Current Topics in Developmental Biology, Volume 34

Current Topics in Developmental BioIogy Volume 34

Series Editors Roger A. Pedersen

and

Reproductive Genetics Division Department of Obstetrics, Gynecology, and Reproductive Sciences University of California San Francisco, CA 94143

Gerald P. Schatten Department of Zoology University of Wisconsin, Madison Madison, W I 53706

Editorial Board Peter Grijss Max-Planck-Institute of Biophysical Chemistry, Gottingen, Germany

Philip lngham Imperial Cancer Research Fund, London, United Kingdom

Mary Lou King University of Miami, Florida

Story C. Landis National Institute of HealthiNational Institute of Neurological Disorders

and Stroke, Bethesda, Maryland

David R. McClay Duke University, Durham, North Carolina

Yoshitaka Nagahama National Institute for Basic Biology, Okazaki, Japan

Susan Strome Indiana University, Bloomington, Indiana

Virginia Walbot Stanford University, California

Founding Editors A.A. Moscona Alberto Monroy

Current Topics in Developmental Biology

Edited by

Roger A. Pedersen Reproductive Genetics Division Department of Obstetrics, Gynecology, and Reproductive Sciences University of California, San Francisco San Francisco, California

Gerald P. Schatten Department of Zoology University of Wisconsin, Madison Madison, Wisconsin

Academic Press San Diego

London

Boston

New York

Sydney

Tokyo

Toronto

Front coverphorograph: The bacterium C. crescentus. From Chapter 6 by Roberts et al. For details see legend to Fig. I .

This book is printed on acid-free paper.

@

Copyright 0 1996 by ACADEMIC PRESS All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher.

Academic Press, Inc.

525 B Street, Suite 1900, San Diego, California 92101-4495, USA http://w.apnet.com United Kingdom Edition published by Academic Press Limited 24-28 Oval Road, London NWI 7DX, UK http://w.hbuk.co.uk/ap/ International Standard Serial Number: 0070-21 53 International Standard Book Number: 0-12-153134-1 PRINTED IN THE UNITED STATES OF AMERICA 96 97 9 8 9 9 00 01 EB 9 8 7 6 5

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3

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1

Contents

ix

Contributors

Preface

xi

1 SRY and Mammalian Sex Determination Andy Greenfield and Peter Koopman

I. Introduction 1 The SRY Gene 2 The SRY Protein 7 Other Sex-Determining Genes General Conclusions 17 References 18

11. 111. IV. V.

13

2 Transforming Sperm Nuclei into Male Pronuclei in Vivo and in Vifro D. Poccia and P. Collas

I. 11. 111. IV. V. VI. VII.

Introduction 26 Changes in Nuclear Proteins 32 Chromatin Decondensation 41 Formation or Adjustment of Nucleosomes 52 Nuclear Envelope Disassembly and Assembly 55 Male Pronuclear Activities 74 Conclusions 78 References 79

3 Paternal Investment and lntracellular Sperm-Egg Interactions during and Following Fertilization in Drosophila Timothy L . Karr I. Introduction

89

11. Sperm Structure and Production in Drosophila

92

vi

Contents

94 111. Sperm Transfer, Storage, and Utilization IV. Syngamy (Sperm Penetration), Pronuclear Maturation, Migration, and Karyogamy 94 V. Structural Analysis of a “Sperm-Derived Structure” in the Developing Zygote 98 VI. Genetics and Molecular Biology of Fertilization and Early Embryonic Development in Drosophilu 100 VII. Cytoplasmic Incompatibility 103 107 VIII. Speculative Models of Sperm Function in the Fertilized Egg IX. Conclusions and Perspectives 111 References 112

4 Ion Channels: Key Elements in Gamete Signaling Albert0 Darszon, Arturo M v a n o , and Carmen Beltrdn

I . Why Are Ion Channels Important in Fertilization? 11. 111. IV. V. VI. VII.

117 Gamete Generalities 118 Influence of the Ionic Environment on Spermatozoa 121 124 Long-Range Communication between Gametes Short-Range Communication between Gametes: The Acrosome Reaction 144 Do Ion Channels lhm the Egg On? Concluding Remarks 148 References 149

5 Molecular Embryology of Skeletal Myogenesis ludith M. Venuti and Peter Cserjesi

I. Introduction

169

11. MyoD Family of Myogenic Basic-HLH Factors (mHLHs) 111. Developmental Expression of mHLHs 175

IV. V. VI . VII . VIII.

171

mHLH Factors in Invertebrate and Nonmammalian Vertebrates 182 Mutational Analysis of mHLH Function 188 Early Activation of the Myogenic Program 194 MEF2 Family of Transcription Factors Summary and Conclusions 198 References 199

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Contents

6 Developmental Programs in Bacteria Richard C. Roberts, Christian D. Mohr, and Lucy Shapiro 207 I. Introduction: The Concept of Development among Bacteria 11. Some Examples of Development among Bacteria 209 111. Control of Cellular Differentiation during the Cuulobucter crescerzfusCell Cycle 226 245 IV. Conclusions and Future Perspectives References 246

7 Gametes and Fertilization in Flowering Plants Darlene Southworth 1. Introduction 259 11. Male Gametes 260 111. Female Gametes 271 IV. Double Fertilization 273 V. Summary 275 References 276

Index

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This Page Intentionally Left Blank

Contributors

Numbers in parentheses indrcate the pages on which the authors' contributions begin

Carmen Beltran (1 17), Departamento de GenCtica y Fisiologia Molecular, Instituto de Biotecnologia, Universidad Nacional Autdnoma de MCxico, Cuernavaca, Morelos 6227 1, Mexico P. Collas' (25), Department of Food Science, Agricultural University of Norway, As, Norway Peter Cserjesi (169), Department of Anatomy and Cell Biology, Columbia College of Physicians and Surgeons, New York, New York 10032 Albert0 Darszon (117), Departamento de Genttica y Fisiologia Molecular, Instituto de Biotecnologia, Universidad Nacional Autonoma de Mtxico, Cuernavaca, Morelos 6227 1, Mexico Andy Greenfield (I), Centre for Molecular and Cellular Biology, The University of Queensland, Brisbane, Queensland 4072, Australia Timothy L. Karr (89), Department of Organismal Biology and Anatomy, University of Chicago, Chicago, Illinois 60637 Peter Koopman (l), Centre for Molecular and Cellular Biology and Department of Anatomical Sciences, The University of Queensland, Brisbane, Queensland 4072, Australia Arturo Lievano (1 17), Departamento de GenCtica y Fisiologia Molecular, Instituto de Biotecnologia, Universidad Nacional Autonoma de MCxico, Cuernavaca, Morelos 6227 1, Mexico Christian D. Mohr (207), Department of Developmental Biology, Stanford University School of Medicine, Stanford, California 94305 D. Poccia (25), Department of Biology, Amherst College, Amherst, Massachusetts 01002 Richard C. Roberts (207), Department of Developmental Biology, Stanford University School of Medicine, Stanford, California 94305 Lucy Shapiro (207), Department of Developmental Biology, Stanford University School of Medicine, Stanford, California 94305 'Present Address: Department of Biochemistry, Norwegian College of Veterinary Medicine, 0033 Oslo, Norway. IX

X

Contributors

Darlene Southworth (259), Department of Biology, Southern Oregon State College, Ashland, Oregon 97520 Judith M. Venuti (169), Department of Anatomy and Cell Biology, Columbia College of Physicians and Surgeons, New York, New York 10032

Preface

The field of developmental biology is fabulous for many reasons, and perhaps foremost among the many strengths of our discipline is its inclusiveness. This volume highlights this aspect: An international group of dynamic scientific contributors explores a range of developing systems wider than those included in the classic dogma, using a wealth of experimental protocols benefitting from molecular, cellular, genetic, and biophysical approaches. Although developmental biology has typically been viewed as the exclusive purview of eukaryotes, the chapter entitled “Development Programs in Bacteria” by Richard C. Roberts, Christian D. Mohr, and Lucy Shapiro undermines the foundation of this narrow perspective. Determination and differentiation during development are major questions now being addressed most successfully, and the article by Andy Greenfield and Peter Koopman examines the exciting topic of “SRY and Mammalian Sex Determination.” “Molecular Embryology of Skeletal Myogenesis” by Judith M. Venuti and Peter Cserjesi admirably reviews this lively example of a different type of differentiation. The molecular mechanisms of fertilization emerge as a subtheme in this otherwise eclectic volume of Current Topics in Developmental Biology. The article by Darlene Southworth on the “Gametes and Fertilization in Flowering Plants” reminds us of the wealth of research opportunities in these fascinating systems. Albert0 Darszon, Arturo LiCvano, and Carmen Beltran consider what is perhaps the earliest of events during the fertilization process in their chapter “Ion Channels: Key Elements in Gamete Signaling.” Because the goal of fertilization is the union of the parental genomes within the activated egg, the emergence of the male pronucleus is a critical event; this is considered by Dominic Poccia and Philippe Collas in their article “Transforming Sperm Nuclei into Male Pronuclei in Vivo and in Vitro.” The precise nature of the parental contributions to the zygote and the embryo is still not completely understood, and the contribution by Timothy L. Karr examines the “Paternal Investment and Intracellular SpermEgg Interactions during and Following Fertilization in Drosophila. ” Together with other volumes in this series, this volume provides a comprehensive survey of major issues at the forefront of modem developmental biology. These chapters should be valuable to researchers exploring development in plant, animal, and now prokaryotic systems, as well as students and other professionals who want an introduction to current topics in cellular, molecular, genetic, and biophysical approaches to developmental biology. This volume in particular will xi

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Preface

be essential reading for anyone interested in sex determination, reproduction, fertilization, inheritance, cell-cycle regulation, ionic signaling, embryo formation, morphogenesis, muscle development, and differentiation. This volume has benefitted from the ongoing cooperation of a team of participants who are jointly responsible for the content and quality of its material. The authors deserve full credit for their success in covering their subjects in depth yet with clarity and for challenging the reader to think about these topics in new ways. We thank the menbers of the Editorial Board for their suggestions of topics and authors. We thank Liana Hartanto, Heather Aronson, and Diana Myers for their exemplary administrative and editorial support. We are also grateful to the scientists who prepared articles for this volume and to their funding agencies for supporting their research. Gerald P. Schatten Roger A . Pedersen

1 SRY and Mammalian Sex Determination Andy Greenfield’ and Peter Koopman1I2 ‘Centre for Molecular and Cellular Biology and 2Departnient of Anatomical Sciences The University of Queensland, Brisbane Queensland 4072. Australia I. Introduction 11. The SRY Gene A . Gonadogenesis B. Timing and Tissue Distribution of SRY Transcription C. The Structure of SRY Transcripts

D. Conclusions 111. The SRY Protein A. The High Mobility Group (HMG) Box B. DNA-Binding Properties of SRY C. SRY: A Transcriptional Activator? D. Conclusions IV. Other Sex-Determining Genes A. Mullerian Inhibitory Substance (MIS) B . Steroidogenic Factor 1 (SF-I) and WT-I

c. sox9

D. Other Loci-DSS and Tus V. General Conclusions References

1. Introduction Two facts feature in any discussion of the genetic basis of sex determination in eutherian mammals. First, the development of an embryo into a male or a female is dependent on whether the bipotential embryonic gonad differentiates into an ovary or a testis. Once the “choice” of the ovarian or testicular pathway of development has been made, all subsequent sexually dimorphic characteristics are the consequence of the hormonal output of the gonads (Jost, 1947). Second, the mammalian Y chromosome is a dominant determinant of maleness (Ford et al., 1959; Jacobs and Strong, 1959; Welshons and Russell, 1959). In normal circumstances, it is the presence or absence of the Y chromosome, not the number of X chromosomes, which determines the sex of the individual. Taken together, these two central tenets allow the geneticist to frame the question of the genetic basis of sex determination in mammals in terms of which gene or genes on the Y chromosome are required for the initiation of testis development. The isolation of the predicted gene(s), known as the testis determining factor (TDF) Currrnr Topics tn Dcvelopmrnral Biology. V i l 34 Copyright 0 1996 by Academic Press, Inc All rights of reproduction In any lorn reserved

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Andy Greenfield and Peter Koopman

in humans and testis-determining Y gene (Tdy) in mice, was the subject of an intense international research effort which culminated in 1990 with the identification of the human SRY gene (Sinclair et a l . , 1990). A search through a 35-kb region of the human Y chromosome, the minimum known to be necessary for male sex determination, resulted in the identification of a gene exhibiting all the predicted properties of TDF: it was conserved on the Y chromosome of all mammals tested, it was expressed in the developing male gonad prior to overt testis differentiation, and it encoded a DNA-binding protein of obvious regulatory potential. Final proof of the identity of SRY and TDF came in 1991 in the form of a chromosomally female mouse transgenic for the murine Sry gene: this mouse developed as a normal male, albeit sterile due to the presence of two X chromosomes and the absence of Y chromosomal genes required for spermatogenesis (Koopman et al., 1991). Sry was thus shown to be the only Y-linked gene (though by no means the only gene) required for testis determination in mammals. Six years have now passed since the identification of SRY. The details of the search for TDFITdy have been documented elsewhere (Goodfellow et al., 1993). This review will focus on our current understanding of the biology of SRY and sex determination. We shall pay particular attention to what is known of the biochemical basis of SRY function and its relationship to other genes in the sex determination pathway. Most importantly, we shall attempt to identify those areas in which our ignorance is greatest and address some of the issues which might concern researchers in sex determination over the next 6 years. Data reviewed here will be primarily from studies of mice and humans; the symbol “Sry” will be used to refer exclusively to the murine gene and “SRY” to that of humans and other mammals.

II. The SRY Gene A. Conadogenesis

It is clear from the above introduction that the function of the SRY gene is known: its activity results in the development of a testis from the bipotential embryonic gonad. Precisely how this result is achieved, however, is unclear. Before discussing what is known of the structure and expression of the SRY gene, we shall briefly review the cellular basis of gonadogenesis in the eutherian mammalian embryo to set the scene for more detailed discussion of SRY function. In the mouse, the gonad has its origins in the genital ridge, a structure which arises as a thickening of the mesonephros at about 10.5 days post coitum (dpc). The mesonephros and genital ridge together are known as the urogenital ridge. The developing gonad comprises four known cell lineages common to both males and females: the germ cells and at least three somatic cell types, steroid

I . SRY and Mammalian Sex Determination

3

cells, supporting cells, and connective tissue. Primordial germ cells migrate from their origin in the primitive streak along the dorsal mesentery and arrive in the genital ridge between 10.5 and 12 dpc. The somatic portion of the genital ridge is derived from mesenchyme and overlying coelomic epithelium, as well as from cells which migrate from the adjacent mesonephros (Buehr et a l . , 1993; Upadhyay et al., 1981). The supporting cell lineage is the first to exhibit sexspecific differentiation. In males, this lineage produces the Sertoli cells at around 12.5 dpc, while in females it differentiates into the ovarian follicle cells. Alignment of the Sertoli cells results in the formation of characteristic testis cords. The first known product of the Sertoli Cells is Mullerian inhibitory substance (MIS), also known as anti-Mullerian hormone, a glycoprotein which causes regression of the female reproductive tract anlagen. The other primary testicular hormones are testosterone and dihydrotestosterone, which are produced from the male derivatives of the steroid cell precursors, the Leydig cells. These promote the development of the Wolffian duct system into the epididymis, vas deferens, and seminal vesicles. It is the absence of these testicular hormones which results in the “default” ovarian pathway of development. No female counterparts to these hormones, required for development of the female genitalia, have been identified. If asked to predict, from the above discussion, the timing and sites of Sry activity during testis determination and differentiation, one would likely say between 10 and 12 dpc in the developing male gonad. In addition, if we were to predict a precise function for S r y , it would likely be in determining the fate of one or more of the gonadal cell lineages discussed above, possibly including the direct activation of genes encoding or regulating the production of the testicular hormones.

