope and a subsequently swelling of the nucleus as it transforms from an elongated to spherical organelle. This is accompanied by a decondensation of the chromatin and, at later stages, a reconstitution of the degraded nuclear envelope. This transformation was observed in chicken ova removed from the upper part of magnum after in vivo fertilization (Okamura and Nishiyama 1978b; Perry 1987) as well as after IVF (Nakanishi et al. 1990). As a consequence of polyspermy many male pronuclei are formed in the GD. The number of pronuclei observed by different authors in the GD of naturally fertilized chicken ovum was as follows: 1-60, average

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10 (Fofanova 1965); 20-60 (Bekhtina 1966); 5-15 (Perry 1987); 6-16 (Waddington et al. 1998); and 1-30 in the case of IVF (Nakanishi et al. 1990). Prior to syngamy, the association of the female and male pronuclei, the supernumerary male pronuclei are evenly scattered in the GD and are indistinguishable from the pronuclei destined to form the zygote nucleus. At the onset of male pronuclei formation, the female nucleus was arrested in metaphase of the second meiotic division. Formation of the female pronucleus commences with the completion of its second meiotic division at the time when male pronuclei formation is nearly completed (Bekhtina 1966; Perry 1987). Syngamy takes place when a fertilized ovum has progressed to the posterior part of the magnum, i.e. about 3-5 h after ovulation (Perry 1987; Waddington et al. 1998) or about 4 h following IVF (Nakanishi et al. 1990).

10.4.7

The Fate of Supernumerary Male Pronuclei

The supernumerary male pronuclei, which do not take part in syngamy, gradually disperse to the periphery of the GD where they can undergo one or even two rounds of mitotic division, but all of them degenerate and disappear at the early cleavage stage (Fofanova 1965; Bekhtina 1966; Perry 1987; Gupta and Bakst 1993; Waddington et al. 1998). The mechanisms responsible for the degradation and disappearance of supernumerary male pronuclei in early avian embryos remain unknown. In the polyspermic egg of the newt Cynops pyrrhogaster displacement of the supernumerary male pronuclei from their points of entry in the animal hemisphere into the vegetal hemisphere has been observed (Iwao et al. 1993). However, unlike birds, the accessory pronuclei never undergo mitotic divisions and degenerate before the first cleavage division.

10.5

POSSIBLE INVOLVEMENT OF DNASES IN LATE CYTOPLASMIC BLOCK TO POLYSPERMY

In mammals, transgenic animals can be produced by the microinjection of exogenous DNA directly into a pronucleus of the fertilized ovum (Brinster et al. 1985). In birds, direct injection of DNA into zygote pronuclei is not possible because: 1) they are masked by the presence of yolk spheres and 2) they cannot be distinguished from the supernumerary male pronuclei. So in birds the injections of DNA are done into the central part of the GD, where the zygote pronuclei are usually located (Perry 1987; Waddington et al. 1998). DNA injection into the ooplasm of a fertilized avian ovum rarely, if at all, leads to chromosomal integration of the foreign DNA. Most of it gradually disappears during early embryonic development (Sang and Perry 1989; Perry et al. 1991), probably contributing to the low efficiency of the method for obtaining transgenic chickens (Love et al. 1994) and quails (Naito et al. 1994). The fate of supernumerary sperm and foreign DNA in the GD led us to suspect that avian oocyte DNases are responsible for the degradation of excess DNA in early embryos. In Japanese quail oocytes DNase I and DNase

#%" Reproductive Biology and Phylogeny of Birds II activities were high and capable of digesting, under in vitro conditions, both naked lDNA/HindIII substrate as well as DNA contained in quail sperm (Stepinska and Olszanska 2001, 2003). It was also found that the activities of both DNases increased during oogenesis and reached maximum in mature preovulatory oocytes and ovulated ova. Stepinska and Olszanska (2001, 2003) speculated that these enzymes degraded the DNA of supernumerary male pronuclei resulting from polyspermic fertilization. Interestingly, the presence of high DNase activities in the GD would explain the low efficiency of production of transgenic birds by DNA injection into GD since the exogenous DNA would be targeted by the DNases. DNase I and II activity is not limited to the GD of ovulated quail ova but is found outside the GD in the thin layer of cytoplasm surrounding the yolk subjacent to the IPVL (Stepinska and Olszanska 2003). No DNase activities were found in the yellow yolk of ovulated quail ova (Stepinska and Olszanska 2003), which is consistent with the observation of visually unchanged sperm in the yolk of fertilized ova removed from the uterus (Bekhtina 1966). In contrast to avian ova, DNase I and II activities could not be observed in ovulated mouse ova under in vitro conditions (Stepinska and Olszanska 2003). Boone and Tsang (1997) also showed the absence of DNase I in rat oocytes from antral follicles, although they found the presence of the enzyme in the younger oocytes from preantral follicles. Monospermic fertilization of mammalian ova is ensured by an early block to polyspermy activated immediately after penetration of a single sperm. This block depends on modifications of the ZP (cortical granule-mediated zona reaction) and/or the oolemma (cortical granule-independent plasma membrane block) (see Yanagimachi 1994). Thus, from a functional perspective, the DNase activity in mature mammalian oocytes would be needless or even harmful for the fertilization process, since the enzymes could destroy the single sperm. However, polyspermic fertilization of mammalian ova has often been observed under a variety of experimental conditions, such as IVF, or excess of sperm in the case of AI. This usually results in abnormal embryo development, polyploidy, mosaics etc. (see Yanagimachi 1994). Thus, in mammals, when several sperm do overcome the block to polyspermy, these sperm may remain functional and they may participate in cleavage divisions causing developmental abnormalities. It seems that in birds the lack of an early block preventing multiple sperm penetration into the ovum might be compensated by the high DNase activities in the GD. While not a true block to polyspermy, the enzymes participating in the degradation of the supernumerary male pronuclei would certainly help minimize polyploidy and subsequent embryo mortality. Given the high activity of the GD-DNases around the time of pronuclei formation, why are the male and female pronuclei destined for syngamy not destroyed? The germinal vesicle (nucleus) of F1 follicle (largest oocyte destined to ovulate) is quite large and visible by eye at the center of the GD prior to germinal vesicle breakdown (GVBD). According to the studies of

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Bekhtina (1966) and Yoshimura et al (1993), the germinal vesicle enters the maturation phase by resuming meiosis and is transformed from a spherical to a discoidal form as it moves towards the surface of the GD. With extrusion of the first polar body, the second metaphase plate is formed and GVBD has commenced (Bekhtina 1966; Yoshimura et al. 1993). Given its vast volume, it is assumed that GVBD and the subsequent liberation of the nucleoplasm dilutes the impact of DNases on the pronuclei involved in syngamy and activation of the ovum (Stepinska and Olszanska 2003). Studies on natural polyspermic fertilization in the newt ovum indicated a possible influence of a gradient of the maturation/M phase-promoting factor (MPF) activity on the behavior of accessory male pronuclei (Iwao and Elinson 1990; Iwao et al. 1993; Sakamoto et al. 1998). A higher level of MPF activity was observed in the center of the animal hemisphere of the newt ovum versus a lower level in the vegetal hemisphere where the accessory male pronuclei degenerate. An injection of cytoplasm from unfertilized ova or germinal vesicle material from full-grown immature oocytes into fertilized newt vegetal hemisphere rescue accessory male pronuclei that subsequently form extra bipolar spindle (Iwao and Elinson 1990). These results suggested cytoplasmic factors present in the animal but not in the vegetal hemisphere control entry of the pronuclei into the M phase and that the nucleus is involved in the control of MPF activation. It has also been demonstrated that nuclear material is indispensable for the increase of MPF activity during oocyte maturation in several amphibian species (Gautier 1987; Iwashita et al. 1998; Sakamoto et al. 1998). Our knowledge concerning the role of MPF in oocyte maturation and its presence in fertilized ova of birds is rather scarce. It has been reported that the largest vitellogenic quail oocytes contain MPF in the GD and that the MPF activity is markedly increased during oocyte maturation (Mori et al. 1991). Injection of a GD extract from the mature oocytes causes maturation of fullgrown immature Xenopus oocytes. No MPF activity was found outside the GD in either immature or mature quail oocytes. It has been suggested that the gradient of MPF activity may play a role in the behavior of pronuclei in avian ova after polyspermic fertilization (Stepinska and Olszanska 2003). DNase I and II might be responsible for the degradation of all but one of the numerous sperm entering the avian ovum. The choice of the single sperm predestined to form the zygote nucleus might depend on its point of entry into the avian ovum (Fig. 10.8). The one sperm that does enter the center of GD is presumably drenched with nucleoplasm and therefore might be saved from the degrading activity of DNases and would be subject to the action of MPF and nuclear MPF-stimulating factor(s). Other sperm entering periphery of the GD and nonGD region, would lack the joint effect of MPF and MPF stimulating nuclear component(s) and would be degraded by DNases present there. It should be noted here that the probability of many sperm entering at the central region of a GD is greatly reduced since, as mentioned before, penetration of IPVL over GD is much less frequent in the center (corresponding to the nuclear territory

#%$ Reproductive Biology and Phylogeny of Birds

Fig. 10. 8 Hypothetical behavior of sperm depending on their point of entry into the avian egg during polyspermic fertilization. Regions of sperm entry: A. Center of GD; the single sperm takes part in the formation of the zygote nucleus. B. Periphery of GD; the sperm form pronuclei but finally are degraded by DNases. C. Non-GD region; the sperm are degraded by the DNases. After Stepinska, U. and Olszanska, B. 2003. Zygote 11: 35-42, Fig. 5.

of GD) than in the surrounding ring (corresponding to the cytoplasmic region of GD) (see 10.4.3.4).

10.6

IS POLYSPERMIC FERTILIZATION IN BIRDS OBLIGATORY?

The findings that a greater proportion of inseminated sperm reach the site of fertilization in the female reproductive tract in birds than in mammals (Birkhead et al. 1993) and that the number of sperm reaching the ovum at fertilization is positively correlated with ovum size in a variety of avian species (Birkhead et al. 1994) are consistent with the idea that a larger ovum requires more sperm in order to be fertilized. In the past, it was generally believed that polyspermic fertilization in birds is necessary to ensure activation of their megalecithal ova. The conclusion that a minimum six accessory spermatozoa were required for normal development of chicken ovum supported the above idea (Fofanova 1965). However, other data provide evidence against the hypothesis of obligatory polyspermy in birds. First, it was observed that avian ovum may be fertilized even when few sperm are found in the OPVL of the GD region (Wishart 1987; Malecki and Martin 2002) or even when only one sperm penetrated the IPVL over that region (Wishart 1997). Second, if polyspermy were obligatory, we would expect a large number of sperm to be concentrated in the center of the GD, the most likely location of the female pronucleus (Perry 1987; Waddington et al. 1998). However, as mentioned earlier, the sperm penetration of IPVL, although higher in the GD region than outside it, is much lower in the very center of the GD than in the ring immediately around the center (Bramwell and Howarth 1992b; Birkhead et al. 1994). Third, excessive sperm penetration into the GD has been observed to cause abnormal development of avian embryos (Fofanova 1965; Bramwell et al. 1997). Finally, conclusive evidence against the necessity of polyspermy in birds was supplied by the recent study of Hrabia et al. (2003)

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which demonstrated that a single sperm introduced by ICSI into the GD of quail ovum was capable of activating the ovum for embryonic development. Thus, it seems that the large avian ovum, with a relatively small GD, requires physiological polyspermy in order to increase the chance of a sperm entering the right place in the GD, but not for fertilization process to commence. Otherwise, in the case of monospermy, a single sperm entering a non-GD region would prevent the ovum being fertilized.