B. Timing and Tissue Distribution of SRY Transcription

When expression of murine Sry was analyzed using the sensitive RT-PCR method, transcripts were detected in adult testis and 11.5 dpc male urogenital ridge samples (Gubbay et al., 1990). More detailed analysis showed that fetal expression is confined to gonadal tissue, does not require the presence of germ cells, and is limited to the period in which testes begin to form (Koopman e t a / . , 1990; Hacker el al., 1995; Jeske et al., 1995). A semiquantitative RT-PCR assay of urogenital ridge RNA samples reveals a profile of Sry transcription starting at 10-10.25 dpc, reaching a peak at 11.25-12 dpc, and ceasing by 13.5 dpc (Jeske et al., 1996). This profile shows a striking correspondence with the key events in testis differentiation and the narrow time window in which transcripts can be detected is likely to reflect the importance of the timing of S t y expression. Further evidence that the timing of Sry expression is critical comes from the observation that when the Mus musculus domesticus-derived M u s poschiavinus

4

Andy Greenfield and Peter Koopman

Y chromosome is placed on a C57BL/6 ( M u s tnusculus musculus) background, XYpos females and hermaphrodites are produced (Eicher and Washburn, 1986; Eicher et al., 1982). This is an example of a more general phenomenon known as B6.YD0m sex reversal. The B6lY-S sex reversal has been interpreted as the result of “precocious” C57BL/6 ovarian-determining genes overriding the masculinizing effects of the YPos Sry allele (Eicher and Washburn, 1986). This explanation supports the idea that the timing of Sry expression is normally sufficiently fine-tuned to preempt any default ovarian differentiation, a fine tuning which can be disrupted in a composite genomic environment. Indeed, differences in the timing of testis differentiation in the presence of YPOS and YC57Bf-16 are well documented (Palmer and Burgoyne, 199 lb). Interestingly, Sry transcription continues beyond the normal period in at least one case of B6.YDom sex reversal (Lee and Taketo, 1994). (See Section 11, part C for further discussion of the role of S l y in B6.YDOmsex reversal.) Sry transcripts are detected exclusively in the genital ridge portion of the murine urogenital ridge (Jeske et al., 1995). What is known of the cell types which transcribe Sty? Testis development can proceed normally in mutant mice lacking germ cells (McLaren, 1985). (This situation contrasts with that found in females, where germ cells are required for the proper development of ovaries). This leaves the three somatic lineages as candidates. The best clues come from the analysis of the cellular make-up of testes in XX ++ XY chimeric mice (Burgoyne et al., 1988; Palmer and Burgoyne, 1991a). While the Leydig cell population in such chimeras displays equal contributions from XX and XY cells, Sertoli cells are almost exclusively XY. It seems likely, therefore, that Tdy acts in the supporting cell lineage and that it is the differentiation of pre-Sertoli into Sertoli cells which constitutes the only cell-autonomous action of Tdy. These data suggest, therefore, that Sry expression in Sertoli cell precursors is the initial step in a commitment to the testicular pathway. In addition, it would appear that Sertoli cells are central in directing the other gonadal cell lineages toward the testicular pathway. The recruitment of a few XX cells to the pathway, however, suggests that some cell-cell interaction is required even in the initial step of testis determination and differentiation. Does a function for SRY exist besides the determination of gonadal sex? Expression studies in humans and marsupials suggest this possibility. Expression of SRY has been detected by RT-PCR in a wide range of human fetal tissues and in adult heart, liver, and kidney, as well as testis (CICpet et al., 1993). Whether these nongonadal transcripts are translated is unknown. A similar picture emerges in the case of fetuses from the marsupial Macropus eugenii, which exhibit widespread transcription of SRY (Harry et al., 1995). However, the absence of such widespread expression in the mouse fetus and the lack of any phenotype other than sex reversal and gonadal dysgenesis in human XY females harboring mutations within SRY argue against a function for SRY beyond gonadal development.

I , SRY and Mammalian Sex Determination

5

Curiously, Sry transcripts have been detected both in mouse (Zwingman et al., 1993) and human (Fiddler et a / . , 1995) preimplantation embryos, and while it has been proposed that this expression may be related to the faster growth of male preimplantation embryos relative to females, it is not known to have any biological significance. C. The Structure of SRY Transcripts

Defining the structure of the SRY transcript is important for our understanding of how the gene is regulated and in determining the structure of the sex-determining SRY protein. The mouse Sry locus is known to be capable of producing at least three distinct transcripts, but data obtained from studies on the predominant transcript found in adult testes caution against attributing a function to all of these. The adult testis is the most amenable tissue for the analysis of Sry expression given the rarity of the embryonic transcript and the minute amounts of RNA which can be extracted from a single dissected urogenital ridge. Expression in the adult testis begins between 2 1 and 28 days post parturn, coincident with the appearance of round spermatids, and is germ cell dependent (Rossi et al., 1993). Northern analysis of adult testis RNA reveals a transcript of approximately 1.3 kb (Koopman et al., 1990). Attempts to isolate an Sry cDNA from embryonic and adult testis cDNA libraries have been unsuccessful, but clones were recovered from a cDNA library derived from RNA extracted from COS cells transfected with Sry (Cape1 et al., 1993). Analysis of these clones, in conjunction with 5' RACE-PCR, RNase protection experiments, and RNase H digestion studies revealed the major adult testis transcript to be circular and nonpolyadenylated. It is believed that this transcript structure arises as a consequence of the organization of the murine Sry locus, which consists of 2.8 kb of unique sequence flanked by two inverted repeats of at least 15 kb (Gubbay er at., 1992) (Fig. 1). The existence of a promoter within the first direct repeat would result in the production of a transcript containing homologous sequences at both ends, facilitating the formation of a hairpin loop structure. This structure could then be resolved into a circle by a normal splicing reaction utilizing conveniently situated splice donor and acceptor motifs. The circular transcript has no known function. It has no known equivalents outside of the Mus species and it is not substantially bound to polysomes; it is unlikely, therefore, to be translated. The majority of Sry transcripts in the genital ridge are not circular. suggesting the existence of a genital ridge promoter distinct from that used in the adult testis during circle formation, and residing within the unique sequence portion of the gene. Defining the structure of this transcript has proven difficult. Traditional approaches such as cDNA library screening have yielded no full-length clones, and researchers have resorted to RACE-PCR and RNase protection experiments

6

Andy Greenfield and Peter Koopman Splice acceptor HMG t+Y box

a. Srylocus

< < < a < 15kb)

b. Genital ridge transcript

c. Adult testis transcript

I I

Splice donor

Y

>>>>>>> Inverted repeat - 3’arm (>15kb)

Unique sequence region (2.8kb)

T

A A A Open reading frame

fftt---y--13

Y

A

A

A

... /

/

/

+ Stem-loop structure

Circular RNA Splicing

Fig. 1 Structure of the mouse S l y gene and its transcripts. (a) Schematic representation of the mouse S l y locus, which consists of 2.8 kb of unique sequence surrounded by at least 15 kb of inverted repeat sequence on either side. The open reading frame contains the HMG box and splice acceptor and donor sequences (Y)5’ and 3’ of the HMG box, respectively. (b) Transcript of Sry generated in the genital ridges. This transcript is initiated from a promoter within the unique sequence region (0).consists of a single exon, terminates within the 3’ arm of the inverted repeat, and is polyadenylated. It contains a 1.2-kb open reading frame with the HMG box at its 5’ end. (c) Transcript of Sry generated in mouse adult testes. This transcript is initiated form a different promoter, within the 5’ arm of the inverted repeat (0)and terminates within the 3’ arm of the inverted repeat. The exact start and end points of this transcript have not been determined, and it is not known whether this transcript is potyadenylated. Base pairing between the repeated ends of this transcript is likely to generate a stem loop structure, allowing the splice donor to splice onto the splice acceptor sequence, thus generating a circular RNA molecule of about 1.4 kb. Not to scale.

to determine the precise start site(s) and polyadenylation site(s) (Hacker et al., 1995; Jeske et ul., 1995). The predominant genital ridge transcript is a 4.5-kb linear molecule (Hacker et al., 1995), though a rarer transcript of 3.5 kb is also detectable, which terminates upstream of the major transcript polyadenylation site (Jeske et al., 1995). Both transcripts consist of a single exon with an 1185bp open reading frame (ORF; Fig. 1). The predicted S r y protein is 395 amino acids long and contains, along with an HMG box DNA-binding motif (see Section 11), a 223-residue glutarnine-rich region at its C terminus. The possible functional significance of this region of the polypeptide is discussed in Section 11. There is evidence for only one type of human SRY transcript: a linear, singleexon, polyadenylated molecule of approximately 1 kb in length, encoding a

1 . SRY and Mammalian Sex Determination

7

predicted protein of 204 amino acids (Behlke et al., 1993; Clepet et a l . , 1993; Su and Lau, 1993; W a i n et ul., 1992). Several transcription start sites have been reported, possibly reflecting differences in the source material. The polyadenylation site is consistently reported as being 133 bp after the stop codon that ends the ORF. The structure of the sex-determining genital ridge transcript has not been determined in humans due to the lack of tissue availability.

D. Conclusions

A picture of the role of Sly in testis determination emerges from this preliminary discussion which can be summarized as follows: at a critical time of around 10 dpc, cells of the supporting cell lineage in the bipotential, embryonic male gonad begin to transcribe Sry. This transcription is initiated by factors produced by the autosomes and/or X chromosome which are likely to be present in both sexes. This linear, polyadenylated message is produced for approximately 3 days and then production ceases, by which time the supporting cells have differentiated into Sertoli cells and these have aligned into the characteristic testis cords. It is not known when commitment to the male pathway occurs in the pre-Sertoli cells nor what factors or signals apart from cell-autonomous Sry activity are involved in this masculinization. What is clear is that Sry is the only Y chromosome gene required to set the ball rolling. The next section reviews what is known about the structure, function, and evolution of the SRY protein.

111. The SRY Protein A. The High Mobility Group (HMG) Box

Sequence analysis of the mouse and human SRY open reading frames revealed a 237-bp region of homology to a protein motif known as the HMG box (Gubbay et al., 1990; Sinclair et al., 1990).This motif is found in a large number of DNAbinding proteins composing the HMG box superfamily (Laudet et al., 1993; Ner, 1992). The HMG box was first recognised as a region of homology between HMG nonhistone proteins such as HMGl and hUBF. SRY belongs to a subclass of HMG box-containing proteins which bind to specific DNA sequences with high affinity and are likely transcriptional regulators, such as TCFl (van de Wetering et al., 1991) and LEF-1 (Giese et al., 1991; Travis et al., 1991). Indeed, the isolation of the SRY gene allowed the identification of a new family of HMG box genes expressed during mammalian development known as SOX genes (for SRY-related HMG box genes) (Gubbay et al., 1990; Denny et al.,1992; Wright et al., 1993). The HMG box of SRY is 79 amino acids long and mediates DNA binding,

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Andy Greenfield and Peter Koopman

interacting with the DNA double helix across the minor groove (van de Wetering and Clevers, 1992). This DNA-binding capacity is critical to SRY function: several cases of human XY sex reversal are the direct consequence of mutations within the HMG box of SRY (Berta et al., 1990; Harley et a l . , 1992; Jager et al., 1990; Nasrin et a l . , 1991). SRY binds to the consensus sequence AITAACAAT/A, as defined by binding site selection experiments (Harley et a l . , 1994). Available evidence suggests that SRY functions by binding to particular sites in the genome and, extrapolating from data concerning known transcription factors such as TCF- I , affects the expression of one or more “downstream” genes in the testis-determining pathway by so doing. As would be predicted for a putative transcription factor, data demonstrate sequestering of SRY protein in the nucleus of the cell, a localization dependent on a highly conserved nuclear localization signal in the amino terminus of the HMG box domain (Poulat et al., 1995). Downstream target genes have yet to be identified, nor is it known whether SRY activates or represses their activity, though the recessive inheritance of certain forms of human XX sex reversal suggests SRY might repress the activity of a repressor of testis determination (McElreavey et a l . , 1993). Recent studies on the in vitro properties of mouse and human SRY protein do not provide a clear answer to the question of whether SRY acts as activator or a repressor and even raise the possibility of species-specific differences in the mechanism of SRY activity. First, we shall look at what is known of the DNA-binding properties of SRY.

B. DNA-Binding Properties of SRY

The model of SRY binding to specific target sequences in order to effect transcriptional control must accommodate two observations. First, SRY exhibits low binding specificity. This can be seen from the number of substitutions in the target site which still allow binding of SRY (Harley et al., 1992). Second, an abundance of proteins exist which share an identical or near-identical sequence specificity, e.g., other members of the Sox protein family (Denny et al., 1992). How is functional specificity achieved in vivo? One possible answer involves the idea that specificity is achieved by proteinprotein interaction. Thus, SRY may bind to a particular target only in the presence of other, presumably tissue-specific, proteins in a nucleoprotein complex. This would involve the contact between one or more proteins and specific SRY residues, likely to reside outside the HMG box. This model predicts the occurrence of mutations at just such residues in cases of sex reversal. However, mutations in SRY associated with human sex reversal have been detected only in the HMG box (Berta et a l . , 1990). Moreover, comparison of SRY proteins from different species reveals a lack of conservation outside the HMG box (Tucker and Lundrigan, 1993; Whitfield et a l . , 1993). We shall return to this lack of conserva-

1 . SRY and Mammalian Sex Determination

9

tion later and discuss its implications for a single model of SRY function in mammalian sex determination. Recent studies of SRY-DNA complexes offer a different answer to the specificity problem, suggesting that SRY acts as a transcriptional modulator via effects on DNA geometry. Human SRY has been shown to distort DNA upon binding, inducing the double helix to bend some 80" and studies on binding of SRY to the enhancer of CD3e show the bending to be centered on the recognition sequence GAACAAAG (Ferrari et al., 1992). Subsequent studies on the binding of mutant human SRY proteins from sex-reversed individuals show that SRY's DNA-binding and -bending activities can be separated and, in addition, that DNA bending is required for SRY function (Pontiggia et al., 1994). One mutant SRY protein studied exhibited a 100-fold reduction in binding affinity but did not affect DNA bending at all. A second mutant protein showed only a 3-fold reduction in binding affinity, but reduced the angle of bending from 75 to 56". Perhaps most importantly, target sites with altered sequences were observed to bind SRY, but the angle of bend induced varied from site to site. This observation is the crux of the solution to the specificity problem proposed by Pontiggia et al.; what might distinguish a functional from a cryptic SRY binding site is not the differential affinity of SRY for those sites. Rather, the proposal contends that binding of SRY results in a conformational change in the DNA propitious for the formation of a transcription-related nucleoprotein complex only in the case of a genuine target site interaction. Note that these analyses of the effects of SRY binding to DNA are entirely consistent with SRY being an activator or a repressor; stereospecific modulation of DNA structure might result in transcriptional repression or activation. Indeed, the options of activation or repression by DNA bending may even exist at any single SRY target site, a situation shown to be the case for the bacterial mercury-detoxification genes which may be activated or repressed by binding of the transcription factor MerR to their promoter elements (Ansari et al., 1995). We turn now to what is known about the structure of SRY in other species and review the evidence for a mouse-specific trans-activation domain.

C. SRY: A Transcriptional Activator?

Alignment of SRY sequences indicates that conservation is largely restricted to the HMG box (Tucker and Lundrigan, 1993; Whitfield et al., 1993). The N-terminus shows heterogeneity in length in nonprimate phyla, while the C-terminal, nonbox region is heterogeneous in length even within the primate group. Notably, this region is greatly extended in mouse Sry, consisting of 314 amino-acid residues, compared to only 70 in human SRY (Fig. 2). Determination of the structure of the murine, genital ridge Sry transcript (section I) indicates that this region is present in the sex-determining Sry protein and may have a species-

Andy Greenfield and Peter Koopman

10 HMG box

2

Mouse

Human

58

79

314 amino acids

68

Fig. 2 Comparison of mouse and human SRY protein structure. Neither the amino acid sequence nor the number of amino acids amino- and carboxy-terminal to the HMG box are conserved between these two species. Marsupial SRY has a different structure again (not shown).

specific role to play. Indeed, the discovery of a function for this domain may allow us to choose between two alternative explanations of the high degree of sequence divergence and high frequency of nonsynonymous mutations observed between nonbox SRY open reading frames from different species; either the nonbox regions are functionally unconstrained or species-specific adaptive divergence has occurred. Evidence for a role in transcriptional activation mediated by the long C-terminal region of mouse Sry does exist. For 223 residues this region consists of 20 repeating units of 2- 13 glutamine residues separated by a reiterated histidinerich spacer sequence (Phe-His-Asp-His-His with minor variations) (Gubbay et al., 1992; Tucker and Lundrigan, 1993), and analysis of the nucleotide sequence reveals regions highly repetitive for the trinucleotide CAG, a glutamine codon (Fig. 3). Glutamine-rich regions are characteristic of transcriptional activation domains found in other DNA-binding proteins, such as Spl (Mitchell and Tjian, 1989). Recent experiments, involving assaying activation of a GAL4-responsive reporter gene (Dubin and Ostrer, 1994) have shown that mouse S t y can act as a transcriptional activator in vitro and that this activation function maps to the glutamine/histidine-rich domain. No activation was observed under similar conditions for human SRY. These data raise the possibility that mouse and human SRY function differently in the manner in which they effect transcriptional regulation, supporting the idea of adaptive divergence of the SRY protein. Human SRY has previously been shown to activate transcription of reporter constructs in vitro (Cohen et al., 1994), but evidence for functionally separable DNA-binding and transcriptionalactivation domains exists only for murine Sry. If such a divergence of the mechanism of transcriptional regulation exists between mice and humans, one would predict a potential loss of function for human SRY when active on a murine genomic background. Interestingly, human SRY genomic fragments do not cause sex reversal in XX transgenic mice (Koopman et al., 1991). However, the exact reason for this lack of sex-reversing activity is unknown and may reflect regulatory or structural incompatibilities. Differences in the DNA-binding and -bending capacities of mouse and human SRY protein are well documented (Giese et al.,

11

1. SRY and Mammalian Sex Determination

a

HMG box 2

79

CAG repeats 54

223 (20blocks of 4 -12 aa)

29 aa

Mus musculus 91 (7.5blocks) M. domesticus

Block 3

Poschiavinus 129

b

LL

10 x CAG

12xCAG

FHNHHQQQQQ FYDHHQQQQQQQQQQQQ FHDHHQQKQQ FHDHHQQQQQ F HDHHHHHQEQQ FHNH HQQQQQ FHDHQQQQQQQQQQQ FH D H HQQK QQ FHDHHHHQQQQQ FHDHQQQQQQ FHDHQQQQHQ FHDHPQQKQQ FHDHPQQQQQ FHDHHHQQQQKQQ FH D H H Q Q K Q Q FHDHHQQKQQ FHDHHQQKQQ FHDHHQQQQQ FHDHHQQQQQQQQQQQQQ FHDQQ

Fig. 3 CAG-repeat region in mouse Sty. (a) Over two-thirds of the Mus musculus musculus Sry protein consists of a glutamine-rich region encoded by repeats of the trinucleotide CAG. These glutamine residues are arranged into 20 blocks of 2- 13 glutamine residues separated by a histidinerich spacer sequence (see b). Mus musculus domesricus subspecies lack over half of this repeat region due to the presence of a premature stop codon in the eighth repeat block. The foschiavinus strain shows a variation in the number of CAG repeats found in the third block (Coward et al., 1994), which normally contains 12 CAG repeats. This variation may or may not explain the sex reversal seen when the foschiavinus Sry allele is bred onto a C.57-strain background. (b) Amino acid of the glutaminerich region showing the repeated block stmcture.

1994), a situation which contrasts with the relatively homogeneous group formed by human and primate SRY HMG boxes (Pontiggia et ul., 1995). Despite the above remarks, the possibility still exists that no significant differences exist between mouse and human SRY at the functional level. In vitro analyses of transcriptional activation potential may hold no significance for the function of SRY in vivo. Evidence that the murine Sry C-terminal, glutaminerich domain functions in vivo is not convincing. As we have already discussed in

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Section I, breeding the Y chromosome from certain Mus musculus domesticus strains into the inbred strain C57BL/6J (96) results in hermaphroditic and sexreversed progeny (Eicher and Washburn, 1986; Eicher et al., 1982; Nagamine et ul., 1987a,b). The degree of B6.YDOm sex reversal is dependent on the strain contributing the domesticus Y chromosome, some resulting in no sex reversal at all (Biddle et al., 1988). It has been suggested that observed polymorphisms in the number of CAG repeats in the third block between different domesticus-type Sry alleles might account for the variation in degree of B6.YDOmsex reversal associated with them (Coward et al., 1994) (Fig. 3). This model supports the idea of a function for the glutamine-rich domain, viewing mutations in this region as potentially disruptive to Sry function on a B6 background, possibly disturbing the protein's secondary structure or affecting contact with another protein. However, this argument for functional significance for the CAG-repeat domain is undermined by the observation that a stop codon, truncating the Sry protein in the eighth CAG block, is found in all domesticus subspecies, while a frameshift mutation results in the complete absence of any C-terminal glutamine residues in two distantly related murine species, Mastomys hildebrantii and Hylomyscus alleni (Tucker and Lundrigan, 1993) (Fig. 3). The fact that the glutamine-rich region is completely dispensable for function in S r y proteins from these two Old World mice suggests caution in attributing functional significance to the CAG-repeat block in the domesticus subspecies. A transgenic approach to determine how well-defined Sry variants function on a C57BL/6J background might help to clarify the question of which regions of the S t y gene are responsible for B6.YD0m sex reversal. We will discuss B6.YDomsex reversal as a means of isolating additional sex-determining genes in Section 111.

D. Conclusions

SRY-related sequences are found on the Y chromosome of all mammals tested, including two marsupial species, Sminthopsis macroura and macropus eugenii (Foster et al., 1992). This observation suggests that SRY has been the mammalian Y-linked sex-determining gene for at least 100 million years (Hope et al., 1990). However, conservation outside of the HMG box is almost nonexistent. The evidence that Sry is able to activate transcription via a domain outside the HMG box is difficult to reconcile with the lack of conservation of this domain. Further, differences between the human and mouse proteins may reflect species-specific differences in the biochemistry of SRY function, but there is no direct evidence for a mouse-specific trans-activation domain functioning in vivo and thus no direct evidence for species-specific functions. Concomitant with any selection for divergence in SRY would be divergence in the downstream targets of SRY activity, and it is hard to imagine how significant changes in both effector and target genes could be brought about simultaneously while allowing the crucial process of sex determination to proceed in any individual. This hypothesis would

1 . SRY and Mammalian Sex Determination

13

also predict the existence of a variety of mammalian sex-determining pathways. The alternative model restricts function to the HMG box, explaining the divergence outside this region as a consequence of drift, rather than speciesspecific adaptation. The ability of SRY to bend DNA suggests a mechanism by which the protein might rely on the HMG box alone for its function. This second model is thus simpler and more plausible. A final caveat remains concerning the functional implications of phylogenetic studies in the Rodentia: multiple copies of SRY exist in some Old World rodents (Nagamine, 1994), and care must be taken in excluding the possibility of functional redundancy within a gene family as an explanation of sequence divergence. We now turn our attention to downstream targets of SRY activity with a view to addressing once again the “activator or repressor?” question and we widen the discussion to include other genes thought to play a role in sex determination.

IV. Other Sex Determining Genes What are the genes that regulate the expression of SRY? The precise window of Sry expression exclusively in the developing male gonad suggests tight control by other gene products. Which genes are the targets of transcriptional regulation by SRY? Given that the fate of four separate cell lineages must be determined in the gonad, it is assumed that SRY activity initiates a complex cascade of gene activity resulting in testis formation. Six years after the isolation of SRY the answers to these questions still elude us. However, candidates for these upstream and downstream genes exist, either in the form of isolated sequences or loci which have been mapped but not cloned. One would also predict the existence of genes not involved in sex determination per se, but in the formation of the indifferent gonad, and candidates for these exist too. In this final section we review what is known about the most promising candidates for the above functions.

A. Miillerian Inhibitory Substance (MIS)

Piecing together the cascade of gene activity required for gonadogenesis will no doubt rely on a detailed understanding of the cell biology of gonadogenesis. The credentials of the oldest candidate for direct activation by SRY, MIS, derive from data concerning its own role in testis differentiation. The existence of MIS was predicted by Alfred Jost ( 1 947), based on his observation that synthetic androgen could stimulate the development of male structures in a castrated male fetus, despite Mullerian duct development remaining unaffected. The testis must, he deduced, produce a second hormone (l’hormone inhibitrice) which inhibited the development of female structures derived from the Mullerian ducts, namely the

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uterus, oviducts, and upper vagina. MIS, a member of the transforming growth factor p protein family (Cate et al., 1986), is the first known product of the Sertoli cells. In the mouse, Mis transcripts are first detected in the testis at 12.5 dpc (Munsterberg and Lovell-Badge, 1991). The function, timing, and site of expression of Mis in the male mouse fetus make it an obvious target for activation by Sry. Evidence for a role for SRY in activation of MIS expression is twofold: first, purified SRY protein has been shown to bind an upstream element in the promoter of MIS in vitro (Haqq et al., 1993); second, cotransfection of a gonadal ridge cell line with an SRY expression plasmid and a MIS-promoter reporter construct results in activation of the reporter gene (Haqq et a!. , 1994). However, it has also been shown that while mutation of the MIS promoter abrogates binding of SRY, no affect is observed on reporter gene activation when MIS-promoter constructs containing those same mutations are used in cotransfection experiments (Haqq et al., 1994). Thus, it appears that SRY activates MIS expression via an as yet unidentified intermediary or intermediaries. The question of whether SRY acts directly to activate or repress transcription remains open, since SRY might activate an activator of MIS expression or repress a repressor thereof. Whatever the case may be, it is clear that MIS expression is not dependent on the presence of SRY, since it is also secreted by the granulosa cells of the postnatal ovary. Generation of Mis-deficient mice by gene targeting demonstrates that Mis is not required for testis development (Behringer et al., 1994). M i s - / M i s - homozygous males develop normal testes that produce functional sperm, but are mostly infertile, because the predicted additional development of female reproductive organs interferes with sperm transfer into females. Older male homozygotes also exhibit Leydig cell hyperplasia, indicating that Mis is a negative regulator of Leydig cell proliferation or function. These data are consistent with observations made on transgenic male mice chronically expressing human MIS (Behringer et al., 1990), which show feminization of the external genitalia, impairment of Wolffian duct development, and undescended testes, presumably due to androgen deficiency caused by Leydig cell dysfunction. Transgenic females overexpressing MIS display absence of Mullerian ductderived structures, loss of germ cells, and eventual appearance of seminiferous tubules containing Sertoli cells. This phenotype is similar to the bovine freemartin, a female fetus exposed to a male twin’s blood during development and exhibiting gonadal sex reversal. Despite the association between MIS overexpression and sex reversal, however, it is worth reiterating that MIS is not required for testis differentiation.

B. Steroidogenic Factor 1 (SF-1) and WT-1 Recent data have shown that the orphan nuclear receptor SF- 1, a key regulator of adrenal corticosteroids (Lala et al., 1992), is essential for gonadal development

1. SRY and Mammalian Sex Determination

15

and sexual differentiation. Mice honiozygous for a null allele in the SF-1 gene lack adrenal glands and show complete gonadal agenesis (Luo et a!. , 1994). By 12 dpc both male and female null mice show loss of gonadal tissue through apoptosis, and by 12.5 dpc gonads are absent. Male and female null mice also develop female internal genitalia. What function does SF- 1 have in the developing gonad? Data exist supporting a role for SF-1 in regulating M I S gene expression, consistent with the persistence of female internal genitalia in SF- I-deficient mice. The MIS promoter contains a conserved nuclear receptor half site (AGGTCA) which is critical for expression of the gene in Sertoli cells and which is bound by single Sertoli cell nuclear protein (Shen er a l . , 1994). This nuclear protein is recognized by a specific anti-SF-1 antibody. In addition, SF-I and M I S exhibit concordant spatial and temporal expression in Sertoli cells during embryonic development. The detection of SF-I transcripts as early as 9.5 dpc in the developing gonad and the early arrest of gonadal development in SF-1-deficient mice suggest it must regulate other genes required for gonadogenesis. SF-1 is also known to bind to a sequence common to the promoters of several steroidogenic genes, including P450 aromatase, and may be involved in the regulation of androgen production in the steroidogenic lineage. The sexually dimorphic pattern of SF-1 expression observed in the developing gonad (Shen er al . , 1994) may be evidence for direct activation of SF-1 by SRY, although peak expression of SF-I well after the narrow window of SRY expression argues against this hypothesis. Targeted disruption of the Wilms’ tumor-associated gene WT-I also results in a failure of gonad development (Kreidberg ef a / ., 1993). The initial unaffected stages of genital ridge development, followed by cell death, are a situation reminiscent of SF- 1 deficiency. It seems clear that SF- 1 and WT- 1 play important roles in the development of the gonadal primordia, rather than sex determination per- se, establishing an environment in which SRY can act to commit cells to the male fate. It will be interesting to determine the nature of any functional interactions between WT-1, SF-1, SRY, and their gene products.

c. sox9 Data from knockout mice are clearly proving invaluable in determining the role of penes other than SRY in gonadogenesis One approach to the identification of additional sex determining loci is the use of positional cloning strategies to identify genes whose disruption causes naturally occurring cases of sex reversal. One such example is the gene for autosomal XY sex reversal (SRAI) which resides on human chromosome 17q (Tommerup et al., 1993). Curiously, SRAIdependent sex reversal is associated with campomelic dysplasia (CD), a skeletal malformation syndrome characterized by congenital bowing, angulation of the long bones, and defects of cartilage formation (Houston et al., 1983; Lee et a l . , 1972). Recently, the SRY-related gene SOX9 was shown to map some 80 kb

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Andy Greenfield and Peter Koopman

distal to a chromosome 17 translocation breakpoint found in some CD patients (Foster et al., 1994 Wagner et al., 1994). Nontranslocation patients were found to harbor mutations in SOX9 which would be expected to destroy gene function (Foster et al., 1994; Wagner et al., 1994; Kwok et al., 1995). Expression studies of murine sox9 showed it to be expressed predominantly in mesenchymal condensations throughout the developing embryo before and during cartilage deposition (Wright et al., 1995). These observations suggest that CD may be caused by defective chondrocyte differentiation. But how is this related to sex determi nation? Sox9 transcripts are detected in the developing mouse gonad by RT-PCR (J. Kent, Y. Jeske, and P. Koopman, unpublished data), consistent with a role in gonadogenesis. Three possible explanations exist for the effect of disruption of SOX9 function on testis differentiation. First, SOX9 may act upstream of SRY in the testis-determining pathway, its expression resulting either directly or indirectly in SRY transcription. Second, the converse may be true. In both cases additional, tissue-specific factors must be involved in gene regulation as SOX9 and SRY are not always coexpressed. Third, SOX9 and SRY proteins may interact directly to affect the regulation of one or more genes involved in testis differentiation. Direct activation or protein-protein interaction require the expression of SRY and SOX9 to overlap at some point in the same gonadal cell lineage. The cell specificity of gonadal SOX9 expression is not currently known. Ultimately, a role for Sox9 in murine testis development must be proven by the generation of Sox9-deficient mice by gene targeting. Clearly, however, the sex reversal associated with human SOX9 mutations indicates that this gene can be unequivocally allocated to the mammalian sex determination pathway.