10.7 10.7.1

ASSISTED REPRODUCTIVE TECHNOLOGIES Artificial Insemination

Included in AI technology are aspects of semen collection, semen evaluation, semen dilution and storage, insemination of the female, and evaluation of hen fertility. The first comprehensive manual on the subject is that from Lake and Stewart (1978). Somewhat more comprehensive is a text by Bakst and Wishart (1995) and laboratory guide book by Bakst and Cecil (1997) addressing AI technology. For captive bird propagation, there is excellent review by Gee et al. (2004). Fundamentally, AI is a 2-step procedure. First, the semen is collected and second, the semen is inseminated. What transpires between these two steps is a function of management objectives, the skill level of the personnel involved, and the degree to which purposeful selection of breeder stock and their progeny drives the breeding program. Manual semen collection in birds was first described by Burrows and Quinn (1937) and the same basic “abdominal massage” technique is still in use today. A gentle massaging and stroking of the cloacal region and abdomen elicits an ejaculatory response from the male. The semen can be collected and left undiluted, or “neat” prior to AI. However, unless inseminated within 30-45 min of semen collection, the sperm will lose their viability. An alternative approach is to collect and dilute the semen with a semen extender that has been formulated to prolong the survival and viability of sperm. Semen dilution and storage has been reviewed several times over the years. Unfortunately, the technology here has not advanced significantly since the 1980s (Christensen 1995). While some have added antibiotics and anti-oxidants to commercially available extenders, conclusive prove of their efficacies remains equivocal (Surai 1999; Breque et al. 2003). The benefits of using semen extender are numerous, with more possibilities as the duration to store sperm in vitro increases. By diluting the semen with extender, one increases the ratio of male:female for breeding purposes. For example, when using neat semen, or when birds are allowed to copulate, the ratio of male: female can be 1:8 or 1:10. After dilution of semen with an appropriate extender, the ratio of male: female can be 1:20 to 1:30. The potential ramifications of using extenders for dilution and the storage of semen include: increased selection pressure on male breeders, an opportunity to evaluate semen quality, an opportunity to inseminate a known number of sperm, use of fewer sperm per insemination, and better male management.

#%& Reproductive Biology and Phylogeny of Birds That being said, no significant changes have been made in semen collection, dilution, and insemination over the past few decades. What has improved are the means by which semen is evaluated and fertility estimated (see Bakst and Cecil 1997). In this millennium, some poultry breeders are selecting males based on sperm mobility through a viscous medium, referred to as the sperm mobility assay (Froman et al. 1999). They found a strong positive correlation in reproductive success (paternity) and the ability of sperm from a given male to progressively move through a viscous medium. This observation appears to be applicable to both domestic and feral birds (Pizzari et al. 2002). Devices that integrate progressive motility and sperm concentration into a sperm index are available on the market and have proven to be useful to some. Careful dissection of fresh laid eggs can provide considerable information about the fertility status of a flock. By examining the morphology of the GD, a trainer observer can determine if the ovum was fertilized or left unfertilized. At the time of lay, a fertilized GD will be at the blastoderm stage of development (Wilson 1995; Wishart 1995; Bakst et al. 1998). This blastoderm will be symmetrical, uniformly opaque, and about 3 mm in diameter. An unfertilized GD is small, asymmetrical and be characterized by variable number of vacuoles sometimes visible by eye. If one is unsure if the GD was fertilized, two additional procedures can be quickly performed. With a fine pipette, puncture the ovum just outside the GD area and bring the pipette tip to the center of the GD. Gently aspirate some of the white yolk from the GD region and mix with a nuclear fluorescent stain such as bis-benzimide. If cells are present, whether from an early dead or a viable embryo, their nuclei will intensely fluoresce. If done carefully the PVL overlying the GD is still intact. This can be removed, washed clean of adhering yolk material and either stain with the same fluorescent nuclear stain to reveal sperm embedded in the OPVL, or the PVL could be examined using the PVL hole assay (see 10.4.3.4 and 10.4.4). The PVL hole assay has provided a means to estimate flock fertility as well as the duration of fertility for individuals hens. While time consuming to perform, the procedure is simple and provides objective results.

10.7.2 In vitro Fertilization The development of IVF and embryo manipulation techniques in mammals has significantly improved our understanding of the mechanisms of fertilization, activation of the ovum, and early embryonic development. In birds, IVF and associated technologies are not extensively developed, possibly due to the difficulties in both obtaining and manipulating mature yolk-laden ova. Howarth (1971) reported the first successful production of chicken embryos by IVF. Recently ovulated chicken ova retrieved from the body cavity or infundibulum were co-cultured with fresh sperm. The fertilization status of the ova was examined after 24 h of in vitro culture by hematoxylin staining of nuclei, which confirmed the presence of multicellular embryos. More recently, Nakanishi et al. (1990) provided direct evidence of IVF of chicken oocytes by

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demonstrating formation of male and female pronuclei after staining sections of a GD with 4,6-diamino-2-phenylindole (DAPI). Nakanishi et al. (1990) also concluded that early nuclear events, such as pronuclear formation, association of the male and female pronuclei, and elimination of supernumerary pronuclei, between in vivo and IVF during 1-4 h of in vitro culture were essentially the same. Tanaka et al. (1994) obtained viable chicks by transplanting in vitro fertilized ova into the oviduct of recipient hens followed by incubation of the eggs. However, the method used by those authors is complicated and requires surgical intervention. The oocytes used for IVF by all above authors were ovulated and recovered immediately from the body cavity or oviduct. Olszanska et al. (2002) successfully showed that quail embryos would develop normally after IVF of oocytes which had been ovulated in vitro. After IVF with fresh semen, and 20-22 h incubation, 14% of the blastoderms were between Stages IV-VI (see Eyal-Giladi and Kochav 1976) and contained DAPI-stained nuclei. Yet, under identical conditions, 25% of the “embryos” appeared normal but did not contain nuclei. Olszanska et al. (2002) suggested that the GD underwent segmentation without accompanying divisions of the nuclei. This phenomenon underlines the importance of monitoring the presence/absence of nuclei in early avian embryos, especially ones derived by IVF, to distinguish true cleavage from segmentation.

10.7.3

Intracytoplasmic Sperm Injection

Methods of direct injection of sperm into an ovum, referred to as intracytoplasmic sperm injection (ICSI) have not only resulted in normal offspring and transgenic animals, but increased our knowledge of some aspects of mammalian fertilization. For the first time an ICSI method in birds was reported by Hrabia et al. (2003). An intracytoplasmic injection of a single sperm into a quail ovulated ovum activated the GD and led to fertilization and the onset of development. The efficiency of fertilization after ICSI of ova was similar to that reported by Olszanska et al. (2002) for conventional IVF using 1-2 ¥ 104 sperm per ovum. The blastoderms after ICSI and 24 h culture reached Stages II-VII (see Eyal-Giladi and Kochav 1976), which is also comparable to the results of Olszanska et al. (2002). It was also shown that injection of a single testicular sperm or elongated spermatid into a quail ovum resulted in fertilization and embryo development to Stages VI-VII. These results indicated that both testicular sperm and elongated spermatids were capable of activating the ovum, form pronuclei and begin early embryonic development when introduced into the ovum. In contrast, injection of a round spermatid into quail ovum did not result in successful fertilization. The findings of Hrabia et al. (2003) provide direct evidence against polyspermy being obligatory in birds.

10.8

ACKNOWLEDGMENTS

We are very grateful to B. Olszanska and G. Wishart for their helpful comments on the manuscript.

#& Reproductive Biology and Phylogeny of Birds

10.9

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n n

Index

A A. renalis medius 41 Abdominal Aperture 153 Abdominal Ostium of the Oviduct 153 Acanthizidae 20 Accessory Ducts 155 Accessory Sex Organs or Glands 37 Accipitres 11 Accipitridae 11, 13, 35, 149, 358 Achaetops 23 Acid Phosphatase Activity 100 Acrocephalidae 23 Acrocephaline Warblers 23 Acrocephalus 147, 448, 451, 457, 478, 517, 543, 549 Acrosin 565, 570, 571, 572, 582, 583, 585 Acrosome 282, 283, 285, 287, 289, 291, 293, 295, 297, 299, 335, 339, 350, 360, 365-369, 371-374, 376, 377, 379, 381, 383-385, 388-392, 394-397, 399-403, 405-407, 410, 411, 414-420, 422, 423, 425, 426, 428, 429, 433, 434, 436, 438450, 452, 454, 455, 458-461, 463, 467469, 471-473, 475, 476, 478-488, 490, 491, 493, 494, 496, 497, 499-502, 506, 508, 532-534, 556, 562, 563, 565, 567, 581 583, 584 Acrosome granule 283, 285, 287, 288, 295 *

Acrosome Reaction (AR) 295, 376, 377, 379, 508, 562, 563, 583, 584 Acrosome Vesicle 293, 295, 299, 350, 360, 367-369, 371, 373, 374, 376, 377, 379, 383, 385, 386, 390, 392, 397, 399, 400, 403, 405-407, 410, 411, 414-417, 419, 420, 425, 426, 428, 429, 433, 434, 436, 438-440, 442, 443, 450, 454, 458, 488 Actin 44, 72, 85, 376, 377, 379 Actions of FSH on Follicular Development 202 Actitus 441 Activin 51, 53, 110, 158, 202, 232, 257259, 261, 264, 273, 275 Additional Apertures 155 Adenohypophysis 190, 192, 194, 233, 234, 239, 327 Adrenal Gland 40, 96, 97, 103, 165 Adsorptive 72, 83 Aedon 357, 463, 467 Aegithalidae 23 Aegithinidae 20, 21 Aegothelidae 8, 10, 12 African Stonechats 207 Agapornis 362, 428, 434 Agelaius 141, 262, 304, 364, 449, 481, 486, 487, 520 Alauda 357, 463, 465

Some taxa given in tables (e.g. Table 8.3) are not listed in this index.

#' Reproductive Biology and Phylogeny of Birds Alaudidae 21, 358, 463 Albumen 149, 166, 167, 171, 201, 557 Alcae 12 Alcedinidae 9, 12, 14 Alcidae 14, 29, 358, 440 Alectoris 209, 226 Alkaline Phosphatase Activity 100 Allometric 540 Allometry 540 Alpine Accentors 513 Amazonetta 8 Ambient Temperature 319 American Avocet 519, 550 American Robin 363, 449, 450, 452, 454, 456, 467-470, 534, 536 AMH Synthesis 161 AMH Transcription 161 Amino Acids 184, 199, 568 Ammodramus 363 Amniote Spermatozoon 350 Amniote Symplesiomorphies 360 Amniote Synapomorphy 354, 360, 364, 365, 436 Amphibians 157, 236, 247, 350, 354, 553 Ampulla 37, 164, 165, 168 Anas 3, 7, 8, 31, 35, 44, 46, 55, 57, 59, 68, 79, 88, 91, 92, 101, 115, 131, 199, 208, 251, 280, 305, 322, 324, 325, 326, 330, 331, 332, 343, 356, 359, 361, 382, 405, 406, 409, 410, 425 Anatidae 7, 32, 126, 128, 129, 131, 137, 139, 142, 143, 281, 241, 358 Anatomical Nomenclature 168 Androgen 38, 51, 82, 94, 105, 106, 113, 166, 199, 226, 233, 236, 241, 256, 257, 263, 314, 315, 317, 319, 329, 333, 342, 346, 531 Androgen Binding 51, 106, 113, 342 Androgenic Hormones 166 Androstenedione 51, 200, 256, 257, 263, 274 Anhingidae 8, 11 Annulus 283, 291, 293, 294, 300, 350, 352, 360, 365, 369, 370-373, 379, 380, 383, 387, 389, 391, 399, 400, 403, 404, 407410, 413, 414, 418, 421, 425, 428, 433, 434, 439, 442, 443, 447, 451, 478, 491, 494, 497, 500, 533 Anser 109, 157, 250, 328, 342, 272

Anseranas 7, 131 Anseranatidae 7, 131 Anseriformes 4, 7, 28, 31, 34, 116, 117, 123, 125, 126, 140, 143, 157, 297, 355, 356, 358, 359, 361, 383, 405, 410, 427, 497, 502, 527, 569 Anseriforms 99, 353, 410, 426, 429, 436, 443 Antbirds 19, 30, 207 Anthus 357, 465, 467 Ant-pittas 19 Ant-thrushes 19, 445 Apicolateral Junctional Complexes 93 Apodiformes 8, 9, 10, 12, 339, 357, 358, 362, 410, 411, 418, 448, 497, 498, 502, 506 Apomorphy 2, 354, 355, 450, 496, 501 Apoptosis 158, 159, 178, 179, 221, 244, 249, 252, 255, 265, 267-270, 272-276 Apostlebird 21 Appendix Epididymidis 96-98 Appendix Paradidymidis 40 Apterygidae 6, 129, 200, 360 Apus 289, 297, 339, 357, 362, 411, 413, 414, 418, 448, 498, 499, 500, 506, 510 Aramidae 9, 11 Arcanator 24 Ardeidae 13, 358 Ardeotis 15 Artamidae 20 Arteria cranialis renalis 41 Arteria pudenta 41 Arteria renalis caudalis 41 Arteria testicularis 41 Arteriae testiculares accessoria 41 Artificial Insemination 113, 505, 509, 553, 577, 580-582, 584, 586 Asymmetrical Testes 517 Atrichornithidae 20 Auks 27, 440 Australasian Treecreepers 20 Australian Babblers 20 Australian Magpie 20 Australian Robins 21 Autoantibodies 53 Autoimmunocytes 53 Aves 3, 26, 28-35, 113, 115, 145, 147, 178, 181, 339, 346, 352, 355, 357, 360, 421, 438, 497, 499, 504-506, 508