D. Other Loci-DSS

and Tas

XY sex reversal has been associated with duplication of the short arm of the human X chromosome (Ogata et al., 1992; Scherer et al., 1989). This phenotype has been shown to be due to the presence in these XY individuals of two active copies of an Xp locus termed DSS, for dosage sensitive sex reversal (Bardoni et al., 1994). These patients have grossly intact copies of SRY, but exhibit varying degrees of gonadal dysgenesis, ranging from incompletely differentiated testes to streak gonads. However, deletions of the DSS critical region do not disrupt testis differentiation, suggesting that DSS is an ovary-determining gene. It is possible that DSS is a remnant of an ancestral sex determining mechanism which operated by dosage before the evolution of X interaction. Recently, mutations in a novel gene isolated from the DSS critical region have been shown to result in X-linked adrenal hypoplasia congenita and hypogonadotropic hypogonadism (Muscatelli et al., 1994; Zanaria et al., 1994). This gene, termed DAX-I (for DSS-AHC critical region on the X chromosome, gene

1. SRY and Mammalian Sex Determination

17

I ) , is a member of the steroid hormone receptor superfamily. Like SF-1, DAX-I is expressed in both the adrenal glands and the genital ridge and may play a role in gonad development. It will be interesting to determine whether transgenic mice carrying extra copies of the DAX-I gene exhibit gonadal anomalies. Once the DSS gene has been isolated it will also be interesting to establish whether it interacts with SRY, since its ovarian determining function makes it a candidate for repression by the testis-determining gene in males. Autosomal sex reversal loci have been detected in the mouse. The Tus gene, for T-associated sex reversal, is a dominant mutation located on chromosome 17, which results in the development of ovaries or ovotestes in XY C57BL/6J mice (Washburn and Eicher, 1983, 1989; Washburn et al., 1990). In addition, an autosomal or X-linked locus, acting as a modifier of the domesticus-type Spy allele, is responsible for the phenomenon of B6.YDomsex reversal discussed in sections I and 11. However, B6.YDom sex reversal appears to be inherited in a complex fashion, suggesting the involvement of more than one autosomal locus. When mapped, these loci may be isolated and will provide novel entry points into the sex-determining pathway. The products of these genes may act upstream or downstream of Spy, either failing to correctly regulate expression of the domesticus-type Sly allele or failing to respond to its activatingirepressing activity. It is noteworthy that several of the genes discussed in this section encode putative transcription factors: S F - I , WT-I, SOX9, DAX-I, and, of course, SRY. However, a description of important sex-determining genes cannot consist of a list of transcription factors. We know that SRY is likely to masculinize the supporting cell lineage in the initial stages of testis formation, but this masculinizing signal must also be directed to the other gonadal cell lineages, presumably by diffusible factors, cell-surface receptors, and other signal-transducing elements. How might the genes encoding these additional elements be identified? The DNA-binding properties of SRY, as discussed in Section 11, do not appear to allow selection of downstream targets by biochemical means. Novel techniques such as differential display of mRNA (Liang and Pardee, 1992) may prove useful in isolating transcripts which are expressed in a stage- and/or sexspecific fashion in the developing gonad, properties expected of sex-determining genes. In addition, positional cloning of loci implicated in human sex reversal is likely to be a fruitful approach to this problem, as in the case of SOX9.

V. General Conclusions The reader of this review may be struck by its lack of definitive statements about the function of SRY in gonad development. Many of the conclusions drawn here have been cautious and qualified; such is the nature of our understanding of sex determination 6 years after the isolation of SRY. We are unable to offer direct

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Andy Greenfield and Peter Koopman

answers to several important questions: what is the mechanism by which SRY protein regulates the activity of its target gene(s)? Do species-specific functions exist for SRY? Which gene(s) does SRY regulate and how is its own expression controlled? Does it activate or repress the activity of its target gene(s)? But the absence of definitive answers to these questions should not obscure recognition of the progress which has been made. A wealth of data has been amassed on the properties of SRY and its gene product in a range of species, and if this rapid progress continues we can soon look forward to answers to some of the above questions.

Acknowledgments We thank Josephine Bowles, Susan Wheatley, Jill Kent, and Edwina Wright for their helpful comments on this manuscript.

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Hacker, A , , Capel, B., Goodfellow, P., and Lovell-Badge, R. (1995). Expression of Sty, the mouse sex determining gene. Developmenr 121, 1603-1614. Haqq, C. M., King, C.-Y., Donahoe, P. K., and Weiss, M. A. (1993). SRY recognizes conserved DNA sites in sex specific promoters. Proc. Natl. Acad. Sci. USA 90, 1097- 1101. Haqq, C. M., King, C.-Y., Ukiyama, E., Falsafi, S . , Haqq, T. N., Donahoe, P. K., and Weiss, M. A. (1994). Molecular basis of mammalian sex determination: Activation of Miillerian inhibiting substance gene expression by SRY. Science 266, 1494- 1500. Harley, V. R., Jackson, D. I., Hextall, P. J., Hawkins, J. R., Berkovitz, G. D., Sockanathan, S., Lovell-Badge, R., and Goodfellow, P. N. (1992). DNA binding activity of recombinant SRY from normal males and XY females. Science 255, 453-456. Harley, V. R., Lovell-Badge, R., and Goodfellow, P. N. (1994). Definition of a concensus DNA binding site for SRY. Nucleic Acids Res. 22, 1500-1501. Harry, J. L., Koopman, P., Brennan, F. E., Graves, J.A.M., and Renfree, M. B. (1995). Widespread expression of the testis-determining gene SRY in a marsupial. Nature Gene!. 11, 347349. Hope, R., Cooper, S . , and Wainwright, B. (1990). Globin macromolecular sequences in marsupials and momotremes. Aust. J . 2001.37, 289-313. Houston, C. S . , Opitz, J. M., Spranger, J. W., Macpherson, R. I . , Reed, M . H., Gilbert, E. F., Herrmdnn, J., and Schinzel, A. (1983). The campomelic syndrome: Review, report of 17 cases, and follow-up on the currently 17-year-old boy first reported by Maroteaux et al. in 1971. Am. J . Med. Genet 15, 3-28. Jacobs, P. A., and Strong, J. A. (1959). A case human intersexuality having possible XXY sex determining mechanism. Nature (LondonJ 183, 302-303. Jager, R. J., Anvret, M . , Hall, K., and Scherer, G. (1990). A human XY female with a frame shift mutation in the candidate testis-determining gene SRY. Nature (London) 348, 452-454. Jeske, Y. W. A., Bowles, J., Greenfield, A., and Koopman, P. (1995). Expression of a linear Sry transcript in the mouse genital ridge. Nature Genet. 10, 480-482. Jeske, Y. W. A , , Mishina, Y., Cohen, D. R., Behringer, R. R., and Koopman, P. (1996). Analysis of the role of Amh and Fral in the Sry regulatory pathway. Mol. Reprod. Devel., in press. Jost, A. (1947). Recherches sur la differentiation sexuelle de l’embryon de lapin. Archs Anat. Microsc. Morph. Exp. 36, 271-315. Koopman, P., Gubbay, .I.Vivian, , N., Goodfellow, P., and Lovell-Badge, R. (1991). Male development of chromosomally female mice transgenic for Sry. Nature (London) 351, 117-121. Koopman, P., Miinsterberg, A., Capel, B., Vivian, N., and Lovell-Badge, R. (1990). Expression of a candidate sex-determining gene during mouse testis differentiation. Nature (London) 348, 450-452. Kreidberg, J. A., Sariola, H., Loring, J. M., Maeda, M., Pelletier, J., Housman, D., and Jaenisch, R. (1993). WT-1 is required for early kidney development. Cell 74, 679-691. Kwok, C., Weller, P. A., Guioli, S . . Foster, J. W., Mansour, S . , Zuffardi, O., Punnett, H. H., Dominguez-Steglich, M. A., Brook, J. D., Young, I. D., Goodfellow, P. N., and Schafer, A. J. (1995). Mutations in SOX9, the gene responsible for campomelic dysplasia and autosomal sex reversal. Am. J . Hum. Genet. 57, 1028-1036. Lala, D. S . , Rice, D. A . , and Parker, K. L. (1992). Steroidogenic factor 1, a key regulator of steroidogenic enzyme expression, is the mouse homolog of fushi tarazu factor 1. Mol. Endocrinol. 6, 1249-1258. Laudet, V., Stehelin, D., and Clevers, H. (1993). Ancestry and diversity of the HMG box superfamily. Nucleic Acids Res. 21, 2493-2501. Lee, C. H., and Taketo, T. (1994). Normal onset, but prolonged expression, of Sry gene in the B6.YDom sex-reversed mouse gonad. Dev. Biol. 165, 442-452. Lee, F. A , , Issacs, H., and Strauss, J. (1972). The “camptomelic” syndrome. Short life-span

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dwarfism with respiratory distress, hypotonia, peculiar facies, and multiple skeletal and cartilaginous deformities. Am. J . Dis. Child. 124, 485-496. Liang, P., and Pardee, A. B. (1992). Differential display of eukaryotic messenger RNA by means of the polymerase chain reaction. Science 257, 967-971. Luo, X., Ikeda, Y., and Parker, K. L. (1994). A cell-specific nuclear receptor is essential for adrenal and gonadal development and sexual differentiation. Cell 77, 48 1-490. McElreavey, K., Vilain, E., Herskowitz, I . , and Fellous, M. (1993). A regulatory cascade hypothesis for mammalian sex determination: SRY represses a negative regulator of male development. Proc. Natl. Acad. Sci. USA 90, 3368-3372. McLaren, A. (1985). Relation of germ cell sex to gonadal development, In “The Origin and Evolution of Sex” (H. 0. Halvorson and A . Monroy, Eds.), pp. 289-300. Liss, New York. Mitchell, P. J., and Tjian, R. (1989). Transcriptional regulation in mammalian cells by sequencespecific DNA binding proteins. Science 245, 371-378. Munsterberg, A., and Lovell-Badge, R. (1991). Expression of the mouse anti-Mullenan hormone gene suggests a role in both female and male sexual differentiation. Development 113, 613624. Muscatelli, F., Strom, T. M., Walker, A . P., Zanaria, E., Recan, D., Meindi, A , , Bardoni, B., Guioli, S . , Zehetner, G., Rabl, W., Schwarz, H. P., Kaplan, J.-C., Camerino, G., Meitinger, T., and Monaco, A. P. (1994). Mutations in the DAX-I gene give rise to both X-linked adrenal hypoplasia congenita and hypogonadotropic hypogonadism. Nature (London) 372, 672676. Nagamine, C. M. (1994). The testis-determining gene, SRY, exists in multiple copies in Old World rodents. Genet Res. 64, 151-159. Nagamine, C. M., Taketo, T., and Koo, G. C. (1987a). Morphological development of the mouse gonad in rda-l XY sex reversal. Diferentiation 33, 214-222. Nagamine, C. M., Taketo, T., and Koo, G. C. (1987b). Studies on the genetics of rda-1 XY sex reversal in the mouse. Diferenziariun 33, 223-23 I . Nasrin, N., Buggs, C., Fu Kong, X., Carnaza, J., Goebl. M . , and Alexander-Bridges, M. (1991). DNA-binding properties of the product of the testis-determining gene and a related protein. Nature (London) 354, 3 17-320. Ner, S . S . (1992). HMGs everywhere. Curr. Biol. 2, 208-210. Ogata, T., Hawkins, J. R., Taylor, A., Matsuo, N., Hata, J., and Goodfellow, P. N. (1992). Sex reversal in a child with a 46,X,Yp+ karyotype: Support for the existence of a gene(s), located in distal Xp, involved in testis formation. J . Med. Genet. 29, 226-230. Palmer, S . J., and Burgoyne, P. S . (1991a). In situ analysis of fetal, prepuberal and adult XX CJ XY chimaeric mouse testes: Sertoli cells are predominantly, but not exclusively, XY. Development 112, 265-268. Palmer, S . J., and Burgoyne, P. S. (1991b). The Mus musculus domesticus Tdy allele acts later than the Mus musculus musculus Tdy allele: A basis for XY sex reversal in C57BL/6-YPOs mice. Development 113, 709-714. Pontiggia, A., Rimini, R., Harley, V. R., Goodfellow, P. N., Lovell-Badge. R., and Bianchi, M. E. (1994). Sex-reversing mutations affect the architecture of SRY-DNA complexes. EMBO J . 13, 61 15-6124. Pontiggia, A . , Whitfield, S . , Goodfellow, P. N., Lovell-Badge, R., and Bianchi, M. E. (1995). Evolutionary conservation in the DNA-binding and -bending properties of HMG-boxes from the SRY proteins of primates. Gene 154, 277-280. Poulat, F., Girard, F., Chevron, M.-P., GozC, C . , Rebillard, X., Calas, B., Lamb, N., and Berta, P. (1995). Nuclear localization of the testis determining gene product SRY. J . Cell. Biol. 128, 737-748. Rossi, P., Dolci, S . , Albanesi, C . , Grimaldi, P., and Geremia, R. (1993). Direct evidence that

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the mouse sex-determining gene Sry is expressed in the somatic cells of male fetal gonads and in the germ cell line in the adult testis. Mol. Reprod. Devel. 34, 369-73. Scherer, G.,Schempp, W., Baccichetti, C . , Lenzini, E., Bricarelli, F. D., Carbone, L.D.L., and Wolf, U. (1989). Duplication of an Xp segment which includes ZFX locus causes sex inversion in man. Hum. Genet. 81, 291-294. Shen, W.-H., Moore, C.C.D., Ikeda, Y., Parker, K. L., and Ingraham, H. (1994). Nuclear receptor steriodogenic factor 1 regulates the Miillerian inhibiting substance gene: A link to the sex determination cascade. Cell 77, 651-661. Sinclair, A. H., Berta, P., Palmer, M. S . , Hawkins, J. R., Griffiths, B. L., Smith, M. J., Foster, J. W., Frischauf, A.-M., Lovell-Badge, R., and Goodfellow, P. N. (1990). A gene from the human sex-determining region encodes a protein with homology to a conserved DNA-binding motif. Nature (London) 346,240-244. Su, H . , and Lau, Y.-F. (1993). Identification of the transcriptional unit, structural organization, and promoter sequence of the human sex-determining region Y (SRY)gene, using a reverse genetic approach. Am. J . Hum. Genet. 52, 24-38. Tommerup, N . , Schernpp, W., Mienecke, P., Pedersen, S . , Bolund, L., Brandt, C., Goodpasture, C., Guldberg, P., Held, K. R., Reinwein, H., Saaugstad, 0. D., Scherer, G.,Skjeldal, O., Toder, R., Westvik, J., van der Hagen, C. B., and Wolf, U. (1993). Assignment of an autosoma1 sex reversal locus (SRAI) and campomelic dysplasia (CMPDI) to 17q24.3-q25.1. Nature Genet. 4, 170-174. Travis, A , , Amsterdam, A., Belanger, C., and Grosschedl, R . (1991). LEF-I, a gene encoding a lymphoid-specific with protein, an HMG domain, regulated T-cell receptor 01 enhancer function. Genes Dev. 5 , 880-894. Tucker, P. K., and Lundrigan, B. L. (1993). Rapid evolution of the sex determining locus in Old World mice and rats. Nature (London) 364, 715-717. Upadhyay, S . , Luciani, J.-M., and Zamboni, L. (1981). The role of the mesonephros in the development of the mouse testis and its excurrent pathways. In “Development and Function of Reproductive Organs” (A. G.Byskov and H. Peters, Eds.), pp. 18-27. Excerpta Medica, Amsterdam. van de Wetering, M., and Clevers, H. (1992). Sequence-specific interaction of the HMG box proteins TCF-1 and SRY occurs within the minor groove of a Watson-Crick double helix. EMBO J . 11, 3039-3044. van de Wetering, M., Oosterwegel, M., Dooijes, D., and Clevers, H. (1991). Identification and cloning of TCF- 1, a T lymphocyte-specific transcription factor containing a sequence-specific HMG box. EMBO J . 10, 123-132. Vilain, E., Fellous, M. J., and McElreavey, K. (1992). Characterisation and sequence of the 5’ flanking region of the human testis-determining factor SRY. Methods Molec. CeII. Biol. 3, 128- 134. Wagner, T., Wirth, J., Meyer, J., Zabel, B., Held, M., Zimmer, J., Pasantes, J., Bricarelli, F. D., Keutel, J., Hustert, E., Wolf, U., Tommerup, N., Schempp, W., and Scherer, G. (1994). Autosomal sex reversal and campomelic dysplasia are caused by mutations in and around the SRY-related gene SOX9. Cell 79, 11 I I- 1120. Washburn, L. L., and Eicher, E. M. (1983). Sex reversal in XY mice caused by dominant mutation on chromosome 17. Nature (London) 303, 338-340. Washbum, L. L., and Eicher, E. M. (1989). Normal testis determination in the mouse depends on genetic interaction of locus on chromosome 17 and the Y chromosome. Genetics 123, 173-179. Washbum, L. L., Lee, B. K., and Eicher, E. M. (1990). Inheritance of T-associated sex reversal in mice. Genet. Res. 56. 185-191.

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Welshons, W. J., and Russell, L. B. (1959). The Y chromosome as the bearer of male determining factors in the mouse. Proc. Natl. Acad. Sci. USA 45, 560-566. Whitfield, L. S . , Lovell-Badge, R., and Goodfellow, P. N. (1993). Rapid sequence evolution of the mammalian sex determining gene SRY. Nature (London) 364, 713-715. Wright, E., Hargrave, M. R., Christiansen, J . , Cooper, L., Kun, J . , Evans, T., Gangadharan, U., Greenfield, A , , and Koopman, P. (1995). The Sry-related gene Sox-9 is expressed during chondrogenesis in mouse embryos. Nature Genet. 9, 15-20. Wright, E. M., Snopek, B., and Koopman, P. (1993). Seven new members of the Sox gene family expressed during mouse development. Nuckic Acids Res. 21, 744. Zanaria, E., Muscatelli, F., Bardoni, B., Strom, T. M., Guioli, S., Guo, W., Lalli, E., Moser, C., Walker, A. P., McCabe, E.R.B., Meltinger, T., Monaco, A. P., Sassone-Corsi, P., and Camerino, G. (1994). An unusual member of the nuclear hormone receptor superfamily responsible for X-linked adrenal hypoplasia congenita. Narure (London) 372, 635-641. Zwingman, T., Erickson, R. P., Boyer, T., and Ao, A. (1993). Transcription of the sex-determining region genes Sry and Zfy in the mouse preimplantation embryo. Proc. Narl. Acad. Sci. USA 90, 814-817.

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2 Transforming Sperm Nuclei into Male Pronuclei in Vivo and in Vitro D. Poccia Department of Biology Amherst College Amherst, Massachusetts 01002

P. Collas Department of Food Science Agricultural University of Norway As, Norway

I. Introduction A. Comparative Overview of Male Pronuclear Development in Vivo B . Comparisons of Cell-Free Preparations 11. Changes in Nuclear Proteins A. Modifications of Sperm Proteins and Exchange for Maternal Histones B. Summary and Speculations 111. Chromatin Decondensation

A. Conditions Promoting Decondensation in Vivo B. Conditions Promoting Decondensation in V i m C. Summary and Speculations IV. Formation or Adjustment of Nucleosomes A . Amphibians B. Fruit Flies C. Sea Urchins V. Nuclear Envelope Disassembly and Assembly A . Removal of the Sperm Nuclear Envelope and Initiation of Nuclear Envelope Formation B . Nuclear Envelope Formation C. Role of Lamins D. Nuclear Pores E. Summary and Speculations VI. Male Pronuclear Activities A . Replication B. Reinitiation of Transcription C . Summary and Speculations VII. Conclusions References

Currenr Topics in Developmenral5io/ugx Vol 34

Copyright 0 1996 by Academic Press, Inc. All rights of reproduction in any l o r n reserved

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1. Introduction The sperm nucleus shares characteristics with nuclei of some other terminally differentiated cell types such as red blood cells or lens cells. It is genetically inactive, with highly condensed chromatin. Unlike the nuclei of terminally differentiated cells, nuclei of sperm successful in fertilization are reactivated. These sperm nuclei are transformed by egg cytoplasm following fertilization into functional nuclei resembling those of active somatic cells. The formation of the male pronucleus is critical in permitting the male genome to contribute its half of the zygotic gene complement to the embryo and adult to follow. If the process is deficient, accurate transmission of the male genome may be compromised, with severe consequences for the embryo. In order to form a male pronucleus, the egg cytoplasm has to undo much of what was done during spermatogenesis in forming the sperm nucleus. Its most important activities are the replacement or modification of special DNA-associated proteins used to package the sperm genome, decondensation and adjustment of the sperm chromatin to a configuration consistent with the reinitiation of replication, transcription and mitosis, and the replacement of the sperm nuclear envelope lacking pores with a new nuclear envelope containing pores. These three processes transform the dormant sperm nucleus into a functional male pronucleus. This transformation has been studied at several levels. The morphological outlines of male pronuclear development have been known from light microscopy analyses for over a century (Wilson, 1898). Much further work and the addition of ultrastructure analysis has enriched our views of the process. Experimental approaches in which pronuclear development is perturbed by chemical and other inhibitors, the extension of techniques for microinjection of sperm nuclei and their subsequent analysis, the use of polyspermic eggs for isolation of male pronuclei or augmentation of transcription signals, sensitive reagents and techniques particularly fluorescent antibodies and autoradiography, and most recently the development of cell-free systems have added more molecular information and insight into the regulation of pronuclear transitions. Relatively little attention has yet been paid to the genetic analysis of pronuclear development. Preferred model systems should combine in vivo and in vitro analyses: in vivo because such transformations are benchmarks against which other observations must be compared and in vitro because precise manipulations of conditions are required to investigate the details of molecular mechanisms. We consider in this review the four organisms in which cell-free systems prepared from egg lysates have been devised to analyze male pronuclear formation: amphibians (particularly Xenopus laevis and Bufo japonica), fruit flies (Drosophila melanogaster), surf clams (Spisula solidissirnu), and sea urchins (mostly Lytechinus pictus). This group represents four different fertilization strategies and encompasses vertebrates and invertebrates. We will examine where data from these four systems are

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in accord and where they are not. We will emphasize recent in vitro results, compare them where possible with the corresponding observations in vivo, and speculate on some unresolved issues. Each organism has unique advantages, and a combination of approaches using these organisms, and hopefully a few others, will extend our ideas about how male pronuclei form and deepen our molecular understanding of how this is achieved. This comparative approach, while currently limited in certain types of data for each of the organisms, will we believe ultimately allow for some general principles to emerge. Our intention is a comparative rather than an exhaustive treatment. Many excellent reviews are available which deal in depth with various topics discussed here. The reader is referred in particular to the following: general comparative reviews of the ultrastructure and regulation of male pronuclear development (Gurdon and Woodland, 1968; Longo, 1973, 1981, 1985, 1991; Longo and Kunkle, 1978), in vivo studies on the biochemistry of nuclear proteins (Bloch, 1969; Poccia, 1986, 1987, 1989, 1991, 1995; Kasinsky, 1989; Poccia and Green, 1992; Green et al., 1995), the use of in vitro systems for chromatin remodeling (Shimamura et al., 1989; Laskey et al., 1993; Almouzni and Wolffe, 1993; Katagiri and Ohsumi, 1994), transcription and replication studied with in vitro systems (Becker et al., 1994; Dimitrov and Wolffe, 1995), nuclear architecture including lamins and pores (Nigg, 1992; Rout and Wente, 1994; Gerace and Foisner, 1994; Hernandez-Verdun and Gautier, 1994), and nuclear envelope assembly and disassembly in vitro (Cox and Hutchinson, 1994; Hutchison et al., 1994).

A. Comparative Overview of Male Pronuclear Development in Vivo

Eggs are normally fertilized at one of four maturation stages depending on species (Table I). These are meiotic prophase I (germinal vesicle stage), meiotic metaphase I, meiotic metaphase 11, or after completion of meiosis (pronuclear stage). Oogenesis arrests at these stages, and completion of meiosis or resumption of the cell cycle is normally triggered by the fertilizing sperm. Although the eggs of a given species are typically fertilized at only one stage, some eggs can be fertilized experimentally at stages other than their normal one, this being prevented in nature by avoiding proximity of the gametes (for example, by sequestration in the female) or incapacity of the gamete membranes to fuse. After gamete fusion that defines the onset of fertilization, the sperm nucleus gains access to egg cytoplasm and pronuclear formation ensues. Male pronuclei then follow one of two paths. In some organisms, the male and female pronuclei fuse together to form a zygote nucleus prior to mitosis. In others, mixture of the parental chromosomes is delayed until the first mitosis at which time each pronucleus loses its nuclear envelope and the condensed parental chromosomes

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Table I Comparison of Model Organisms for Studying Pronuclear Formationa

Organism Amphibian Fruit fly Surf clam Sea urchin

Fertilization type

Stage at fertilization

External Internal External External

MI1 MI GV PN

Length of embryo cycles (min)

Duration of DNA synthesis (min)

40 10

10

25-30 60-90

4 -

10-13

Syngamy type “Ascaris” “Ascaris” “Ascaris” “Sea urchin”

a Abbreviations: GV, germinal vesicle stage; MI, first meiotic metaphase; MII, second meiotic metaphase; PN, pronuclear stage. See text for references.

intermix and become surrounded by a common nuclear envelope. These paths represent respectively the so-called “sea urchin type” and “Ascaris type” distinguished by Wilson (1925). The four model organisms represent the four stages of fertilization and both paths to the zygote nucleus. Each of the model organisms has advantages and disadvantages for studying pronuclear transformations in vivo. Amphibian eggs, like those of most vertebrates, are fertilized at meiotic metaphase 11. Fertilization is external and easily induced. Exceptionally large quantities of eggs are available. The cell cycle in early X . laevis embryos takes about 40 rnin with about 10 min of this occupied by DNA synthesis (Myake-Lye et al., 1983). The large eggs lend themselves to microinjection experiments. Electron microscopy studies are few and eggs are highly pigmented and large so detailed light microscopy of pronuclear morphology has been limited, especially in living cells. Fruit fly eggs are fertilized at meiotic metaphase I. Fertilization occurs internally. The exceptionally long sperm enters a micropyle at the anterior end of the egg (Karr, 1991). Meiosis is complete by 10 rnin and approach of the pronuclei occurs 16-17 min after egg laying (Schneider-Minder, 1966). The first nine cycles of nuclear reproduction last 10 rnin with a replication period of 4 rnin (Kriegstein and Hogness, 1974). Cycles 10-13 last about 15 min, so that it takes just over 2 hr for the first 13 divisions producing -6000 nuclei. The syncytium then undergoes cleavage divisions to produce a cellular blastoderm. The syncytial stages are unique to Drosophila among the four model organisms. Drosophila has a great potential advantage over the other organisms in its wellunderstood and easily manipulated genetics. In vitro fertilization is problematic, but microinjection experiments are feasible. Surf clam fertilization is external in sea water and easily induced. Because fertilization is at the germinal vesicle stage, subsequent steps of the meiotic cycle and female chromosome behavior can be followed in parallel with male pro-

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nuclear development. Arrested eggs are probably released from the G21M border at fertilization by a mechanism involving MAP kinase (Shibuya et al., 1992). Early cell cycles take about 15-30 min (Allen, 1953). Large numbers of eggs can be obtained and biochemical studies have been performed on cell cycle progression, inhibitor studies of pronuclear development in vivo have been reported, and excellent electron microscopy descriptions are available. Surf clams can be made polyspermic, but this procedure has not been exploited for nuclear protein analysis as it has for sea urchins. Sea urchin eggs are normally fertilized at the pronuclear stage, having completed meiosis. Gametes are abundant. Fertilization is external in sea water and easily induced. Early cell cycles are 60-90 min with 10- to 13-min periods of DNA synthesis (Hinegardner et al., 1964). Unfertilized eggs can be partially released from Go arrest without fertilization by raising intracellular pH with ammonia or they can be fully activated with Ca*+ ionophores. Excellent electron microscopy descriptions and inhibitor studies have been reported as have microinjection studies. The sea urchin is the only organism for which extensive studies of in vivo transitions in pronuclear histones and chromatin physical structure have been made (using male pronuclei isolated from polyspermically fertilized eggs). Genetic analysis is limited.

B. Comparisons of Cell-Free Preparations

Cell-free systems supporting pronuclear formation have been reported for amphibians, fruit flies, surf clams, and sea urchins. Each of the organisms produces large quantities of eggs which are easily procured and all but Drosophila produce large quantities of easily obtainable sperm as well. In the original procedures with amphibians, sperm nuclei were permeabilized with 0.05% lysolecithin, which removes most of the nuclear envelope (Lohka and Masui, 1983a). This procedure has become standard for preparation of sperm nuclei in all four systems, although sometimes extraction with the nonionic detergent Triton X- 100 is used which may result in more thorough removal of the envelope. The eggs of all four organisms have large stockpiles of nuclear constituents stored for the rapid cleavage divisions that follow during embryogenesis. These constituents include histones, nuclear envelope precursors such as membranes and lamins, and various chromatin and membrane assembly factors. Since frogs, clams, and urchins are externally fertilized or activated, populations of eggs can be obtained synchronized for cell cycle stages. For Drosophila this is impractical and lysates are prepared from unsynchronized batches of embryos pooled from several individuals, usually from 0-2 or 0-6 hr. Amphibian extracts so far are the only ones reported capable of in vitro cycling. The first and by far best characterized cell-free system for analysis of pronuclear development is from the frogs X . laevis and Rana catebesiana and from

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the toad B . japonicus. Amphibian egg lysates support replication, transcription, nuclear envelope assembly, chromatin decondensation, cell cycle progression, and chromatin assembly (Almouzni and Wolffe, 1993). Cell-free systems from the fruit fly Drosophila support chromatin decondensation of amphibian and avian sperm and limited replication. Nuclei form nuclear envelopes and incorporate lamins. Homologous sperm is not used, and extracts are taken from unsynchronized early embryos. Since the embryos have undergone several nuclear cycles they may contain somewhat depleted pools of nuclear precursors. Egg lysates of the surf clam Spisula support surf clam sperm chromatin decondensation and nuclear envelope formation. Activated oocyte cytoplasm is more effective than unactivated cytoplasm. Envelope assembly is accompanied by lamin and nuclear pore assembly. Sea urchin egg lysates of L . pictus support sea urchin sperm chromatin decondensation and pronuclear envelope formation as well as limited replication of frog nuclei. Fertilized egg cytoplasm is more effective than unfertilized and nuclear envelope formation is accompanied by lamin assembly. Since preparation of extracts may affect the extent of pronuclear development, procedures and conditions for the four model organisms are given in detail below. Buffers used to date differ in a number of constituents, and not all variations of such important factors as protease inhibitors, protein synthesis inhibitors, phosphatase inhibitors, pH, or Ca2+ concentrations have been explored. Inclusion of ATP and an ATP-generating system is common but not universal.