Index Avian Egg 150, 176, 576 Avocets 13, 440, 540 Axoneme 61, 297, 299, 350-352, 360, 364, 365, 370, 371, 379-371, 374, 376, 379, 383, 385-387, 389, 391-393, 396, 397, 399, 400, 401, 403, 404, 408, 409-411, 413, 415, 417, 418, 420, 421, 423, 425, 426, 428, 431-434, 436, 438, 439, 440, 442, 443, 447, 451, 452, 454-456, 458, 460, 461, 463, 469, 470, 472-480, 482486, 488, 493, 496, 497, 500, 532 Aythya 356, 382, 405 B Babblers 20, 23 Balaenicipitidae 8, 440 Ballowitz 349, 355, 371, 405, 415, 417-419, 426, 441, 443, 456, 461, 463, 467, 477, 501 Bank Swallows 520 Barbets 8, 13, 26, 32, 34 Barn Swallows 517 Basal Cells 87, 89, 90, 95, 321 Basal Lamina 46, 50, 53, 54, 56-58, 61, 63, 80, 96, 153, 155, 157, 158, 162-164, 200, 254, 255, 271, 307, 309, 323-325, 335, 367, 431 Basal Plate or Capitulum 291 Batis 21 Batises 21 Baya Weaver 203 Bayesian Analysis 16, 23, 24 Bearded Tit 21, 135, 144 Beavan’s Bullfinch 451, 490, 533 Bee-eaters 9 Bengalese Finch 299, 300, 493, 544 Berrypeckers 20 b-catenin 159, 175 Biological Clock 181-183, 229 Blackbirds 25, 147, 503, 520, 533, 543, 549, 550 Blackcap 356, 457, 478 Black-headed Grosbeaks 523, 547 Black-necked Stilt 441, 519 Blackpoll Warbler 538 Black-throated Blue Warblers 518 Blastoderm 166, 262, 557, 578, 580

#'

Block to Polyspermy 553, 570, 573, 574 Blood-epididymal Barrier 71, 108 Blood-testis Barrier 49, 51, 53, 102, 104, 105, 110, 111, 279, 337, 338 Blue Tit 486 Blue-flycatchers 21 Body Temperatures 516 Bombycillidae 23, 358, 467 Bonasa 7 Boobies 8, 11 Boundary Tissue 53-59, 61, 87, 89, 95, 112 Bowerbirds 20, 504 Brachypteracidae 9, 12, 14, 31 Bradypterus 23 Brain 182, 184, 188, 189, 194, 197, 198, 200, 202, 205-207, 210, 216, 218-222, 224, 225, 228, 231, 233-240, 262, 272, 327 Brambling 521 Branta 128, 259, 361, 405, 410 Breeding 40, 109, 124, 126, 128, 133-135, 138, 141, 147, 148, 158, 162, 166, 181186, 188, 199, 200, 203-209, 211, 213, 215, 221, 224, 226, 228-231, 233, 235237, 242, 246, 248, 250, 251, 253, 258, 266, 267, 271, 272, 274, 275, 312, 315319, 335, 339-347, 505, 513, 515, 521528, 530-532, 543-548, 550, 551, 553, 577, 586 Breeding Strategies 182 Bristleheads 20 Broadbilled Sapayoa 17 Bucerotidae 9, 12, 15 Budgerigars 509, 522, 551 Bulbuls 23 Bullfinch 448, 451, 457, 490, 492, 501, 502, 504, 533, 542, 544 Buntings 25 Buphagus 24 Bush Warblers 23 Bush-shrikes 21 Bustards 11, 26, 33 Butcherbirds 20 Butterflies 540, 546 Buttonquails 9 Ballowitz 349, 355, 371, 405, 415, 417-419, 421, 423, 425, 426, 440, 443, 456, 461, 463, 467, 477, 501, 504, 532, 543

#'

Reproductive Biology and Phylogeny of Birds C

Cacatuidae 13-15, 26 Caiman 352, 354, 355, 509 Cairina 280 Calcareous Shell 166 Calcarius 25, 527 Calidris 357, 382, 441, 443 Callaeatidae 20 Caluromys 352, 508 Canary 21, 208, 236, 364, 410, 455 Canary-flycatchers 21 Capacitation 85, 315, 339, 559, 562, 580, 583 Cape Sparrow 457 Cape Weaver 364, 450, 484, 485 Capitonidae 2, 12-15, 34 Capitulum 291, 292, 385 Caprimulgiformes 8, 9, 12, 32, 355, 358, 362, 410, 411, 415, 497, 498 Caprimulgus 289, 291, 297, 355, 362, 415418, 426, 443, 498, 499 Carbendazim 83, 334 Cardinalidae 25 Cardinalis 364, 449, 481, 488 Carduelis 218, 233, 356, 357, 364, 451, 455, 457, 463, 464, 526, 548 Cariamidae 11 Carpodacus 211, 355, 356, 526 Carrion Crow 448, 451, 453, 456-458 Caspase-3 161, 268, 269 Casuariidae 3, 6, 124, 360 Casuarius 124, 360 Cathartidae 3, 9, 13, 35, 149 Catoptrophorus 441 cDNA 198, 199, 218, 219, 223, 235, 236, 238, 586 Cellular Associations 280, 301, 302, 305, 306, 347, Centriole 283, 285, 289, 291, 300, 338, 345, 350, 351-353, 355, 360, 367, 369, 371-374, 376, 397, 377, 379-381, 383, 385-387, 389, 391, 393, 395-397, 399, 400, 403-405, 407, 408, 410, 411, 413415, 417-423, 425, 428, 431 434, 436, 439, 442, 443, 446, 451, 455, 463, 472, 473, 475, 478, 480, 482, 483, 486, 494, 496-498, 500, 502, 509 Centropus 516 Certhia 363, 455, 478, 488

Certhiidae 23, 24, 363, 455, 478 Certhioidea 19, 23, 363, 455, 478 Cettiidae 23 Chachalacas 7 Chalaziferous Region 168 Charadrii 12, 34, 440 Charadriidae 12, 15, 358, 440, 441, 533 Charadriiformes 9, 11, 12, 15, 28, 29, 33, 353, 355, 356, 358, 362, 405, 418, 426, 427, 440, 443, 448, 498, 499, 501, 533 Charadrius 383, 441, 533 Chatshrike 21 Chelidon 356, 463 Chelonia 350, 352-354, 371, 374, 381, 436, 494, 495, 496, 505 Chick Embryos 150, 153, 159, 161, 164, 176, 242 Chick-quail Chimeras 165 Chiffchaff 457, 478 Chionidae 13 Chloris 356, 357, 364, 455, 463, 464, 467, 526, 548 Chordae Retis 64, 66, 70, 72 Chromatoid Body 291, 292, 335, 337, 338, 345, 346 Chrysomitris 357, 463 Ciconiidae 3, 9, 10, 13, 34 Ciconiiformes 8, 10, 149, 358, 362, 440, 443, 498, 500 Ciliated Cells 66, 68, 73, 74, 77, 80, 83, 84, 89, 94, 99, 163, 168, 171, 172, 328-323 Cinclidae 23, 24 Circadian Rhythm 203, 211, 212, 226 Circadian Rhythmicity 229, 240, 270 Circannual Rhythms 230, 272, 316 Circus 40 Cirl Bunting 364, 455, 456, 488 Cisticolas 23 Cisticolidae 23 Classification and Phylogeny 1, 37 Cliff Swallows 523, 532, 545 Climacteridae 20 Cloaca 37, 41, 85, 86, 95, 120-124, 127129, 131, 133-137, 139, 142, 146, 158, 162, 166, 172, 175, 201, 434, 519 Cloacal Protuberance 85-87, 112, 134136, 145-148, 199, 319, 516, 548, 551 Clutch Size 245, 246, 248, 250, 258, 266, 272, 530, 531

Index CNS 182-184, 188, 193 Coerebidae 358, 467 Colies 8 Coliiformes 8, 361, 499 Columba 15, 31, 38, 102, 112, 232, 287, 289, 293, 318, 355, 357, 362, 412, 418, 419, 421, 423, 425, 448 Columbidae 9, 14, 15, 358, 410 Columbiformes 8, 31, 297, 355, 358, 359, 362, 392, 411, 418, 427, 448, 494, 497500, 502, 569 Commericial Turkey 166 Common Magpie 521 Common Tern 85, 252, 274 Comparative Methods 137, 514 Compensation Hypothesis 518, 547

Connecting Duct 61, 72, 73, 83, 85 Connecting Piece 292, 300, 385, 397, 399, 400, 404 Conopophagidae 19 Continuous Layer 166, 168, 557, 571 Contopus 362, 444, 445, 447 Control of Avian Reproduction 216 Coprodeum 122, 172, 173, 175 Copulation 87, 98, 102, 103, 115, 116, 120, 122-124, 129, 131, 133-138, 140, 144-147, 185, 198, 222, 312, 318, 335, 504, 514, 518-521, 525, 530, 535, 544, 549, 559, 560 Copulatory Structures 116, 120, 136, 137, 144 Coraciidae 9, 12 Coraciiformes 9, 12, 15, 361 Corcorax 21 Cormorants 8, 11 Corn Bunting 457, 490 Coronaves 4, 5, 9, 10, 410, 415, 423, 426, 498, 499, 500, 501 Corticosterone 211, 235, 262, 272, 532, 545 Corvida 19, 355, 357-359, 363, 444, 446, 448, 449, 452, 455-459, 461, 501, 502 Corvidae 17, 21, 113, 346, 358, 363, 427, 445, 451, 455-459, 461 Corvoidea 17, 19, 20 Corvus 85, 208, 233, 271, 328, 355, 357, 363, 427, 446, 448, 449, 451-460, 494, 501, 504, 521

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Cost of Flight Hypothesis 518 Cotingas 20, 445, 446 Cotingidae 17, 20, 345, 358, 445, 446 Coturnix 39, 44, 47, 48, 50, 54, 55, 57, 60, 65, 67, 68, 83, 84, 98, 100, 103, 108, 110, 157, 176, 198, 204, 224, 226, 230, 232234, 236, 240, 242, 254, 259, 264, 277, 281, 287, 302, 309, 310, 313, 334, 336, 340, 342, 344, 347, 361, 385, 387, 388, 390-394, 405, 421, 423, 448, 451, 457, 497, 502, 507, 508, 511, 534, 551, 584, 585, 587 Coucals 516, 517, 548 Coursers 13 Cracidae 7, 33, 126, 131, 132, 136, 143 Cranes 9, 11, 31 Crescent-chests 19 Crex 356, 382, 438, 440 Cristae 59, 60, 289, 295, 323, 325, 326, 352, 354, 365, 369, 370, 373, 387, 397, 404, 407, 408, 413, 428, 433, 436, 470, 480 Crocodile + Avian Stock 355 Crocodiles 116, 117, 120, 121, 350-354, 371, 436, 443, 498 Crocodiloid Spermatozoon 371 Crocodylia 116, 350, 352, 354, 355, 381, 436, 496 Crocodylus 352-355, 494, 495, 506, 509 Crossbill 207, 231, 251, 271 Crotophaga 289, 293, 344, 362, 424-426, 500, 508, 509 Cryptic Female Choice 535, 538, 539, 544, 546 Cuckoos 3, 9, 10, 25, 27, 30, 32 Cuculidae 9, 10, 14, 30, 32, 358 Cuculiformes 3, 9, 10, 344, 355, 358, 361, 426, 498-500, 508, 509 Cuculus 355, 425, 426 Culicicapa 21 Curassows 7 Currawongs 20, 21 Cyanistes 485, 486 Cyanocitta 363, 447, 454, 456 Cycle of the Seminiferous Epithelium 281, 301-303, 305-307, 311, 312, 336, 340, 343 Cynops 573, 585 Cypselus 357, 362, 411, 413

#'" Reproductive Biology and Phylogeny of Birds D Dapple-throat 24 Darters 8, 11 Dasyornis 20 Deconychura 427, 445 Dehydrogenases 94, 99, 113 Dendrocitta 203, 226 Dendrocolaptidae 17, 19, 20, 358, 445 Dendrocalaptinae 446 Dendrocopos 355, 357, 412, 423, 425, 448 Dendrocygninae 7 Dendroica 363, 391, 427, 513, 518, 538, 546 Dense Fibers 299, 336, 339, 353, 369, 371, 372, 376, 379, 381, 385, 387, 393, 396, 399, 407-409, 413, 417, 421, 423, 425, 428, 433, 436, 439, 440, 442, 446, 447, 451, 456, 461, 463, 467, 469, 471-476, 478, 480, 482, 483, 486, 488, 494, 496, 497, 501, 502 Desmin 72 Development 49, 95, 102, 103, 105, 107, 109, 110, 112, 113, 137, 141, 142, 149, 150, 157-159, 161, 162, 166, 175-179, 182-186, 188, 189, 194, 199, 201, 202, 204, 208, 221, 227, 230, 232, 235, 239245, 247, 249-251, 253-257, 259, 262267, 270-276, 279, 283, 287, 289, 292, 295, 300-302, 306, 312, 313, 317, 320, 321, 323, 336, 337, 340, 343-345, 347, 352, 391, 413, 418, 428, 440, 447, 450, 453, 454, 478, 488, 506-509, 517, 542, 553, 557, 573, 574, 576-579, 581-585, 587 Developmental Abnormalities 516, 574 Dicruridae 21 Didelphis 352, 510 Diethylstilbestrol 161, 179, 585 Differentiation of the Left Oviduct 150 Diomedeidae 13, 251 Dipnoi 352 Dippers 23 Distal Centriole 283, 285, 289, 291, 300, 338, 351-353, 355, 360, 367, 369, 371374, 376, 379, 381, 386, 387, 389, 391, 393, 396, 399, 400, 403-405, 407, 408, 410, 411, 413-415, 417, 418, 420, 421, 422, 425, 428, 431-434, 436, 439, 443,

446, 451, 463, 472, 473, 475, 478, 483, 486, 494, 496-498, 500 Divers 8 Diving-petrels 11, 13 DNA Hybridization 3, 7, 26, 27, 30, 33, 34, 117, 118, 120, 441, 498, 500, 504, 509 DNA Sequences 1, 25, 26-31, 33, 34, 494, 505, 583 DNase 573-575, 585 Domestic Fowl 38, 40, 63, 95, 98, 101, 102, 104, 106, 108-113, 150, 175, 177, 193, 198, 229, 230, 232, 238, 239, 241, 275, 280, 321, 338, 342, 343, 345, 347, 392, 503, 505, 507, 508, 510, 521, 538, 543, 547, 580-586 Domestic Turkey 133, 538 Donacobius 23 Double Oviducts 149 Doublets 353, 354, 365, 369-372, 385, 387, 389, 391, 393, 399, 400, 405, 408, 410, 417, 418, 420, 421, 423, 433, 434, 436, 442, 456, 461, 470, 473, 475, 482, 483 Dromaius 38, 109, 318, 341, 360, 361, 364, 365, 374, 448, 451, 496, 584 Drongos 21 Dryoscopus 21 Duck 7, 54, 73, 128-130, 146, 157, 162, 210, 224, 280, 338, 343, 356, 382, 405, 418, 425, 433, 555, 569 Ductuli Aberrantes 97, 98, 103 Ductuli Efferentes Distalis 72 Ductuli Efferentes Proximalis 72, 73 Ductus Aberrans 97, 98 Ductus Conjugens 61, 65, 72, 83, 98 Ductus Deferens 37, 39, 41, 44, 61, 64, 71, 82, 83, 85, 86, 89, 90, 94, 95, 100, 106, 107, 110, 113, 135, 300, 328, 333, 430, 434 Ductus Epididymidis 61, 64-66, 83, 84, 91, 97, 98, 100, 101, 105-107, 110, 328 Dunnock 144, 335, 527, 544 Dynein Arms 370, 371, 387, 399, 408, 431, 476 E Early Embryonic Development 150, 166, 557, 573, 578, 579

Index Ectoparasite 532 Efferent Ducts/Ductuli Efferentes 61, 63, 65, 72-75, 77, 80-83, 85, 93, 95, 98, 99103, 105-109, 111, 113, 314, 328, 334, 344 Egg Formation; Sperm Selection 166, 557 Egg Production 143, 162, 166, 171, 252, 276, 559 Egg-mass 166 Ejaculate Size 520, 535, 539, 549 Elminia 21 Emberiza 356, 357, 364, 455-457, 465, 467, 488, 490, 533 Emberizidae 25, 134, 449, 455, 457, 488 Empidonax 534 Emx-2 157 Endonuclear canal 285, 287, 289, 293, 295, 355, 360, 367, 369, 371, 372, 374, 376, 377, 379, 381, 383-385, 392, 399, 403, 407, 410, 415-419, 431-434, 436, 438, 439, 494, 496, 497, 500 Endpiece 350, 360, 365, 366, 369, 371, 374, 376, 381, 383, 385, 387-389, 393, 396, 400, 401, 405, 407-410, 413, 415, 471, 473, 475, 477, 483, 532 Environmental Clues, Stimuli or Effects 183, 184, 188, 216, 243, 250, 270, 316, 319, 321, 327 Environmental Signals 181-184, 186-188, 205, 315, 318 Epididymal Duct/Ductus Epididymidis 61, 71, 83, 85, 87-92, 94-97, 100, 110, 112, 315, 328-330, 333, 515 Epididymis 37, 39, 40, 41, 44, 46, 61, 6365, 71, 72, 75, 83, 85, 86, 89, 94-98, 100104, 107-112, 177, 199, 314, 315, 327, 329, 334, 335, 339, 340, 342-344, 346, 386, 434, 510 Epithelio-mesenchymal Transformation 157 Epithelium 45-49, 51, 54, 57, 58, 61, 64, 66-68, 70-73, 75, 80-82, 84, 85, 87-90, 95-100, 102, 104-107, 110, 112, 113, 141, 150, 153, 155, 157-159, 162, 163, 164, 166, 168, 171, 172, 174, 175, 244, 253, 256, 271, 279, 280, 281, 282, 283, 301303, 305-308, 311, 312, 314, 321-324,

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328-330, 333-345, 557-561 Epithelium of the Müllerian Duct 157 Epoophoron 165, 176 Eptatretus 377, 508 Estrildidae 230, 364, 493 Estrogen Receptors 82, 99, 100, 108, 161, 247 Estrogenic Hormones 161 Ethynyloestradiol 162, 176 Eudromia 287, 293, 361, 377, 378, 496 Euplectes 364, 450, 484, 485 Eupodotis 15 Eurasian Bullfinch 448, 451, 457, 490, 492, 501, 502, 504, 533, 544 Eurasian Tree Sparrow 521 European Quail 204, 230 European Starlings 194, 206, 212-214, 224, 226, 227, 229, 230, 232, 521, 545 Eurylaimidae 17, 19, 33 Eurylaimides 19 Eurypygidae 9, 11, 410 Excurrent Ducts of the Testis 61, 82, 87, 89, 101, 107, 113, 313, 314, 327, 329, 333 Presecretion Phase 320, 333 Reproduction Phase 320, 328, 333 Regressive Phase 320, 329, 521 Refractory Phase 320, 321, 329 External (Coelomic) Glomeruli 150 Extracellular Matrix 157 Extra-Pair Copulations 145, 313, 335, 514, 518, 525, 530, 544, 545 Extremitates Caudales 37 Extremitates Craniales 37 F Fairy Bluebirds 24 Fairy Warbler 21 Fairy Wrens 20, 110, 136, 137, 146, 147, 543, 548, 550 Falcipennis 7 Falco 149, 362, 426, 444, 448, 499, 501 Falconidae 13, 29, 149 Falconiformes 9, 11, 29, 149, 355, 358, 362, 443, 448, 499, 501 Fantails 21 Female Choice 141-145, 525, 535, 538, 539, 544, 546, 548

#'$ Reproductive Biology and Phylogeny of Birds FEPC 142 Fertilization 115, 141, 142, 150, 166, 168, 171, 201, 252, 261, 262, 344, 376, 442, 503, 504, 507, 508, 514, 530, 535, 536, 545, 547, 550, 553, 554, 557, 559-562, 565, 568, 569, 572, 574-579, 583-587 Fibrolymphatic Bodies 122, 123, 125, 127 Fibrous Sheath 285, 291, 294, 336, 339, 350, 352, 354, 360, 365, 369-372, 374, 376, 379, 380, 381, 383, 385, 387, 389, 393, 400, 401, 404, 405, 410, 423, 428, 433, 434, 436, 442, 494, 496, 497 Ficedula 357, 449, 463, 465, 467 Fimbria 168 Fimbriated Region of the Infundibulum 166, 167, 557 Finches 25, 207, 225, 236, 242, 251, 262, 451, 518, 520, 534, 539, 549, 551 Finfoots 9, 11 Fitch Trees 3 Fitch-margoliash 3 Flagellum 282, 283, 289, 291, 299, 300, 341, 345, 372, 374, 380, 388, 389, 391, 392, 396, 399, 400, 401, 404, 413, 417, 418, 423, 425, 426, 429, 433, 438-440, 451, 454, 457, 458, 466, 469-476, 480, 486, 492, 493, 509, 532, 534, 536, 539 Flamingos 8, 10, 32 Flight Costs 138, 518, 532 Fluid-phase Endocytosis 83, 109 Follicle Atresia 265-267 Follicle Selection 246, 252, 255, 258, 260, 264, 266, 270, 276 Follicle Stimulating Hormone (FSH) 51, 194, 197-199, 201, 202, 205, 208, 220, 227, 232, 237, 242, 249, 250, 256-259, 261, 263, 264, 266, 269, 270, 317, 327 Food Availability 204, 243, 250, 251, 316, 318, 321 Forced Extrapair Copulations 129, 138, 139, 142, 143, 528 Formicariidae 17, 19, 358, 445, 446 Francolinus 7 Fregatidae 8, 11, 13, 31 Frigatebirds 8, 11, 31 Fringilla 356, 357, 363, 455, 463, 464, 505, 521 Fringillidae 25, 35, 358, 363, 427, 455, 457, 463, 490

Fulica 356, 359, 382, 438, 440 Furnarii 19 Furnariidae 17, 19, 358, 445, 446 G Galbuliformes 8, 10, 15, 361 Galliformes 4, 7, 26, 28, 32, 33, 75, 94, 116, 117, 126, 131, 140, 143, 157, 218, 252, 297, 300, 301, 355, 356, 358, 361, 383, 405, 410, 427, 448, 497, 502, 569 Galliforms 353, 389, 391, 400, 405, 410, 426, 428, 436, 443, 451 Galloanserae 4, 6, 28, 120, 126, 131, 132, 134, 136, 140, 143, 371, 383, 400, 410, 413, 418, 419, 423, 428, 431, 497, 498, 500, 502 Gallus 38, 42, 43, 64, 67, 71, 74, 76, 78, 79, 81, 96, 97, 101-104, 109, 111-113, 116, 150, 152, 154, 156, 157, 159, 160, 165, 167, 176, 193, 232, 245, 247, 254, 256, 264, 271-276, 280, 312, 343, 345, 346, 355, 356, 361, 382-386, 391-393, 396, 398, 401, 405, 410, 425, 427, 497, 504, 505, 508, 510, 511, 521, 543, 547, 553, 554, 556, 558, 561, 563, 564, 566, 582, 584-586 Gartner’s Ducts 165 GATA-3 157 Gaviidae 10 Gaviiformes 8, 362 Genetic Distances 3 Genetic Paternity 529 Genome Size 540, 541, 546 Geopelia 352, 362, 418, 419, 421, 422, 423 Germ Cells 38, 40, 46, 47, 49, 53, 72, 83, 99, 108, 247, 249, 273, 274, 276, 279, 292, 300-302, 304, 306-308, 311, 312, 321-323, 328, 329, 342, 344, 347, 517, 584 Germinal Disc (GD) 554 Glandular Grooves 168 Glareolidae 13 Glycoprotein Hormones 197 Gnatcatchers 23 Gnateaters 19 GnIH 189, 203, 210, 217-223, 225, 239 GnIH Peptide 218 GnRH 183, 189, 197, 198, 201, 205-207, 212, 214-216, 219-223, 231-238, 243, 250, 251