1. Amphibians The procedures for preparing extracts have been extensively studied and vary depending on the purposes of the experiments (Masui et al., 1984; Lohka and Masui, 1984; Almouzni and Wolffe, 1993). Frog eggs are arrested in meiotic metaphase I1 with high levels of maturation promoting factor MPF. When the large eggs are broken open by centrifugation, Ca2+ is released from internal pools triggering maturation. If lysed in the presence of EGTA to prevent Ca2+dependent cyclin degradation and phosphatase inhibitors to prevent loss of phosphates from MPF, stabilized M-phase extracts can be made. S-phase extracts are made by omitting the EGTA and including cycloheximide to inhibit resynthesis of cyclin. Extracts which cycle in vitro are made in the same way but omitting the cycloheximide. Cells are broken by centrifugation and cytoplasm clarified at 10,OOOg (S 10). Further fractionation can be performed by preparing a high-speed supernatant at 150,OOOg (S150) virtually devoid of membranes from the S10. S 150 is capable of supporting complementary strand DNA synthesis, chromatin assembly, and transcription. In S 10, double-stranded replication and nuclear

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2. Transforming Sperm Nuclei into Male Pronuclei

assembly occur. In the original formulation, Rana pipiens cytoplasm was made by activating eggs by electric shock, washing them in 250 mM sucrose-200 mM KC1-1.5 mM MgC1,-2 mM P-mercaptoethanol-10 mM Tris, pH 7.5, crushing by centrifugation at 15,OOOg for 15 min, and subsequently removing yolk and pigment layers (Lohka and Masui, 1983a).

2. Fruit Flies Because fruit fly eggs are fertilized internally, deposited fertilized eggs (early embryos) are collected over a period of time, pooled, and homogenized. Berrios and Avilion (1990) used 0- to 5-hr embryos which were homogenized using a French press into a buffer of 10% ethylene glycol-250 mh4 sucrose-I00 mM NaC1-2.5 mM MgC1,-1 mM EDTA-2 mM DTT-10 mM Hepes, pH 7.5, containing a cocktail of protease inhibitors. After centrifugation at 12,OOOg for 10 min, the middle clear zone was used. ATP and an ATP-generating system were added for nuclear assembly. Crevel and Cotterill (199 1 ) used essentially the same protocols. Ulitzur and Gruenbaum (1989) homogenized 1- to 6-hr embryos into 250 mM sucrose-2.5 mM MgCl,-50 mM KCl-100 mg/ml cycloheximide5 pg/ml cytochalasin B- 1 mM DTT without protease inhibitors. Homogenates were centrifuged for 5 min at 14,OOOg. Ulitzur et al. (1992) modified these protocols by addition of a cocktail of protease inhibitors. Kawasaki et al. (1994) prepared S25 extracts from 0- to 2-hr embryos, broken with a Dounce homogenizer into 50 mM NaCl-5 mM MgCl,-lmM DTT-50 mM Hepes, pH 7.5, supplemented with a protease inhibitor cocktail.

3. Surf Clams Longo et al. (1994) washed unactivated or artificially activated surf clam eggs in 1 M glycerol buffered with sodium phosphate, pH 8, and lysed them into 0.1 M NaCI-5 mM MgC1,-20 mM Pipes, pH 7.2. Lysis was accomplished by vigorous pipetting, and the extract centrifuged at 10,OOOg for 15 min in the absence of ATP and an ATP-generating system. SlOOs were also prepared. No protease inhibitors were included. A cell-free extract for disassembly of the clam oocyte nuclear envelope was devised by Dessev et al. (1991). Eggs were lysed into 0.1 M KC1-5 mM MgC1,-10 mM EGTA-25 mM P-glycerophosphate-2 mM DTT-20 mM Pipes, pH 7.2, and the homogenate was centrifuged at 20,000g for 10 min. 4. Sea Urchins

-

In the system of Cameron and Poccia (1994) fertilized eggs 15 min into the cell cycle were lysed with a 22-gauge hypodermic needle. The lysis buffer was 150

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D. Poccia and P. Collas

mM NaCl-5 mM MgC1,-25 mM EGTA-110 mM glycine-250 mM glycerol1 mM DTT-1 mM PMSF-10 mM Hepes, pH 8, supplemented with ATP and an ATP-generating system. The final pH of the homogenate was -7. The homogenate was centrifuged at 10,OOOg for 10 min and the supernatant taken. In the system of Zhang and Ruderman (1993), the lysis buffer was 40 mM NaC12.5 mM MgC1,-300 mM glycine- 100 mM potassium gluconate-2% glycerol50 mM Hepes, pH 7.4, including a protease cocktail. Homogenates were centnfuged at 10,OOOg for 15 min, supplemented with ATP and an ATP-generating system and recentrifuged.

II. Changes in Nuclear Proteins Although eukaryotic chromatin typically consists of DNA packaged by wellconserved histones into a regular repeating nucleosomal structure, sperm chromatin DNA is associated with an almost bewildering array of special proteins depending on species (Bloch, 1969; Poccia, 1986; Kasinsky, 1989). Some of these are protamines, small arginine-rich proteins which are incapable of forming nucleosomes. Others are protamine-like. Others are genuine but male germ line specific histones that differ from somatic histones in significant ways. Since amino acid sequences of the nucleosomal core histones (H2A, H2B, H3, and H4) and the central region of the linker histone H1 are rather highly conserved in somatic cells, remodeling of sperm nuclear proteins to the somatic state must occur sometime in the early embryo. In effect, egg cytoplasm must replace the “foreign” sperm proteins with somatic histones. This remodeling takes place rapidly after fertilization in all cases known and may involve modification and/or removal of the sperm proteins and acquisition of stored maternal histones. There is a premium placed on rapid replacement of protamines or protaminelike sperm proteins with histones prior to the first mitosis. Even male pronucleicontaining sperm-specific histones are likely to replace these relatively soon after fertilization. The replacement histones come from a store of preformed maternal histones in all or most eggs. These may be complexed with chaperone proteins such as nucleoplasmin or Nl/N2 as in amphibians (Kleinschmidt et al., 1985). For almost all other organisms, the storage form and location of these nonchromosomal maternal histones is not known, although at least some must be cytoplasmic (Salik et al., 1981). Isolation of male pronuclei from fertilized eggs is formidably difficult because of the high cytoplasm/nucleus ratio. Details of nuclear protein remodeling are only available for the amphibians and sea urchins. Most data for amphibians come from in vitro systems and for the urchins from study of transitions occurring in polyspermically fertilized eggs.

2. Transforming Sperm Nuclei into Male Pronuclei

33

A. Modifications of Sperm Proteins and Exchange for Maternal Histones

1. Amphibians Diverse types of sperm chromatin proteins are found among the amphibians. Bufo sperm nuclei contain no histones, but instead two fast moving protaminetype molecules, P1 and P2 (Takamune et al., 1991). These differ in one of 39 amino acids. Both have arginine clusters and no cysteine, thus resembling fish rather than mammalian protamines. Bufo P2 mRNA is restricted to the testis, appearing first in spermatids (Mita et al., 1991). Xenopus sperm nuclei have six sperm specific proteins (SP1-6) in addition to the core histones but lack H1. They have large amounts of somatic H3 and H4 but little H2A or H2B. SP3-6 are 33-41% arginine and contain little lysine. SP2 is similar to H3 and H4 histones in its lysine/arginine ratio. SP4 (78 amino acids) and SP5 (74 amino acids) are intermediate in composition between histones and protamines. SP3 and SP6 are probably encoded by the same gene. S P l , SP2, and SP4 may also be encoded by a single gene (Ariyoshi et al., 1994). SP proteins appear in the last steps of nuclear condensation during spermatogenesis (Yokota et al., 1991). Xenopus SP4 RNA is restricted to the testis and first seen in primary spermatocytes (Mita et al., 1991; Hiyoshi et al., 1991). R a m sperm DNA is packaged with somatic histones and a sperm-specific H 1-like histone variant (Kasinsky et al., 1985). Sperm chromatin of all three species is nonnucleosomal (Katagiri and Ohsumi, 1994). Transitions in sperm nuclear proteins of several amphibians have been studied. Removal of protamines is rapid. In vivo, protamines are removed from Bufo sperm nuclei within 5 min of fertilization as judged by immunofluorescence (Ohsumi and Katagiri, 1991a). The timing or details of protein post-translational modifications in vivo are not known. For example, it is not known if SP proteins are phosphorylated in vivo before removal but phosphorylation in vitro facilitates their removal (Katagiri and Ohsumi, 1994). In vitro, Bufo sperm chromatin loses its protamines within 1 min in egg extracts (Ohsumi and Katagiri, 1991a). In Xenopus, proteins y and z (two of the SP proteins) are also removed rapidly in an S 150 extract (Dimitrov et al., 1994). The protamine removing activity has been isolated. Oocyte cytosol (but not postneurula or adult cytosol) contains an activity which removes protamines in vitro (Ohsumi and Katagiri, 1991). It purifies as a protein running at 36 kDa on SDS gels which induces decondensation and has been identified as nucleoplasmin, a protein previously characterized as an assembly factor for histones. The protein was independently identified from Xenopus by Philpott et al. (1991). Nucleoplasmin structure and function have been recently reviewed (Laskey et al., 1993). Nucleoplasmin is an acidic heat stable protein which binds to histones in vitro, shields their positive charge, and facilitates ordered nucleosome assembly on DNA. In vivo, nucleoplasmin is bound to histones H2A/H2B, whereas

34

D. Poccia and P. Collas

H3/H4 are complexed with other proteins (Kleinschmidt et al., 1985). Nucleoplasmin binds 12,000 diploid histone equivalents in the maternal storage pool. Nucleoplasmin may function in storage of these large maternal histone pools or may chaperone histones whose synthesis is not S-phase coupled. It apparently has no role in somatic cells. Nucleoplasmin potentially links several early transitions in male pronuclei. Philpott and Len0 (1992) showed that purified nucleoplasmin will remove SP proteins X and Y and assemble H2A and H2B on decondensing sperm chromatin. X and Y are bound to nucleoplasmin in a complex immunoprecipitated with anti-nucleoplasmin antibodies. Thus they hypothesize that nucleoplasmin facilitates an exchange of SP proteins for histones. This exchange correlates with the formation of a nucleosomal ladder in the protamine-depleted male pronuclear chromatin which as sperm chromatin was nonnucleosomal and deficient in H2A and H2B. If heated Rana extracts are used with sperm nuclei, sperm lose their protamines under the influence of the heat-stable nucleoplasmin, but no histones assemble (Ohsumi and Katagiri, 1991a). Under these conditions, the chromatin decondenses extensively but is very fragile. Since nucleoplasrnin is sufficient for frog sperm chromatin decondensation in vitro (see below), the removal of protamines, acquisition of histones, formation of nucleosomes, and decondensation of chromatin are tightly linked in amphibians. Removal of sperm proteins is accompanied by exchange for maternal histones. Approximately 20,000 diploid equivalents of histones are found in amphibian eggs (Adamson and Woodland, 1977; Woodland, 1980). They are stored in the germinal vesicle or cytoplasm and suffice for all of early development (Woodland, 1982). These large pools allow for the assembly of up to 80,000 pg of exogenous DNA per egg in vitro (Laskey et al., 1977). Details on the utilization of these stores in male pronuclei are mostly derived from in vitro studies. In Xenopus or Bufo, replacement core histones in vitro are of the somatic type, except for H2A.X, a large H2A variant (Ohsumi and Katagiri, 1991a). Large amounts of typical somatic H1 are stored in the egg but this protein is not incorporated into the chromatin. Instead, histone H1 . X appears in the pronuclei. H1.X and H2A.X are larger than the corresponding somatic forms. They may be counterparts of the sea urchin histone variants CS H1 and CS H2A involved in male pronuclear chromatin remodeling (see below). H1 .X and H2A.X persist in embryos up to the blastula stage and are replaced at the gastrula stage by their somatic counterparts. Xenopus patterns are similar to Bufo, but Xenopus nuclei acquire four variants of H1 .X (Ohsumi and Katagiri, 1991a). Substitution patterns in embryos are otherwise similar up to late blastula. Typical somatic histone composition is not achieved until gastrulation (Dimitrov et al., 1995). H1.X is likely to be the protein B4, although this has not yet been verified (Hock et al., 1993). B4 mRNA in Xenopus is expressed during oogenesis and embryogenesis through preneurula stages. The 29-kDa protein is 29% homolo-

2. Transforming Sperm Nuclei into Male Pronuclei

35

gous with somatic H1 but lacks the normally well-conserved central globular domain characteristic of most HI molecules (Smith et al., 1988). Much of the core histone replacement involves post-translationally modified proteins (Dimitrov et al., 1994). Some of these are newly synthesized. In Xenopus, during the prereplicative period of rapid chromatin decondensation in vitro, deacetylated H2B, H2A, and H2A.X assemble (Fig. 1). H2A.X is stored in a complex with nucleoplasmin. The H2A and H4 from the sperm are phosphorylated in egg extracts and H2A.X and H2A from the storage pool are also phosphorylated. H2B is not. Sperm H2A is phosphorylated by a kinase in the sperm itself. Histone phosphorylation might involve release from storage but is not essential for assembly or nucleosome spacing. After replication, all core histones are incorporated. Incorporation of newly synthesized H3 and H4 requires replication. During replication, H3 from the pool and diacetylated H4 accumulate.

A RBC

Sperm Nuclei

plus Egg Extract

RBC

Sperm Nuclei

Fig. 1 In v i m remodeling of Xenopus sperm nuclear proteins. Two-dimensional gel electrophoresis of proteins of Xenopus luevis sperm before (A) and after (B) remodeling in Xenopus egg high-speed extract. RBC, Xenopus red blood cell histone control. 0 and 1, deacetylated and monoacetylated forms of H4. X , y, and z, sperm-specific proteins. B 4 is H1-type molecule. t, HMG2; s, H2A.X. Within 10 min, y and z are lost, deacetylated H2A.X and HMG2 appear. Taken from Dimitrov er ul. (1994) with permission.

36

D. Poccia and P. Collas

Linker histone analysis is complicated by the absence of a typical H 1. Prior to replication, B4 and the high-mobility group protein HMG2 are incorporated into chromatin (Dimitrov et a/., 1994). Postreplicatively phosphorylated HMG2 and B4 are taken up. The role of B4 in nuclear assembly was assessed by immunodepletion in Xenopus (Dasso et al., 1994). Chromatin decondensation appears normal and nuclei form with an envelope capable of transport and replication, a lamina, and prereplication centers. No differences are detected from control nuclei. Thus B4 appears not to be essential for chromatin compaction or to provide a scaffold for looped structures involved in replication. It was suggested that HMG2 might take over some of these roles. The role of H1 in mitotic chromosome condensation was examined by similar immunodepletion experiments (Ohsumi et al., 1993). H1 .X was depleted from 90-min extracts going into the first mitosis. Chromosomes condensed normally and had typical 200-bp repeat lengths. They differed only in their fragility to pipetting. The experiments suggest that H1 is not required for mitotic condensation.

2. Fruit Flies Most of the literature on Drosophila sperm and pronuclear histones is cytochemical. Mature Drosophila sperm contain arginine-rich histones, not protamines (Das et al., 1964). A candidate for a gene encoding a sperm-specific histone (or transition protein) has been isolated (Russell and Kaiser, 1993). It is an autosoma1 gene expressed in the male germ line with 45% similarity to the red blood cell H5 histone class and has a cysteine-rich region probably related to mammalian protamine sequences. Cytochemically, early embryo nuclei and pronuclei have atypical histones but typical somatic staining patterns appear just before blastulation (Das et al., 1964). An HMG-I-type protein, HMG-D, is found during the first six cycles of Drosophila embryo chromatin which lacks immunochemically detectable H 1. Although not yet demonstrated in male pronuclei, HMG-D has been suggested to keep the chromatin in a less condensed state prior to transcriptional activation, perhaps facilitating rapid cycling (Ner and Travers, 1994). Somatic histones are modified in the early cell cycles. Drosophila nuclear Hl from 0- to 2-hr embryos is not highly phosphorylated, although mitosis takes up a large fraction of the early cycles (Giancotti et al., 1984). Perhaps this reflects the storage form in the maternal pool. H4 and H3 are highly acetylated and probably derive primarily from a cytoplasmic pool for early assembly reactions. H2A is modified, but not H2B. Drosophila cleavage stage nuclei contain a large H2A variant (H2AP) which is selectively phosphorylated (Holmgren et al., 1985). In vitro, Drosophila lysates remove Xenopus sperm-specific proteins X and Y rapidly from the chromatin (Kawasaki et al., 1994).

2 . Transforming Sperm Nuclei into Male Pronuclei

37

3. Surf Clams The sperm of the surf clam contains a high molecular weight nuclear protein with properties intermediate between protamines and histones (Ausio and Subirana, 1982). Its chromatin is nonnucleosomal. In vivo transitions in male pronuclear DNA binding proteins or modifications of sperm proteins in surf clam have not been reported. In early embryos, a switch in histone subtypes occurs at the 32- to 64-cell stage (Gabrielli and Baglioni, 1975, 1977). Some of the histone messengers for the early subtypes are present in the unfertilized egg and may contribute to a pool of histone used in male pronuclear remodeling. Longo et al. (1994) speculate that sperm protamines are released and exchanged for histones in vitro because exogenous histones or protamines in lysates prevent decondensation, which they suggest interferes with the removal of protamines. 4. Sea Urchins

The only biochemical data on male pronuclear transitions have been obtained from polyspermic sea urchin eggs. Sea urchin sperm nuclear proteins and transitions have been extensively reviewed (Poccia, 1987, 1989, 1995; Poccia and Green, 1992; Green et a!., 1995). Sea urchin sperm DNA is packaged exclusively with histones not protamines. Three of the histones are identical to somatic type: H2A, H3, and H4. Two special types, Sp H1 and Sp H2B, are found only in the sperm (and in modified forms during spermatogenesis and immediately after fertilization). They are first expressed in spermatogonia/spermatocytes(Poccia et al., 1989). Their phosphorylated forms are found up to the last steps of spermatogenesis when they become dephosphorylated (Poccia et a l . , 1987). There are usually two and sometimes three variants of Sp H2B. The Sp histones resemble their somatic counterparts in domain structure but contain additional sequences in the N-terminal regions (and for Sp HI in the C-terminal domain) (Poccia, 1987). These regions are highly basic and composed largely of serine-proline adjacent to two basic amino acids (lysine and/or arginine). The tetrapeptide has come to be known as the SPKK motif (Suzuki, 1989b). The tetrapeptide is repeated several times in tandem or nearly so in the N-terminal regions of Sp histones. Regions replete with SPKK motifs are predicated to have p-turn secondary structures (Green and Poccia, 1985; Poccia, 1987; Suzuki, 1989a) possibly coexisting with U-turns (Suzuki et al., 1993) or extended helical conformations (von Holt et al., 1984). These may play a role in their mode of binding to DNA. It has been proposed that SPKK regions in their unphosphorylated state stabilize and/or condense the sperm chromatin possibly through cross-linking of adjacent fibers (von Holt et al., 1984; Green and Poccia, 1985).

D. Poccia and P.Collas

3a

SPKK serines are sites of phosphorylation both during spermatogenesis (Poccia et al., 1987; Green and Poccia, 1988; Hill et al., 1991) and following fertilization (Green and Poccia, 1985). Sp histone phosphorylation occurs within minutes after fertilization, before extensive chromatin decondensation has occurred (Green and Poccia, 1985) or even if decondensation is inhibited (Poccia et al., 1990). The sperm H2A, H3, and H4 are not phosphorylated at this time (Fig. 2). The composition of the male pronuclear chromatin 5 min after fertilization (repertoire of variants, phosphorylation patterns) is very similar to that found during spermatogenesis. Since at least in their phosphorylated states during spermatogenesis, Sp variants are compatible with transcription, replication, or mitosis (Poccia et al., 1987), there is no need a priori to postulate a replacement of Sp histones to achieve reactivation of the chromatin in the male pronuclei. Nonetheless, further changes in histone composition and chromatin structure ensue before and during reactivation. The kinases catalyzing phosphorylation of SPKK sequences have not been identified. Sea urchin sperm are not known to contain Sp histone kinases. Egg cytoplasm has such kinases but whether they are the ones acting in vivo is not certain (Porter et al., 1989; Suzuki et al., 1990; Green et al., 1995). Although SPKK sites resemble those phosphorylated by cdc2 kinases, doses of the drug 6DMAP which block mitotic H1 kinase do not block Sp histone phosphorylation (Poccia et al., 1990). Sp H1 is removed from male chromatin during the first cell cycle, but the extent to which this occurs has only been estimated from polyspermic eggs and is variable (Poccia et al., 1981, Green and Poccia, 1985). Sp H2B remains in nucleosomes in polyspermic eggs and may be diluted during successive rounds of replication in the cleaving embryo or exchanged (Poccia et al., 1984). Early cleavage-stage embryos are, however, devoid of Sp H1 and Sp H2B (Cohen et al., 1975).

B

H3 w2A I

*%0

u4

Fig. 2 I n vivo remodeling of sea urchin sperm histones. (A) Male pronuclear histones 3 min after fertilization, and (B) sperm histones. N , phosphorylated Sp HI; O/P, phosphorylated Sp H2Bs. The earliest detectable changes in male pronuclei are specific modifications of Sp histones. Taken from G . R. Green and D. Poccia, unpublished.

2. Transforming Sperm Nuclei into Malc Pronuclei

39

The effect of phosphorylation is to decrease DNA binding as measured by affinity chromatography, precipitability, thermal denaturation, and dye binding competition assays (Suzuki 1989b; Hill et al., 1991; Green et al., 1993). It has been proposed that the loss of S p HI from the chromatin may occur after such weakening of DNA-protein interactions, possibly by competition with CS H1, a large HI histone stored in the egg that appears in the male pronuclear chromatin immediately following fertilization. In any event, no nucleoplasmin-like molecules have been reported in sea urchin that might serve to remove Sp histones. Additional proteins from the maternal pool are accumulated by the male pronucleus during and following modification of Sp histones. Sea urchin eggs contain stores of histones including CS H2A, CS H2B, H3", and CS H1 (CS standing for cleavage-stage). No variants of the late embryo or sperm (a,@, y, A, or Sp histones) are detected in unfertilized eggs. The maternal storage pool has been demonstrated by histone extraction from whole unfertilized eggs or enucleated egg halves, indicating that at least some of the pool is cytoplasmic (Poccia et al., 1981; Salik et al., 1981). It is not known if these histones are complexed with chaperone proteins. The maternal histones can assemble into male pronuclear chromatin in the absence of protein synthesis (Salik et at., 1981). The pool sizes have been estimated at >25 functional (assembly competent) haploid equivalents in eggs inhibited in protein synthesis and up to several hundred equivalents by direct extraction and separation by gel electrophoresis. Although considerably less histone is stored per egg than in frogs, the number of nuclei produced in an early sea urchin embryo is less by almost 1000-fold, and the egg is likewise about 1000-fold less in volume. Thus the concentration of histone and its ratio to cells at the blastula stage is comparable (Salik et at., 1981). While Sp histones are being phosphorylated, male pronuclei accumulate CS HI from the maternal pool (Poccia et al., 1981; Green and Poccia, 1985). Following these events there is a brief lag period with little change in male pronuclear chromatin composition. Then during replication, which begins at about 30 min postfertilization, the chromatin accumulates three other proteins stored in the egg: CS H2A, CS H2B, and an H3 species H3" (Poccia et al., 1981; Green and Poccia, 1985). CS H2A, H3', H3", and H4 but not CS H2B become phosphorylated during S phase, particularly CS H2A (Green and Poccia, 1989). Phosphorylation of CS H2A occurs on serines in the C-terminal region of the molecule and is enhanced when DNA synthesis is blocked with aphidicolin. Although the major accumulation of maternal histones from the storage pool into male pronuclei occurs during replication, replication is not required for replacement or augmentation of sperm histones. In eggs blocked with aphidicolin, CS H2A and CS H2B accumulation is slowed, but eventually they become the predominant histones of their classes in the chromatin. (Poccia et al., 1984). This implies that the CS forms must displace some of the preexistent sperm histones since under these conditions no new DNA is available for binding

40

D. Poccia and P. Collas

and reduction of nucleosomal spacing is not sufficient for additional histone binding. Eventually in monospermic eggs, Sp variants must be displaced, degraded, or highly diluted, since early embryo chromatin is devoid of these proteins (Cohen et at., 1975). The wealth of in vivo information available for the sea urchin has not yet been matched by in vitro results. Sp H1 and Sp H2B are phosphorylated in sperm nuclei added to cell-free egg extracts and enzymes capable of phosphorylation of purified histones and N-terminal fragments are clearly present in egg lysates (Green el al., 1995). Variable amounts of CS H1 become associated with sperm nuclei in vitro, but later changes have not been detected. Since the nuclei have not been shown to cycle, it is perhaps not surprising that histone assembly does not occur.

B. Summary and Speculations

Analysis of nucleoprotein transitions in the male pronucleus reveals many similarities in the four model systems despite the diverse sperm chromatin compositions. All four eggs have stores of maternal histones which are utilized for chromatin remodeling. Secondary modifications seem to play a role in adjustments, probably by modulating the affinity of the histones for DNA. The first signs of removal or modification of sperm proteins are seen within minutes of fertilization. The initial histone changes are prereplicative or replication independent. Male pronuclear histone composition is of a unique form, neither identical with the sperm nor with later embryonic nuclei which more closely resemble somatic chromatin in composition. The cleavage stage variants and HMGs may play a role in facilitating rapid replication in early embryos. In the maternal pool are unusual histone variants. CS H1 and H1 . X (B4) are larger variants of somatic H1. CS H2A of sea urchins, H2A.X of amphibians, and H2AP of Drosophila are larger variants of H2A. Such variants have also been reported in the dipteran Sciaru (Ruder et af., 1987) and the marine worm Urechis (Franks and Davis, 1983) and may be more common than yet demonstrated. Likewise it is not known how common is the storage of maternal histones with nucleoplasmin-like molecules or other chaperone proteins. H1 function in early embryos may not be typical as suggested by immunodepletion experiments. The mechanisms of protein remodeling may not be specific for species or cellular source of nuclei. Frog extracts can remodel human sperm nuclei (Itoh et al., 1993) or red cell nuclei (Blank et al., 1992). The role of nucleoplasmin as histone exchange and assembly factor and for decondensation has been well demonstrated for amphibians, but whether this function is more widespread is not known. At least in the case of sea urchins, histone exchange is not essential since the sperm lack protamines. In this case, phosphorylation seems to be the critical element for activation and chromatin decondensation.

2 . Transforming Sperm Nuclei into Male Pronuclei

41

III. Chromatin Decondensation Sperm chromatin DNA may be more densely packed than even mitotic chromosomal DNA. The density of DNA in protamine-containing mouse sperm chromatin has been estimated at almost - I pg/pm3 (Pogany et al., 1981) and in histone-containing sea urchin sperm chromatin at -0.2 pg/pm3, similar to mitotic chromosomes (Green and Poccia, 1985). Since such high degrees of chromatin condensation are usually incompatible with normal DNA reactions such as replication and transcription, sperm chromatin must be decondensed to function as pronuclear chromatin. Decondensation may be relatively uniform as in starfish (Longo and Schuetz 1982), molluscs and Ascaris (Longo, 1973). However, in many organisms it is not uniform, certain regions reproducibly decondensing late. In many invertebrates, decondensation is slower at specialized regions of the nuclear envelope. Decondensation may be progressive from periphery to interior (sea urchin) or from midregion to anterior and posterior poles (mammals) (see Longo, 1973). The pattern of male pronuclear decondensation does not necessarily resemble in reverse the condensation occurring during spermatogenesis. In rats and rabbits the patterns are similar; in sea urchins, mussels, and surf clams they are not (Longo, 1973). The ability to support sperm chromatin decondensation depends on the oocyte stage at fertilization. For example, if eggs are artificially fertilized or microinjected without activation of the egg at the germinal vesicle stage, sperm nuclei usually remain condensed. Sometimes the condensation state of the sperm chromatin mimics that of the maternal chromatin, undergoing meiotic maturation, swelling, and condensation in synchrony. In other cases the pronuclei appear to be regulated independently. In cases of physiological (natural) polyspermy, several sperm nuclei in a common cytoplasm may behave independently. Manipulation of living eggs and cell-free systems has helped to define conditions that promote decondensation and others which are not essential. Unfortunately, the relationship between chromatin composition and condensation state remains unclear. In general, cell-free systems mimic in vivo behavior closely.

A. Conditions Promoting Decondensation in Vivo

1. Amphibians The female chromatin completes the second meiotic division in Xenopus eggs at about 15 min postfertilization (Graham et al., 1966). The second polar body is extruded at 20 min as the female pronucleus swells and moves to the egg center. The sperm nucleus is condensed at 10 min postfertilization but then begins to swell and move toward the egg center. By 40 min, the two pronuclei are adjacent

42

D. Poccia and P. Collas

with intact nuclear envelopes. DNA synthesis occurs in both nuclei simultaneously at about 20-30 min. The chromatins only associate at 60-70 min when the nuclear envelopes break down. The sperm nucleus swells to about 20 p.m (a 50-fold increase in volume) by -30 min after fertilization (Gurdon and Woodland, 1968). Fertilization of toad eggs in the germinal vesicle stage results in failure of the male pronucleus to develop (Katagiri, 1974). This is probably accounted for by the absence of germinal vesicle material in the cytoplasm. Swelling occurs when nuclei are microinjected into the oocyte nucleus or egg cytoplasm, but not in the cytoplasm of enucleated oocytes unless a soluble fraction of oocyte nuclear extract is added back (Lohka and Masui, 1983b). Xenopus sperm responds to Rana mature egg cytoplasm by forming male pronuclei or mitotic chromosomes so controlling factors are not species specific (Fig. 3).

2. Fruit Flies In Drosophila, the condensed sperm chromatin undergoes changes in condensation during oocyte maturation. Supernumerary male nuclei in the same cytoplasm disintegrate. The male pronucleus forms an aster during the second meio-

,

*.

8 *

f

..

Fig. 3 Behavior of Xenopus sperm nuclei after microinjection into mature Rana pipiens oocytes. (a) Intact sperm, (b) partially decondensed male pronucleus, (c) complete decondensed male pronucleus, and (d) mitotic chromosomes formed from sperm nuclei. Taken from Lohka and Masui (1983a) with permission.