Index Goblet Cells 171 Goldfinch 364, 457 Golgi Complex 49, 50, 56, 59, 63, 68, 70, 77, 80, 91, 94, 283, 285, 286, 292, 299, 307, 328 Gonad 38, 157, 159, 161, 162, 165, 179, 181, 184, 194, 197, 198, 200, 205, 207209, 212, 213, 247, 317, 320, 516, 517, 545, 547 Gonadal Recrudescence 319, 332 Gonadotropin Releasing Hormone (GnRH) 189, 206, 235, 343 Gonadotropin Synthesis 220, 221, 240 Grallina 21, 359, 363, 456, 461, 462 Grallinidae 363, 456 Granular Body 455, 478, 479, 486, 488 Granular Helix 455, 478-480, 482, 483, 488, 502 Granulosa 159, 200, 201, 202, 253-261, 263-276, 555-557, 570, 585, 586 Granulosa Cells 159, 200, 201, 202, 254261, 263, 264, 267-276, 555-557, 570, 585, 586 Great Crested Flycatcher 444, 447, 449, 455, 534 Great Tit 232, 252, 342, 346, 457, 486 Grebes 8, 10, 32 Greenfinch 356, 357, 455, 464, 467, 526, 548 Grosbeaks 25, 523, 547 Ground-rollers 9 Grouse 7, 28, 116, 132, 238, 525, 549, 550 Growth 112, 158, 162, 164, 179, 193, 197, 199, 202-205, 210, 221, 224, 228, 229, 232, 234, 240, 242-245, 250-259, 261, 262, 266, 268-276, 319, 327, 346, 417, 508, 521, 523, 545, 555, 565, 583, 585, 587 Gruidae 9, 11, 13 Gruiformes 9-11, 15, 30, 32, 33, 356, 358, 359, 362, 418, 438, 498-501 Gruiforms 2, 9, 10, 11, 15, 353, 440, 443 Grus 362, 418, 431, 438-440, 443, 499, 501 Guans 7, 132 Guineafowl 7, 27, 54, 59, 61, 64, 73, 132, 281, 287, 291-293, 295, 302, 315, 328, 329, 352, 361, 392, 396-399, 401, 402, 404, 405, 421, 433, 448, 451, 497

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Gulls 27, 34, 247, 252, 440 Gymnorhina 20 H Haematopodidae 13, 15 Hamerkop 8, 440 Hawks 11, 112, 149, 150 Head 155, 176, 210, 300, 301, 335, 337, 341, 347, 360, 365-367, 369, 372, 374, 376, 377, 379, 385, 392, 405-408, 413, 417, 421, 429, 438, 444, 445, 448, 451, 454, 457, 475-477, 479, 480, 486, 487, 490, 492, 493, 504, 505, 507, 532, 533, 563, 572 Helical Membrane 413, 418, 425, 428, 444, 446, 447, 449, 450, 452, 454, 461, 463, 467, 472, 473, 475, 477, 478, 480, 481, 485, 486, 488, 492-494, 497, 500, 501, 502 Heliornithidae 9, 11, 30 Helmet Shrikes 21 Hemiprocnidae 12, 498 Himantopus 441, 519 Hippolais 355, 356 Hirundinidae 23, 358, 363, 449, 450, 455, 463, 477 Hirundo 85, 113, 251, 252, 274, 356, 449, 463, 477, 517 Hoatzin 3, 8, 10, 27, 30, 34 Homo 64, 280, 305 Honeyeaters 20, 28 Honeyguides 8 Hoopoe 9 Hormonal Cascade 182, 189 Hormone Control 181, 275 Hormones of the Pars Distalis 197 Hornbills 9, 28 House Crow 521 House Finch 211, 231, 526 House Martin 356, 455, 463 House Sparrows 147, 210, 212, 218, 225, 231, 237, 239, 345, 517, 519, 547, 548 Hume’s Ground Jay 16 Hummingbirds 8, 26, 27 Hydatids of Morgagni 165 Hydrobatidae 11, 13, 15, 358 Hydroxysteroid 94, 99, 109, 113, 341 Hylia 23

#'& Reproductive Biology and Phylogeny of Birds Hyliota 21, 29 Hylophylax 207 Hypothalamic Nuclei and Tracts 190 Hypothalamo-hypophysial Portal System 192 Hypothalamo-pituitary Unit 181, 183, 187, 188 Hypothalamus 181, 182, 187-190, 192, 197, 198, 201, 206, 210, 212, 217, 218, 221, 233-235, 270, 276 I Ibidorhynchidae 12 Ibisbill 12 Ichthyophis 153 Icteridae 25, 358, 364, 449, 455, 467, 486, 487, 503, 533, 543 Icterus 364, 486, 487 Immune Responses 532 Implantation Fossa 287, 291, 300, 350, 367, 385, 393, 397, 400, 403, 404, 405, 407, 411 In Vitro Fertilization (IVF) 553, 562, 572574, 579, 584, 586 Indian Tree Pie 203 Indicatoridae 12 Indifferent Gonad 38, 161 Infundibulum 153, 162, 165-168, 171, 192, 201, 536, 557, 559-562, 568, 571, 578, 580 Inhibin 51, 53, 107, 108, 110, 197, 202, 232, 264, 273 Inner Perivitelline Layer (IPVL) 166, 554 Internal Coincidence Model 211 Interphase 281 Interrenal (Adrenal Cortical-like) 165 Interstitial Nodules 165 Interstitial Tissue of the Testis 53, 105, 113 Interstitium 53-56, 59, 61, 321, 323, 326, 341 Intracellular Lipid 175 Intracytoplasmic Sperm Injection 553, 579, 583 Intraepithelial Lymphocytes 68, 70, 80, 87, 95, 101, 104 Intravascular Perfusion Fixation 93 Intromittent Organ 115, 139, 145, 545 Ioras 20

Irenidae 24 Iridoprocne 463, 477 Isthmus 162, 166-168, 171, 172, 201, 557 J Jacamars 8, 28, 34 Jacana 362, 418, 442, 448, 500, 501, 509 Jacanidae 14, 441, 442, 533 Jackdaw 85, 328, 329, 357, 452, 453, 456 Japanese Quail 44, 56, 71, 82, 95, 103, 108, 176, 234, 262, 287, 297, 302, 305, 307, 311, 340, 511, 534, 551, 555 Junctional Complexes 68, 70, 77, 93, 110, 279 K Kagu 11 Kinetics of spermatogenesis 311, 336, 344 Kingfishers 9 Kinglets 21

301, 305,

L Lagorchestes 352, 507 Lampetra 376, 505, 508, 510 Laniidae 21, 358, 455 Lanioturdus 21 Lanius 355, 357, 451-453, 455, 456, 458, 461, 513 Lapwings 440 Lari 12 Laridae 14, 15, 33, 358, 440, 441 Larus 15, 355, 356, 382, 426, 427, 440, 443 Leaf Warblers 23 Least Flycatcher 534 Left Oviduct 122, 150, 162-165, 167, 173, 248, 519, 557 Lek 525, 528 Leydig Cell 53, 55, 59, 60, 62, 102, 103, 105, 112, 319, 326 Life History Stage 182, 184-187 Limpkin 9, 11 Linkage Disequilibrium Hypothesis 540 Linnet 356, 451, 457 Lissamphibia 350, 352, 354 Logrunners 20 Lonchura 299, 300, 340, 364, 450, 452, 490, 492, 493, 507, 510, 544

Index Longitudinal Columns 354, 365, 369, 370, 371 Long-tailed Tits 23 Loons 8, 10 Lophonetta 8 Lovebird 295, 299, 300, 347, 362, 428, 434, 490, 492, 511 Loxia 207, 218, 231, 251, 271 Luscinia 357, 463, 465, 467 Luteinizing (Leutenising) Hormone (LH) 109, 193, 194, 196-202, 204, 205, 208, 211, 215, 220-222, 227, 235, 238, 239, 241, 242, 256-259, 261, 263, 264, 266, 269, 270, 277, 316-318, 327, 341, 343, 346 Lymphoid Enhancer Factor 1 159 M Macgregor’s Bird of Paradise 17 Macgregoria 17, 27 Macropus 107, 285, 340 Macrosphenus 23 Magellanic Plover 13 Magnum 162, 163, 166-168, 171, 172, 201, 557, 571-573 Magpie Goose 7, 131 Malaconotinae 20, 21 Male Germ Cells 40 Mallard 115, 127, 128, 131, 132, 199, 208, 280, 293, 302, 306, 323, 328, 329, 341, 359, 361, 405, 406, 409, 410, 433 Maluridae 20 Malurus 87, 136, 147, 519, 527, 543, 550 Mammals 37, 40, 41, 44, 46, 49, 51, 53, 54, 56, 59, 61, 63, 64, 66, 68, 70-73, 77, 81-83, 85, 86, 94, 98-100, 109, 158, 159, 161, 165, 198-200, 203, 209, 210, 212, 223, 228, 235, 236, 244, 247, 249, 255, 256, 265, 267, 280-282, 285, 289, 291, 292, 297, 300, 301, 302, 303, 305, 306, 312-315, 327, 336, 341, 342, 344, 350, 352, 353, 372, 436, 508, 514, 516, 536, 546, 547, 548, 550, 551, 553, 555, 557, 562, 568, 573, 574, 576, 578, 586 Manakins 20, 445 Manchette 287, 289-293, 297, 300, 344, 345, 415, 417, 418, 428, 492, 509 Mating Systems 126, 135, 141, 142, 346, 527, 528, 536, 538, 549, 550

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Matrix Metalloprotease MMP2 159 Maturation/M Phase-promoting Factor (MPF) 575 Mature Follicular Oocyte 149 Median Eminence 188-194, 198, 205, 215, 217, 218, 226, 228, 232 Megaluridae 23 Megapodes 7, 26, 116, 131, 146 Megapodiidae 7, 26, 118-120, 126, 131, 136, 137 Melampitta 21 Melanerpes 361, 423-425, 447, 454, 500 Melanocharitidae 20 Melatonin 202-204, 210, 215, 222-227, 230, 232, 233, 236, 237, 240 Meleagrididae 7, 15 Meleagris 38, 54, 88, 101, 106, 107, 116, 193, 226, 254, 264, 281, 283-286, 288, 290, 294, 296, 330, 334, 346, 355, 361, 391, 393, 396, 398, 401, 402, 448, 510, 538, 582 Meliphagidae 17, 20, 27, 28, 145 Meliphagoidea 17, 19, 20 Melopsittacus 49, 85, 285, 339, 362, 428430, 432, 434, 448, 499, 506, 509, 522, 551 Melospiza 198, 225, 238, 241, 251 Menuroidea 19, 20 Meropidae 9, 12, 24 Mesenchymal Sheath 157, 158, 164 Mesites 10, 11, 32 Mesitornithidae 9-11, 32, 410 Mesonephric Tubules 150, 159, 165 Mesonephros 97, 104, 150, 152, 153, 157, 161, 162, 165, 166, 178 Mesorchium 37 Mesotocin 192, 193 Metanephros 161, 162 Metaves 4, 9, 10, 410, 411, 423, 497-500 Microrecanalization 83, 101, 105 Midpiece 282, 291, 293, 296, 297, 335, 351, 352, 360, 364-367, 369-374, 376, 377, 379-381, 383-389, 391-393, 395397, 399-405, 407-409, 411, 413-415, 417-421, 423, 425, 426, 428, 429, 431434, 436, 438-440, 442, 443, 445-447, 451, 452, 454-461, 463, 467-469, 472478, 480, 483, 484, 486-488, 490-494, 496-502, 504, 532-534

$ Reproductive Biology and Phylogeny of Birds Mimic Thrushes 23 Mimidae 3, 23, 24, 358, 467 Mitochondrion/Mitochondria 50, 56, 59, 60, 68, 70, 77-80, 91, 94, 95, 217, 263, 287, 289, 293, 295-297, 299, 307, 309, 310, 323, 325-329, 332, 351, 352, 354, 365, 367, 369-372, 374, 376, 378-381, 383, 385-387, 389, 391-393, 396, 397, 399-401, 403, 404, 407, 408, 410, 411, 413, 414, 417, 418, 420, 421, 423, 425, 426, 428, 431-434, 436, 438-440, 442, 443, 446, 447, 451, 452, 454, 455, 458461, 463, 469, 470, 472, 473, 475, 477, 478, 479, 480, 483, 486, 488, 493, 494, 497, 500, 501, 502, 532 Mitochondrial Genomes 4, 32, 507 Mitochondrial Helix 450-452, 454-456, 463, 472-480, 482-485, 488, 490, 493, 502 Mitochondrial Sheath 297, 299, 389, 407, 408, 454, 455, 466, 478, 488, 490, 491 Mockingbirds 3 Molecular Data 1, 10, 32, 497 Molluscs 354 Molothrus 364, 449, 486 Momotidae 9, 12 Monarch Flycatchers 21 Monarchidae 21 Monogamy 129, 138, 142, 312, 318, 340, 519, 527, 528, 530, 536, 546 Monotremes 350, 352 Motacillidae 24 Motmots 9 Moustached Warbler 451, 478, 543, 544 Mucin 171, 175 Mucous Part of the Magnum 171 Müllerian (Paramesonephric) Duct 150, 153-156, 158, 159, 161, 162, 164, 175179 Müllerian Duct Aplasia 157 Müllerian Ridge 150, 157 Multicellular Glands 168, 171 Mus 82, 311 Muscicapa 356, 357, 463, 467 Muscicapidae 23, 24, 358, 363, 449, 463, 467, 472 Muscicapoidea 19, 23 Musophagidae 9, 10, 14, 15 Myiarchus 363, 444, 445, 447, 449, 534

Myofibroblast Cells 56, 57 Myrmecocichla 359, 363, 447, 449-452, 456, 467, 468, 472-474, 477, 479, 483, 502 N Neck 167, 168, 170, 171, 292, 300, 301, 335, 354, 367, 369, 372, 373, 377, 379, 381, 389, 391, 393, 396, 397, 400, 407, 446, 451, 475, 476, 478, 504 Neoaves 4, 8, 9, 120, 133-136, 383, 410, 423, 497-500 Neognathae 4, 29, 117, 132, 358, 383, 494, 497, 498 Neornithes 3, 27, 116, 145, 358, 496 Nephrostome-like Depressions 150, 153 Neural Pathways 182-184 Neuroendocrine Secretions 183, 184, 188 Neurohormones 189, 192 Neurohypophysis 194 Neuromodulator Mechanisms 184 Neurosecretions 182, 190 New World Blackbirds 25, 533 New World Sparrows 25 New World Vultures 3, 9, 27, 35 New Zealand Wattlebirds 20 Nine-primaried Oscines 24, 25, 35 Noduli Epididymidis 98, 103 Non-ciliated Cell Types 72 Non-ciliated Cells 66, 73, 74, 77, 83, 89, 99, 163, 328, 331-333 Non-monophyly 2, 6, 14 Non-passerine Birds 2, 15, 16, 17, 85, 87, 89, 135, 138, 140, 295, 299, 300, 352, 355, 358, 365, 371, 374, 381, 383, 391, 392, 399, 412, 418-420, 423, 428, 431, 433, 435, 436, 440, 444, 445, 446, 448, 451, 456, 475, 486, 500, 516, 533, 534 Nuclear Fossa 360, 367, 383-385, 392, 394, 400, 403, 405, 407, 411, 414, 438, 446, 447, 463, 467, 469, 478, 480, 482, 486 Nucleus 45-48, 60, 63, 70, 75, 77, 79, 80, 91, 93, 95, 96, 119, 191, 203, 210, 217, 218, 261, 276, 280-287, 289, 291-293, 295, 297, 299, 300, 307, 309, 310, 323, 324, 326, 328, 329, 332, 342, 343, 350, 355, 360, 365-369, 372-374, 376, 377,

Index 379, 381, 383-385, 388-397, 399-405, 407, 410, 411, 414, 415, 417-423, 425, 426, 428, 429, 431-434, 436, 438-440, 442-450, 452, 454, 455, 458, 459, 460, 461, 463, 467-469, 471, 472-476, 478, 480, 482-486, 488, 490, 491, 493, 494, 497, 500, 501, 502, 532-534, 541, 554, 570, 572-576, 583, 584 Numida 54, 101, 281, 330, 334, 346, 361, 391, 393, 397, 398, 401, 402, 448 Numididae 7, 120, 126, 132, 136, 143, 401 Nuthatch 23, 356, 455 Nyctibiidae 14, 32 Nymphicus 362, 428, 434, 436, 448, 499 O Ocyphaps 354, 359, 362, 418-421, 423 Odontophoridae 7, 120, 126, 132, 136, 143 Old World Flycatchers 23, 24 Old World Sparrows 24 Opisthocomidae 3, 9, 10, 410 Opsin 210, 238 Oriolus 355, 451, 456, 461 Orthonychidae 20 Oscines 17, 19, 20, 24, 25, 28, 35, 358, 363, 423, 425, 445-447, 449, 451, 454, 457, 458, 467, 494, 500, 501 Osprey 11 Ostium 153, 155, 157, 161, 162, 166-168, 173, 201 Ostrich 6, 27, 40, 41, 44-46, 54, 56, 59, 61, 63, 64, 66, 68, 70-73, 75, 80, 87, 89, 93, 96, 98, 101, 103, 109, 112, 116, 117, 124, 125, 193, 199, 225, 235, 285, 287, 289, 292, 293, 295, 297, 301, 311, 314, 334, 341, 344, 345, 354-356, 360, 361, 365, 366, 368, 371-374, 376, 377, 381, 448, 497, 509, 510, 550, 571, 584 Otididae 11, 13, 15, 26, 33 Outer Perivitelline Layer (OPVL) 166, 557 Ovarian Dynamics 244 Ovarian Growth Factors 202 Ovary 111, 149, 159, 161, 162, 165, 167, 181, 200, 201, 229, 241, 243-250, 252256, 258, 259, 266-268, 270-277, 516, 519, 555, 581, 583 Ovenbirds 19, 445

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Overcrowding 319 Oviduct 66, 101, 122, 146, 149, 150, 153, 155, 158, 161-168, 172, 173, 175, 176, 178, 179, 193, 201, 233, 237, 239, 248, 265, 293, 516, 518, 519, 536, 546, 550, 557, 560, 561, 571, 579, 581, 582, 587 Oviposition 166, 183, 188, 193, 201, 262, 265, 270, 557, 559, 560, 565, 580 Ovulation 149, 166, 171, 184, 197, 201, 238, 243, 244, 253, 254, 259, 261-265, 267, 270, 272, 554, 557, 560, 561, 573, 580 Ovulatory Cycle 166, 167, 238, 255, 264, 271, 557, 560 Ovum 149, 166-168, 171, 201, 536, 539, 540, 553-555, 557, 559, 561, 562, 565, 568-581, 583-586 Owlet-nightjars 8, 10, 415 Owls 8, 27, 202, 272 Oxpeckers 24 Oxyruncus 20 Oystercatchers 13 P Palaeognathae 4, 282, 358 Pandionidae 11 Pangolin 350, 560 Panurus 21, 135, 144, 147 Papilla Ductus Deferentis 41, 85, 328 Papillae 134 Paradidymidis 40, 97 Paradiseidae 17, 20, 21 Paramythia 20 Paraphyly 6, 14, 32, 383, 415, 496 Parasitemia 531 Pardalotes 20 Pardalotidae 20 Paridae 17, 21, 29, 30, 358, 363, 449, 450, 457, 467, 485 Parrots 8, 13, 133, 136, 138, 147, 339, 506 Pars Cranialis Uteri 172, 173 Pars Distalis 190, 192, 194, 197, 201, 202 Pars Nervosa 183, 188, 190, 192-194 Pars Recta Ductus Deferentis 39, 41, 85 Pars Translucens 167, 171 Parulidae 25, 32, 358, 363, 467 Parus 17, 232, 252, 363, 447, 449, 450, 457, 485, 486 Passer 86, 103, 203, 219, 224, 225, 228, 231, 236, 237, 239, 283, 298-300, 335,

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Reproductive Biology and Phylogeny of Birds

338, 356, 357, 364, 449, 450, 452, 455, 457, 464, 467, 471, 479-481, 484, 494, 504, 505, 517, 521, 544, 547, 548, 550 Passerculus 523, 549 Passeri (Oscines) 494, 501 Passerida 18-22, 24, 28, 29, 356-359, 363, 392, 425, 447, 448, 451, 458, 461, 463465, 481, 494, 500-502, 532 Passeridae 24, 364, 449, 450, 455, 457, 463, 467, 479, 484 Passeridan Sperm Type 463, 467 Passeriformes 3, 8, 15, 16, 28, 30, 35, 218, 236, 355-359, 362, 427, 444, 458, 494, 498-502, 506, 507, 532, 547, 569 Passerimorphae 500 Passerines 1-4, 15-17, 26, 27, 28, 33, 37, 61, 85-87, 89, 113, 135, 138, 140, 145, 147, 148, 238, 261, 282, 283, 287, 289, 291, 292, 295, 297, 299, 300, 313, 314, 328, 335, 337, 347, 355, 358, 374, 377, 381, 382, 391, 399, 413, 418, 423, 428, 431, 433, 436, 441, 444-448, 451, 456, 458, 461, 475, 480, 486, 490, 492-494, 496, 499-501, 510, 516, 519, 522, 532535, 540, 544, 547, 549, 551 Passeroidea 19, 24, 363, 455, 478, 479 Pathological Polyspermy 168, 553, 559, 561, 570, 571 Pavo 228, 520, 544 Peafowl 520, 544, 569 Pedionomidae 9 Pelecanidae 8, 11, 440 Pelecaniformes 8, 10, 11, 15, 31, 34, 362, 440 Pelecanoididae 11, 13, 15 Pelicans 8, 11, 15, 440 Penduline Tits 21 Penguins 8, 10, 26, 32, 246, 250, 259, 276 Peptides 181, 189, 190, 192, 193, 197, 216, 218, 220, 222, 223, 235, 236, 240 Perforatorium 287, 289, 293, 295, 299, 334, 350, 354, 360, 367, 369, 371-374, 376, 377, 379, 381, 383-386, 389-392, 394, 397, 399-401, 403, 405-407, 410, 411, 413, 415-419, 423, 426, 428, 429, 431-434, 436, 438-440, 443, 450, 478, 480, 494, 496-498, 500, 503, 508, 533 Perisoreus 357, 449, 452, 453, 456, 459 Peritoneal Funnels 150

Persistent Right Oviduct 164, 165, 178 Persisting Right Genital Organs 149 Petrels 11, 13 Petrochelidon 463, 477, 523 Petroicidae 21 Phaethontidae 8-11, 13, 31, 410 Phalacrocoracidae 8, 11 Phalaropes 137, 441 Phalloid Organ 136, 144, 145, 147, 148 Phallus 41, 98, 99, 108, 115-128, 130-134, 136, 142, 143, 146, 147 Phasianidae 7, 15, 120, 132, 136, 143, 358, 383, 387, 396, 400 Pheasants 7, 31, 116, 132, 237 Pheucticus 523 Philetairus 364, 447, 449-452, 455, 456, 473, 479, 482, 483, 493, 502 Philomachus 357, 412, 441, 448, 533 Phoenicopteridae 9, 10, 32, 410 Phoenicopteriformes 8 Phoeniculidae 9, 12, 15 Photoperiodism 203, 205, 224, 231, 236, 237, 250, 270, 276, 316, 317, 337, 341, 344 Photosensitivity 203, 205, 207, 209, 210, 214, 224, 225, 227, 235 Phylloscopidae 23 Phylloscopus 355-357, 457, 463, 465, 478 Phylogenetic Analyses 2, 120 Phylogeny of Spermatozoa 493 Physiological Monospermy 553 Physiological Polyspermy 553, 554, 577 Pica 357, 452, 453, 455, 456, 521 Picathartidae 21 Picidae 12, 14, 15, 35, 358 Piciformes 8, 10, 12, 15, 26, 30, 32, 34, 355, 357, 358, 361, 423, 448, 498-500 Picoides 15, 34, 412 Piculus 15 Picus 15, 355, 423 Pigeons 8, 31, 38, 111, 193, 247, 377 Pine Grosbeak 493 Pineal Gland (Epiphysis) 202 Pinicola 493 Pipilo 364, 447 Pipits 24 Pipridae 17, 20, 358, 445, 446 Piranga 364, 447, 449, 450, 478

Index Pittas 19 Pittidae 19 Pituitary Gonadotropins 243 Plains Wanderer 9 Platycercus 359, 362, 428, 434 Plectrophenax 25 Pleomorphic Sperm 440 Plesiomorphy 2, 365 Plethornithes 8 Ploceidae 358, 449, 450, 467, 479, 482-485 Ploceus 203, 224, 238, 364, 450, 484, 485 Plovers 12, 13, 440 Plumage Ornaments 518 Pluvianellus 13 Pluvianus 13 Podicipediformes 8 Poephila 251, 275, 299, 452, 453, 493 Polioptilidae 23 Polyandry 313, 514, 527, 528, 549 Polygynandrous 145, 148, 504, 527, 544 Polygyny 142, 312, 313, 318, 346, 525, 528, 530, 550 Polypeptides 110, 197 Pomatostomidae 20 Postcopulatory Female Choice 538 Pratincoles 13 Premeiotic Interphase 281 Preleptotene 281 Leptotene 281, 303, 308 Zygotene 281, 308 Pachytene 281, 303, 308 Diplotene 249, 281, 308 Diakinesis 281 Preovulatory Follicle 246, 254, 263, 264, 276, 277 Primary Folds 162, 163, 175 Primary Follicle 252, 254, 255 Primary Spermatocyte 46, 49, 281, 303, 306, 308, 311, 323, 344 Principal Piece 295, 350, 354, 360, 365, 366, 369, 370-374, 379, 381, 383-385, 387-389, 391, 393, 396, 400, 405, 407409, 413, 414, 418, 423, 425, 428, 433, 442, 445, 446, 451, 452, 487, 488, 500, 532 Prionopidae 21 Proacrosomal Granule 283, 285 Procellariidae 11, 13, 15

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Procellariiformes 8, 10, 11 Proctodeum 98, 120, 122, 124, 127, 132, 133 productive demands on testis 312 Progesterone 51, 200, 201, 229, 257-259, 261-264, 273, 274, 276 Prolactin 194, 197, 204, 205, 208, 211, 213, 214, 220, 227-230, 234, 251, 545 Prolactin Releasing Hormone 213 Promerops 24 Promiscuity 135, 142, 147, 303, 313, 315, 517, 519, 521, 527, 528, 536-539, 543 546, 550 Pronephric Tubules 153 Pronephros 150, 153, 155, 156 Pronephrostomes 153 Pronuclei 261, 553, 572-576, 579 Pro-opiomelanocortin Derived Peptides 197 Protein 40, 51, 75, 77, 82, 83, 94, 99, 100, 106, 108, 109, 111, 113, 158, 166, 171, 176, 178, 179, 197, 200, 210, 220, 224, 232-234, 242, 254-259, 261-263, 267270, 272, 273, 275, 292, 315, 336, 342, 343, 377, 554, 555, 557, 559, 565, 567, 568, 570, 571, 580-587 Protonataria 363 Proximal Centriole 360, 367, 369, 372, 373, 377, 379, 380, 385-387, 389, 391, 393, 395, 397, 399, 400, 404, 407, 408, 411, 415, 419,421, 423, 431-434, 436, 439, 442, 472, 480, 483, 494, 502 Prunella 134, 144-146, 335, 364, 493, 504, 513, 527, 544, 548 Prunellidae 24, 364, 493 Pseudophallus 116 Pseudostratified Columnar Epithelium 168, 171, 172, 175 Psittacidae 8, 13-15 Psittacidae/ Cacatuidae 14 Psittaciformes 8, 26, 27, 120, 339, 357, 359, 362, 418, 428, 448, 494, 498, 499, 502, 506 Psittacus 357, 412, 428, 448 Psophiidae 9, 11 Pteroclidae 9, 410 Pterocliformes 8 Pteroicnemia 360 Ptilonorhynchidae 20

$" Reproductive Biology and Phylogeny of Birds Puffbacks 21 Puffbirds 8 Puffins 137, 440, 441 Pycnonotidae 23 Pycnonotus 522 Pyrrhula 451, 457, 490, 493, 504, 533, 544 Q Quelea 328, 340, 341, 364, 450, 484, 522 Quiscalus 364, 481, 486, 487 R Rails 9, 11, 32 Rallidae 9, 11, 13, 32, 356, 358, 438 Ramphastidae 2, 12-15, 33, 358 Ramus ureterodeferentialis cranialis 41 Ratites 4, 6, 66, 99, 115, 117, 146, 193, 297, 352, 354, 355, 371, 374, 381, 383, 387, 393, 405, 418, 428, 431, 436, 443, 494, 496, 509 Rattus 64, 280, 301, 305, 311, 313 Receptaculum Ductus Deferentis 41, 85, 89, 90, 328 Recessus Uterinus 172, 173 Recrudescence 68, 231, 243-245, 251, 254, 317, 319, 320, 327, 330, 332, 521-523, 532 Recurvirostra 519 Recurvirostridae 12, 15, 358, 440 Red Bishop 364, 450, 484, 485 Red-backed Shrike 355, 357, 452, 453, 455, 456, 461, 513, 533 Red-billed Quelea 364, 450, 484, 522 Red-eyed Vireo 449, 457, 461, 526, 536 Red-legged Partridge 209, 226 Red-vented Bulbul 522, 547 Red-winged Blackbirds 147, 520, 550 Reed Bunting 145, 533, 534, 545 Regressed Testes 38, 51, 63, 323 Regression of the Male Duct 158 Regression of the Right Müllerian Duct 150, 158, 159 Regulation of GnIH Expression 222 Regulidae 21 Relationships Between Neoavian Orders 9 Relationships Within Neoavian Orders 11 Relationships Within Passerida 21

Remizidae 21 Reproductive Cycle (see also Testicular Cycles) 40, 51, 86, 101, 214, 265, 270, 271, 315-321, 327, 328, 329, 333, 334, 338, 551 Rete Testis (RT) 44, 54, 61, 63-68, 71, 84, 101-109, 111-113, 305, 328, 329, 330, 333, 335 Retronuclear Body 354 Retzius 349, 355, 356, 371, 382, 405, 411, 413, 417, 419, 421, 423, 425, 426, 428, 438, 440, 441, 443, 448, 452, 453, 456, 458, 459, 461, 463-465, 467, 475, 480, 501, 503, 508, 532, 549 Rhabdornis 24 Rhea 6, 103, 125, 126, 283, 287, 292, 293, 297, 343, 351, 360, 361, 364, 365, 371374, 376, 377, 381, 496, 508 Rheidae 6, 125, 360 Rhinocryptidae 17, 19 Rhipiduridae 21 Rhodopsin 202, 203, 210, 229 Rhynochetidae 9, 11, 410 Ring Dove 210 Riparia 449, 463, 477, 520 Rockfowl 21 Rollers 9, 31 Rooster 38, 41, 44, 46, 49, 51, 54, 56, 61, 64, 66, 70, 73, 75, 80, 82, 83, 87, 90, 9396, 98, 99, 102, 104, 107-109, 111, 134, 280-283, 285, 287, 293, 295, 300-302, 312, 314, 315, 328, 329, 337, 341, 342, 347, 352, 355, 356, 361, 382, 383, 385, 396-401, 403-405, 418, 421, 451, 508, 511 Rough Endoplasmic Reticulum 58-60, 62, 70, 323 Rudimentary Right Oviduct 164 Rudiments of Mesonephros 165 Rudiments of the Wolffian Ducts 162 Ruff 357, 412, 441, 448, 533 Rufous-collared Sparrows 206, 207, 218 S Sagittariidae 11 Sandgrouse 8, 27 Sandpipers 441, 545 Sapayoa 17, 19, 29 Sarcopterygian Fish 350

Index Savannah Sparrows 523, 524, 526, 549 Saxicola 207, 228 Scolopaci 12 Scolopacidae 14, 358, 441, 501, 530, 535 Scolopax 357, 382, 441, 443 Scopidae 8, 440 Screamers 7, 116, 131 Scrub-birds 20 Season 40, 124, 128, 133-135, 138, 141, 147, 158, 162, 166, 184-186, 199, 200, 203, 205, 209, 221, 235, 239, 245, 248251, 253, 256, 266, 267, 312, 316-318, 327, 345, 346, 504, 515, 521, 523, 526, 527, 532, 543 Seasonal Breeding 204, 209, 230, 237, 315, 319, 344 Seasonal Dynamics 220 Seasonally Breeding Species 40 Secondary Folding 162 Secondary Spermatocyte 46, 281, 303,

308 Secretary Bird 11 Secretory Cells 89, 168, 170-172, 175, 194 Secretory Granules 168, 170, 172, 193, 194 Sedge Warbler 448, 457, 478, 517, 544 Semen Extender 561, 577 Seminal Glomera 135, 314, 434, 519 Seminal Glomus or Sac 85, 86, 89, 96, 199, 313, 314, 320, 330, 333 Seminiferous Epithelium 45, 46, 48, 49, 51, 54, 57, 61, 104, 113, 279-282, 301303, 305-307, 308, 311, 312, 321-324, 334, 336, 337, 340, 342-344 Seminiferous Tubule 46-48, 51, 53-57, 5961, 63, 105, 112, 279, 297, 301, 302, 305, 306, 321, 323, 325, 335 Semipalmated Plover 533 Seriemas 11, 208, 235, 239, 364 Serinus 44, 45, 157, 208, 364 Serosa 44 Serotonin 202, 203 Sertoli Cell 46-53, 61, 63, 104-107, 111, 112, 202, 279, 284, 285, 287, 289, 291, 297, 300, 301, 302, 311, 313, 321, 323325, 327, 337, 339, 413, 415, 488, 490, Sexual Selection 116, 126, 137, 141, 145, 176, 335, 539, 540, 544-546, 548, 580, 585

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Sexually Transmitted Diseases 124, 139, 146 Sharpbill 20 Sharp-tailed Grouse 525, 549 Shearwaters 11, 13 Sheathbills 13 Shell 149, 162, 164, 166, 167, 172, 173, 178, 201, 557 Shell Gland 162, 164, 166, 167, 172, 173, 201, 557 Shell Membranes 149, 166, 172 Shoebill 8, 32, 440 Shrikes 21 Silky Flycatchers 23 Sitta 356, 363, 455, 456 Sittidae 23, 358, 363, 455, 467 Skuas 26, 440 Smith’s Longspur 527 Smooth Endoplasmic Reticulum 49, 50, 59, 60, 62, 285, 300, 327, 490, 491 Snow Buntings 25 Social Mating System 134, 514 Society Finch 493 Song Sparrow 198, 207, 218, 238, 241, 251 Song Thrush 357, 448, 449, 451, 457, 463, 467 Songbirds 26, 32, 198, 199, 216, 220, 223225, 252, 549, 550 SOX 161 Specialized Receptors 182, 184 Speculanas 8 Sperm 40, 53, 70, 82, 83, 85-87, 90, 99, 100, 102, 103, 106-110, 113, 115, 116, 122, 124, 131, 135-139, 141-148, 150, 166, 168, 171-177, 225, 295, 300, 303, 313-315, 318, 328, 333-337, 339, 342, 343, 345, 347, 349-352, 354, 355, 357, 360, 365, 367, 369-372, 374, 376, 377, 379, 381, 383, 385-387, 389-394, 396405, 407, 408, 410-413, 415, 417-419, 421, 423, 425-428, 431, 433, 434, 436, 438-447, 449-458, 461, 463, 464, 466468, 472, 473, 475-481, 483, 485-488, 490, 492, 493, 496-510, 513, 514, 516, 518-521, 525-551, 553, 556-587 Sperm Competition 85, 109, 110, 135, 139, 141-148, 176, 225, 313, 314, 335, 342, 343, 451, 490, 504, 513, 514, 516,

$$ Reproductive Biology and Phylogeny of Birds 520, 526, 527-532, 535, 536, 538, 539, 541-550, 580 Sperm Depletion 109, 342, 520, 527, 530, 544, 548 Sperm Head 337, 347, 377, 379, 407, 417, 421, 438, 451, 475, 476, 487, 492, 505, 532, 563, 572 Sperm Longevity 539, 543 Sperm Production 40, 82, 85, 87, 109, 147, 303, 313, 314, 342, 514, 516, 518521, 525-527, 530, 539, 548, 550 Sperm Production Rate 40, 313 Sperm Size 145, 504, 513, 514, 533-536, 539-543, 545, 546 Sperm Storage 102, 113, 136, 141, 146, 148, 150, 171-177, 313, 335, 347, 392, 434, 493, 504, 514, 516, 536, 537, 539, 543-547, 550, 551, 557-561, 580, 582, 584, 587 Sperm Storage Tubules (SST) 557 Sperm Velocity 536 Spermateliosis 111, 282 Spermatid(s) 46-49, 53, 107, 108, 281302, 307, 308, 311, 334, 336-339, 341, 342, 344-346, 351, 352, 411, 414, 417, 425, 429, 442, 454, 456, 459, 461-463, 477, 490, 505, 507-509, 579 Spermatocytes 46, 49, 108, 280, 281, 292, 301, 302, 303, 306, 308, 311, 321, 323, 337, 344 Spermatocytogenesis 279-281, 301, 302, 304, 306-313, 321, 323 Spermatogenesis 40, 46, 49, 51, 61, 104, 106, 108, 109, 112, 197, 198, 279, 280282, 297, 302, 303, 306, 311, 312, 317, 327, 333, 336-338, 340-345, 347, 349, 365, 425, 447, 488, 506, 509, 510, 511, 521 Spermatogenesis, Duration of 311 Spermatogenic Tissue 531 Spermatogonium/ia 46, 47, 48, 49, 50, 51, 53, 108, 279-281, 301, 304, 306-309, 310, 311, 321, 323, 336, 337, 340, 341, 344 Spermatozoon/Spermatozoa 38, 44, 46, 49, 61, 66, 70, 72, 75, 76, 82, 83, 85, 86, 89, 90, 94, 95, 98, 101-104, 107, 109, 112, 113, 199, 247, 279, 282, 289, 291, 292, 297, 299-301, 312-315, 318, 328,

329, 334-336, 337, 339, 342-346, 349, 350, 352-354, 360, 361, 365, 366, 369, 371-374, 376-383, 385, 387-389, 391393, 396-398, 401, 403-405, 407, 408, 410, 415, 417, 418, 420, 421, 424, 426431, 433, 434, 438-444, 447, 451, 452, 455, 456, 458, 460, 464, 465, 467-469, 470-474, 476-480, 484-487, 492-494, 496, 498, 500, 502-511, 513, 514, 532534, 539, 543, 544, 547, 548, 550, 551, 576, 581-586. Spermiation 297, 300, 340, 507 Spermiogenesis 111, 282, 283, 285, 289, 291, 292, 295, 297, 298, 300, 302, 334, 336, 338-340, 343-347, 351, 362, 411, 418, 426, 442, 450, 454, 455, 479, 488, 490, 503-505, 507-511 Spheniscidae 10, 13, 32 Sphenisciformes 8, 10, 26, 362 Sphenodon 350, 352, 353, 354, 381, 436, 505, 506 Sphenodontida 350, 352, 505, 506 Sphenoeacus 23 Spinus 357, 427, 463, 464 Splanchnopleure 157 Spotted Antbirds 207 Spotted Sandpiper 441, 549 Squamates 350, 352, 421 SST 172, 175, 493, 536, 538, 557-562 Starlings 3, 23, 194, 206, 209, 212-214, 224-230, 232, 235, 236, 241, 266, 274, 276, 521, 545 Steamer Ducks 8 Stem Cell 46, 255, 280, 301, 302, 304, 306, 307, 311, 312, 339, 343 Type A (Ad, Ap1, Ap2) 280, 281, 307 Type B 121, 127, 280, 281, 306-308, 310, 311, 326 Steps of Spermiogenesis 283, 298, 299, 302 Stercorariidae 14, 15 Stercorarius 15 Steroid Biosynthesis 200 Steroids 51, 99, 177, 182, 189, 201, 212, 213, 224, 229, 231, 234, 236, 237, 239, 256, 272, 341 Stilts 12, 440, 550 Storage 85, 90, 100, 102, 103, 113, 135, 141, 146, 148, 150, 166, 171-177, 234,

Index 313, 335, 336, 347, 392, 434, 493, 504, 514, 516, 536-539, 543-547, 550, 551, 553, 557, 558, 559-561, 577, 580-582, 584, 587 Storks 3, 9, 34 Storm Petrels 13 Streptopelia 15, 31, 38, 210, 259, 293, 342, 362, 418, 423, 507 Striated Columns 354, 421 Strigiformes 8, 357, 358, 415, 417, 426, 448 Strix 357, 412, 426, 448 Struthidea 21 Struthio 40, 58, 62, 88, 90, 101, 109, 112, 124, 193, 225, 285, 295, 322, 341, 345, 356, 357, 360, 361, 364-366, 368, 383, 448, 451, 496, 509, 510, 550 Struthionidae 360 Struthioniformes 6, 116, 297, 356, 360, 361, 379, 381, 448, 502 Sturnella 487, 533 Sturnidae 3, 23, 24, 358, 363, 450, 455, 463, 473 Sturnus 73, 102, 194, 206, 224-230, 235, 238, 241, 271, 328, 357, 363, 449, 450, 452, 456, 463, 465, 473, 475, 476, 521, 545 Subacrosomal Cone 350, 360, 431 Subacrosomal Space 293, 295, 350, 360, 367, 376, 383-385, 392, 400, 403, 407, 410, 415, 416, 419, 423, 429, 438, 496, 497 Suboscines 17, 19, 29, 30, 358, 362, 444, 445, 447-449, 451, 458, 501, 502 Sugarbirds 24 Sulidae 8, 11 Sunbirds 24 Sunbittern 11 Superb Fairy-wren 87, 527 Supernumerary Male Pronuclei 573, 574 Supernumerary Testes 516, 547 Surface Mucosa 168 Swallow(s) 20, 21, 23, 146, 517, 520, 523, 532, 545 Swifts 8, 12, 27, 38 Sylvia 23, 230, 356, 457, 478 Sylvietta 23 Sylviidae 23, 358, 457, 463, 477

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Sylvioidea 17, 19, 21, 23, 25, 363, 477 Symmetrical Testes 517, 518 Synapomorphies 3, 12, 350, 365, 371, 495, 502 Synapomorphy of Aves 360 Syngamy 569, 573-575 T Tachycineta 134, 363, 449, 450, 455, 478, 517, 534, 547 Tachyeres 8 Tadorna 355, 405, 425 Taeniopygia 225, 230, 251, 262, 299, 313, 364, 451-453, 455, 493, 513, 544, 549, 581 Tail 285, 289, 291, 300, 343, 365, 366, 369371, 374, 376, 401, 403, 407-409, 419, 425, 429, 433, 445, 451, 456, 461, 463, 466, 469, 471, 477, 480, 493, 507, 508, 518, 532, 534, 541 Tanagers 25 Tapaculos 19 Terminal Segment 62, 63, 105, 111 Terns 252, 440 Testicular Asymmetry 103, 105, 517-519, 523, 543, 546, 547, 549 Testicular Capsule 40, 44-46, 65, 104, 321, 323 Testicular Color 38 Testicular Cycles 241, 279 Acceleration (Accelerative) Phase 320, 327, 333 Active Secretory Phase 320 Culmination Phase 320, 327 Presecretion Phase 320, 333 Progressive Phase 320, 333 Reconstruction (Reconstructive, Regeneration) Phase 320, 329 Regressive Phase 320, 329, 526 Reproductive Phase 320, 327, 328, 333 Testicular Spermatozoa 44, 314, 315, 452 Testiculares Accessoria 41 Testis 37-41, 43-46, 51-54, 56, 61, 63-68, 71, 82, 84-87, 89, 96, 97, 101-113, 145, 159, 161, 177, 181, 197, 199, 206, 221, 225, 226, 229, 233, 234, 247, 279, 303, 305, 306, 311, 313, 314, 319-330, 333, 334-342, 344-347, 452, 462, 513-527, 529-532, 535, 536, 541-543, 545-550

$& Reproductive Biology and Phylogeny of Birds Testis Size 38, 40, 109, 141, 145, 199, 212, 338, 342, 513, 516-518, 521-523, 525527, 529-532, 535 , 542, 543, 545, 546, 548, 549, 550 Testosterone 51, 198-201, 221, 228, 239, 262, 263, 317, 318, 327, 338, 342, 343, 346, 516, 529, 532, 546, 549, 550, 555, 585 Tetrao 357, 360, 383 Tetraonidae 7, 15, 358 Tetrapod(s) 350, 352, 354, 436, 506 Thamnophilidae 17, 19, 30 Thermotolerance 40 Thornbills 20 Thraupidae 25, 358, 364, 449, 450, 467, 478 Thrushes 19, 23, 445 Thryothorus 363, 380, 456 Thyroid Hormones 204, 213, 215, 262, 276, 345 Thyroid-stimulating Hormone (TSH) 197 Timaliidae 23, 24 Tinamidae 123, 357, 360, 377 Tinamiformes 6, 116, 147, 361, 377, 379, 381 Tinamous 4, 6, 26, 116, 117, 122-124, 141, 142, 147, 377, 418, 494, 509 Tip Granule 383, 385, 387, 391, 393, 401, 405, 494, 497 Titmice 21, 29, 33, 229 Tityras 20 Tityridae 20 Todidae 9, 12 Todies 9 Totanus 357, 441 Toucans 13, 26 Tragopan 361, 400, 510 Transferrin 53, 113 Transport 61, 68, 71, 80, 82, 85, 93, 103, 106, 111, 150, 166, 176, 192, 206, 247, 256, 257, 261, 313, 315, 335, 337, 503, 553, 557, 559, 560, 580, 581, 585 Tree Swallow 134, 463, 517, 518, 547 Treecreeper(s) 20, 23, 455, 478 Treeswifts 12 Tringa 357, 382, 441, 443 Trochilidae 14, 26, 358, 498 Troglodytes 363, 455, 478, 479, 521

Troglodytidae 23, 26, 358, 363, 455, 467, 478 Trogon 426, 427 Trogonidae 10, 14, 30, 32, 358 Trogoniformes 8, 28, 33, 358, 405, 426, 427 Trogons 8, 10, 30, 32 Tropicbirds 8, 11, 31 Trumpeters 9, 11 Tubal Ridge 153 Tubenoses 8 Tubular Glands 162, 166, 168, 171, 172, 175 Tubular Neck Region 167, 168 Tubular Shell Gland 167, 172, 173 Tubuli 63, 97, 103, 105 Tubulus Rectus 63 Tufted Titmouse 363, 449, 450, 485 Tunica 44, 45, 175 Tunica Albuginea 44 Tunica Muscularis 175 Turacos 9, 10, 34 Turdidae 23, 24, 358, 363, 449, 450, 457, 467 Turdus 165, 357, 363, 447-452, 454, 456, 457, 463, 464, 467-472, 477, 480, 493, 522, 534, 549 Turnicidae 9, 11, 15 Tympanuchus 525 Tyranni 19, 20, 358, 444, 445, 449, 501 Tyrannida 444 Tyrannidae 17, 20, 358, 427, 444-446, 455 Tyrannus 362, 427, 444-447, 449, 456, 501, 502 Tyrant Flycatchers 20, 445, 446 U Undulating Membrane 380, 425, 445-447, 454, 488, 533 Unilateral Development of Female Genital Organs 149 Unit 61, 63, 64, 71, 72, 82, 83, 85, 87, 9092, 94, 95, 100, 181, 183, 187, 188, 315, 328-330, 333 Upper Neck Region 168 Upupidae 9, 12, 15 Ureterodeferentiales Caudales 41 Uria 356, 382, 440, 504 Urodeles 354

Index Urodeum 41, 85, 120, 175, 328 Uterine Transudate 166 Utero-vaginal Junction (UVJ) 172, 557 Uterus 162, 166, 167, 172, 173, 175, 201, 557, 559, 574 V Vaginal Sphincter 175 Vanellus 355, 357, 382, 440, 443 Vangas 21 Vangidae 21 Vasculosa 44, 45 Vasectomy 83, 104, 105 Vasoactive Intestinal Polypeptide 213, 227, 230 Veniliornis 15 Vimentin 72 Violet-green Swallow 363, 449, 450, 455, 472, 534 Vireo 17, 363, 427, 447-449, 451, 454, 457, 458, 461, 463, 526, 536 Vireonidae 358, 363, 427, 457 Vitellogenesis 200 Vitellogenin (VTG) 257 VTG/VLDL Receptor 257 Vultur 358 W Waders 530, 533, 540 Warblers 17, 23, 25, 518, 546 Waterfowl 31, 32, 34, 116, 131, 145, 261, 527, 528, 545 Wave of the Seminiferous Epithelium 305, 343 Waxwings 23 Weight of the Egg 149, 259 Western Meadowlark 487, 533 White-bellied Yuhina 17 White-crowned Sparrow 187, 191, 194, 195, 217, 218, 221, 225, 228, 229, 233, 234, 240-242, 274, 319, 342

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White-rumped Mannikin 493 White-rumped Munia 493 White-winged Chough 21 White-winged Fairy-wren 519 Wild Birds 40, 68, 102, 115, 127, 150, 165, 244, 249, 318, 319, 335, 519 Willow Warbler 457, 478 Winter Wren 521 Wolffian Duct(s) 85, 94, 97, 99, 104, 109, 150, 153, 155, 157, 162, 163, 165, 166, 176, 177, 315, 337, 342 Wolffian Duct Proteins 94, 99, 109, 315, 342 Woodhewers 445, 446 Woodhoopoes 9 Woodpeckers 8, 28, 33, 34 Wood-swallows 20 Wren 17, 20, 23, 87, 134, 363, 380, 381, 455, 456, 478, 479, 519, 521, 527 X Xenopus 555, 575, 583, 587 Y Yellow Hammer 457, 490 Yellow Warbler 391, 513, 534 Yuhina (Erpornis) 17 Z Zebra Finch 193, 230, 251, 275, 299, 300, 313, 335, 364, 451-453, 493, 504, 513, 519, 534, 536, 544, 569, 581 Zona Pellucida (ZP) 555 Zonotrichia 187, 191, 195-199, 201, 206208, 210-212, 215, 217, 218, 225, 228, 229, 232-234, 236, 240-242, 250, 256, 258, 274, 319, 337, 340, 342, 347, 364, 447 Zosteropidae 23 Zosterops 23