2. Transforming Sperm Nuclei into Male Pronuclei

43

tic division. Its chromatin decondenses after completion of the meiotic divisions of the female and then both pronuclei approach side by side in the interior of the egg, each spherical and similar in size. Both sets of chromosomes condense simultaneously and are arranged on the first spindle separately (Huettner, 1924). The tail remains attached to the male pronucleus throughout (Karr, 1991).

3. Surf Clams The relationship between the male and female pronuclei in Spisulu solidissirnu is complex. Fertilization triggers germinal vesicle breakdown within 12 min, followed by two meiotic divisions and formation of the female pronucleus by 45 min. The outlines of pronuclear transitions have been established in its relatively transparent living egg stained with a DNA vital stain (Luttmer and Longo, 1986). In the germinal vesicle, 19 bivalent chromosomes are apparent. Following fertilization, the sperm nucleus remains unswollen for 10 min while germinal vesicle breakdown initiates. The maternal chromosomes move toward the center of the egg, while the sperm chromatin begins to decondense. By 20 min, the maternal chromosomes move to the cortex in meiotic metaphase I. Meiotic metaphase I1 follows quickly and formation of the second polar body is completed by 40 min postinsemination. Decondensation of the female pronucleus is paralleled at this point by the male pronucleus so that both approach the same size. At 50 min, the pronuclei migrate to the egg center, and the chromosomes condense in the first mitosis. At the electron microscope level, the sperm head is sometimes seen to rotate in the fertilization cone and by 3 min is well incorporated into the egg (Longo and Anderson, 1970). The sperm chromatin with no nuclear envelope boundary begins to disperse. Inner and outer zones of chromatin can at first be distinguished by condensation state, but by 6-15 rnin the male chromatin becomes uniformly dispersed like the maternal. After reformation of a nuclear envelope, the male nucleus undergoes further enlargement becoming ellipsoidal. The two pronuclei approach following meiosis, their nuclear envelopes vesiculate, and chromosomes condense at the first mitosis. Clam male pronuclear transitions have been divided into four phases (Luttmer and Longo, 1988): (A) no changes before GVBD at 15 min, (B) moderate expansion during germinal vesicle breakdown at 15-20 min, (C) condensation during polar body formation at 20-40 min, and (D) major expansion after 40 min. Using a variety of treatments to perturb pronuclear progression such as temperature, pH, and inhibition of microtubules, protein synthesis, and metabolism, a strong correlation was made between the state of the maternal chromosomes and the state of male pronuclei. Swelling at phases B and D is differentially affected, suggesting that these stages of expansion might be regulated by different factors. Na+ is needed for nuclear enlargement, but Na+ deprivation cannot be reversed by increasing internal pH with ammonia, implying the effect

44

D. Poccia and P. Collas

is not mediated by a Na+/H+ exchange. Phase D is insensitive to pH control but ATP-dependent. Protein synthesis appears to be required for phase D, but not B. Polyspermy results in less male pronuclear expansion in vivo, suggesting that factors controlling pronuclear swelling may be present in limiting amounts (Luttmer and Longo, 1988). Blocking protein synthesis delays swelling in phase D but this effect can be overcome by elevating intracellular Ca2+ levels with ionophore A-23187 (Longo et a l . , 1991). The DNA topoisomerase I1 inhibitor teniposide has no effect of sperm chromatin decondensation in surf clam eggs at concentrations which inhibit mitotic chromosome condensation (Wright and Schatten, 1990). 4. Sea Urchins

The sea urchin egg is fertilized after completion of meiosis, the female pronucleus having already formed. Events can be monitored by electron microscopy or in living eggs with vital staining of nuclei (Luttmer and Longo, 1987). In Arbacia, the sperm nucleus rotates in the fertilization cone at 2-3 min. The chromatin, unbound by a nuclear envelope, begins to disperse characteristically from the perimeter towards the core (Longo and Anderson, 1968). This results in a heart-shaped nucleus (appearing ovoid in the light microscope) which gradually acquires a spherical shape (Fig. 4). Three regions of chromatin can be distinguished during decondensation: a condensed core (CC), a coarsely aggregated region (CDC), and finely dispersed region (FDC) extending from inside to periphery, respectively. Membrane vesicles appear surrounding the decondensing chromatin and fuse. The male pronucleus migrates toward the larger female pronucleus, becoming spherical. The male pronuclear chromatin may not be completely decondensed at this point reflecting how much time has occurred before fusion, this depending on the site of sperm entry relative to the position of the female pronucleus. The nuclei may swell to 10 pm in diameter, a volume increase of about 20-fold. Upon fusion, the more highly condensed male chromatin is easy to discern, but soon after it cannot be distinguished from the finely dispersed female chromatin. The processes up to fusion may take 15-20 min and are normally completed before DNA synthesis ensues at about 30 min. Conditions promoting decondensation have been studied in fertilized eggs and in eggs microinjected with isolated sperm nuclei. Fertilization of oocytes at different stages of maturity demonstrates that not all cytoplasmic states promote decondensation (Longo, 1978). Although all stages are fertilizable, no male pronuclear decondensation occurs in germinal vesicle stages, and only limited decondensation has been observed in eggs undergoing meiotic maturation. In none of these cases does a nuclear envelope form. Once acquired, the conditions promoting decondensation persist in embryos as shown by reinsemination experiments (Longo, 1980).

2. Transforming Sperm Nuclei into Male Pronuclei

45

Fig. 4 Heart-shaped sea urchin male pronucleus forming in vivo. PNE, pronuclear envelope; C, centriole; SF, sperm flagellum; CC, condensed chromatin; CDC, coarsely dispersed chromatin; FDC, finely dispersed chromatin; single and double-stemmed mows, male pronuclear remnants at acrosoma1 and centriolar fossae, respectively. Nuclear envelope remnants have been incorporated into the new envelope. Taken from Longo and Anderson (1968) with permission.

Mature egg (ootid) cytoplasm promotes decondensation. The rate of enlargement of the male pronucleus in fertilized mature eggs (estimated by crosssectional area) is linear (therefore the volume increase is a power function), while the female nuclear volume remains constant. As in surf clams, the rate of enlargement is slower in polyspermic eggs, suggesting limiting factors stored in the egg (Luttmer and Longo, 1987). The length of S-phase and the first cell cycle are also increased at > 15 male pronuclei/egg (Poccia et al., 1978). Nuclear swelling is unaffected by colchicine, cytochalasin, or inhibition of protein synthesis, but slowed or prevented by low temperature and by metabolic inhibitors such as azide, cyanide, and oligomycin, consistent with a requirement for ATP (Luttmer and Longo, 1987). Fertilization in Naz+-free sea water, which prevents the cytoplasmic pH rise due to H+/Na+ exchange but not the increase in intracellular Ca2+, blocks decondensation, which can be restored by transfer to normal sea water (Canon and Longo, 1980). Microinjection experiments, which bypass sperm-initiated surface events resulting in egg activation, confirm and extend observations on living monospermic eggs (Cothren and Poccia, 1993). Microinjected permeabilized sperm nuclei only partially decondense in unfertilized eggs. This block is completely relieved by activating eggs with ammonia, raising the internal pH. Under these conditions the eggs show no signs of activation of signalling pathways which result in

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increases in internal Ca2+ such as the inositol triphosphate pathways. No decondensation occurs in microinjected immature (germinal vesicle) stage oocytes, whether they are unfertilized or fertilized, Ca2+-ionophore or ammonia activated. The results confirm a two-phase decondensation process, the first resulting in partial decondensation, which develops during meiotic maturation and can be blocked by the kinase inhibitor 6-DMAP, and the second normally set in motion by fertilization but only requiring cytoplasmic alkalinization. A comparison of results from fertilized and microinjected eggs is shown in Fig. 5.

Primary Oocyte

Unfertilized Egg

Fertilized Egg

Fertilized Oocyte

Unfertilized Egg

Fertilized Egg

Fertilized Oocyte

Unfertilized Egg

Fertilized Egg

Fig. 5 Behavior of fertilizing and microinjected sperm nuclei in living sea urchin eggs. Diagrammatic summary of data from Cothren and Poccia (1993). Microinjected (lower) or fertilizing (upper right) nuclei in primary oocytes (column I), unfertilized eggs (column 2). and fertilized eggs (column 3). Maternal nucleus is shown in the upper left of each egg. DMAP, egg pretreated with protein kinase inhibitor 6DMAP; A23187, egg activated with Ca2+ ionophore; pH, partially activated eggs treated with ammonia to raise intracellular pH.

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B. Conditions Promoting Decondensation in Vitro 1. Amphibians The definitive system for studying male pronuclear decondensation and development was invented by Lohka and Masui (1983a) and has found widespread use as a cell-free system for studying nuclear transformations (Fig. 6). They found that both cytosol and membranous fractions are needed for full decondensation. The particulate or membrane-containing fraction provides a nuclear envelope precursor population. Decondensation occurs in two phases: a rapid decondensation which can take place in cytosol (S150) followed by a slower membrane-dependent swelling. The initial phase (from a compact coiled filamentous structure to an elongated snakelike object threefold longer) occurs within 1-10 min and depends on nu-

Fig. 6 Formation of male pronuclei from Xenopus sperm nuclei in Xenopus egg extracts. (A) Inputpermeabilized sperm nuclei; bar, 1 pm. (B) Decondensed sperm chromatin incubated in egg extract for 5 min; bar, 1 pm. (C) Male pronucleus with nuclear envelope, 180 min of incubation; bar, I prn. (D) Detail of pore-containing nuclear envelope around decondensed chromatin, 120 min of incubation; bar, 0.5 pm. Taken from Lohka and Masui (1983a) with permission.

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cleoplasmin (Philpott et al., 1991; Ohsumi and Katigiri, 1991a). High-speed extracts of Xenopus or Rana imrnunodepleted of nucleoplasmin do not support decondensation (Fig. 7). Activity is restored by addition of purified nucleoplasmin which, unlike other polyanions, is sufficient at physiological concentrations to promote decondensation. These observations can at least partially explain the lack of decondensation of sperm nuclei in immature oocyte cytoplasm in vivo, since nucleoplasmin is stored in the germinal vesicle during oogenesis, becoming phosphorylated upon activation and germinal vesicle breakdown. There is sufficient cytoplasmic activity in mature eggs to decondense up to 30,000 nuclei per egg. Polyglutamic acid can induce decondensation of amphibian sperm but unlike nucleoplasmin also removes core histones (Katagiri and Ohsumi, 1994). The mode of action of nucleoplasrnin appears to involve nuclear protein removal and exchange rather than degradation because intact protamines are coprecipitated with nucleoplasmin by anti-nucleoplasmin antibodies (Philpott and Leno, 1992). Assembly of histones and removal of protamines imply a dual role for nucleoplasmin. It has been suggested that nucleoplasmin binds to a

MOCK

+NPL

-NPL

FRESH

Fig. 7 Nucleoplasmin decondenses Xenopus sperm chromatin. Xenopus sperm nuclei incubated for 10 min with Xenopus egg extract (a) mock depleted, (b) nucleoplasminimmunodepleted, (c) nucleoplasmin depleted with added purified egg nucleoplasmin before nuclear addition, and (d) control incubated sperm. Bar, 50 pm. Taken from Philpott er al. (1991) with permission.

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specific domain rather than acting by simple charge shielding mechanisms (Katagiri and Ohsumi, 1994). Topoisomerase I1 probably helps organize DNA loops in chromatin and may be involved in male pronuclear decondensation. Nuclear swelling can be blocked with topoisomerase I1 inhibitors VM-26 (Newport, 1987) and ICRF-193 (Takasuga et al., 1995), suggesting sperm nuclei must be capable of responding to decondensation factors by some sort of unwinding or decatenation. The latter drug, unlike the former, introduces no breaks in the DNA that might recruit the DNA repair system to further complicate the interpretation. ICRF-193 blocks swelling or dispersion of chromatin but not the initial decondensation phase. However, alteration of chromatin composition and nucleosomal spacing remain unaffected (Katagiri and Ohsumi, 1994). Even though these nuclei are rather condensed, they are capable of replication, though at a slower rate. Amphibian cytoplasm can decondense human sperm if the mammalian protamines are first disulfide reduced (Itoh et al., 1993). Thus the factors involved are not species specific. It has long been established that nuclei such as erythrocyte nuclei decondense in amphibian egg cytoplasmic extracts (Barry and Merriam, 1972). Erythrocyte nuclear decondensation occurs in two phases reminiscent of sperm decondensation. H5, H2A, and H4 are phosphorylated and swelling requires ATP in the first phase during which time the HI variant H5 is released (Blank et al., 1992). In the second phase, which also requires ATP, lamin L,,, is incorporated, and DNA synthesis initiates. Thus the mechanisms for decondensing chromatin are not cell type-specific to the nuclear source in vivo or in vitro.

2. Fruit Flies Embryo extracts decondense Xenopus sperm nuclei in two phases. Phase 1 occurs rapidly, but phase 2 occurs very slowly in the heterologous system (Kawasaki et al., 1994). Decondensation in phase 1 can be achieved by soluble factors that are heat resistant and resistant to the alkylating agent N-ethylmaleimide (NEM); phase 2 requires membranes (Kawasaki et al., 1994) Decondensation is complete by about 30 min (Berrios and Avilion, 1990). Other studies indicate the swelling may occur in multiple phases, with initial enlargement followed by a temporary condensation and then further swelling (Ulitzur and Gruenbaum, 1989). These phases are: ( 1 ) condensed, 0-5 min; (2) decondensed, 5-10 min; (3) recondensed, occurring in about 50% of the nuclei, 3040 min; and (4) swollen interphase-like thereafter. Swelling requires ATP and can be inhibited with the topoisomerase I1 inhibitor novobiocin up to but not in the fourth phase (Ulitzur and Gruenbaum, 1989). An attempt to isolate a nucleoplasmin-like decondensing factor from flies resulted in purification of a 22-kDa heat-stable protein immunologically distinct from frog nucleoplasmin (Kawasaki et al., 1994). The protein is present in

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embryo nuclei throughout development and is sufficient for decondensation of Xenopus sperm nuclei. However, if it is immunodepleted from Drosophilu extracts, decondensation still occurs, suggesting the presence of more than one heat-stable decondensing activity in flies.

3. Surf Clams Decondensation in vitro resembles events observed in vivo. In unactivated and 4-min extracts, there is an unexpected slow increase in decondensation (since the germinal vesicle is not yet broken), but in later extracts much faster and more extensive swelling is promoted (Longo et ul., 1994). In 15-min extracts, 90% of the nuclei swell but maximal sperm head enlargement in vitro occurs in 65 min postmeiotic cytoplasm at spermiegg ratios of 1. The last chromatin to decondense is in the middle portion of the nucleus associated with the sperm nuclear envelope in the region subjacent to the acrosome. Decondensation is inhibited at low temperature and requires cytoplasm. Lysed germinal vesicles have no effect on swelling when added to the lysates. EDTA, EGTA, histone, protamine, and 6DMAP block the percentage and extent of decondensation in vitro, with most effects being reversible. These results were interpreted to implicate histone/protamine exchange and Ca2+ and phosphorylation requirements in decondensation. An involvement of nucleoplasmin has not been established in surf clam. An abundant nucleoplasmin-like phosphoprotein of 49 kDa on SDS gels has been isolated from surf clam germinal vesicles, but its effects on decondensation or chromatin assembly were not reported (Herlands and Maul, 1994).

-

4. Sea Urchins

Sea urchin nuclei decondense in the system of Zhang and Ruderman (1993) to small diameter spheres. Only 5-10% of Xenopus sperm nuclei in this system were well swollen, with others partially decondensed or irregularly shaped. Only those fully enlarged showed signs of replication (see below). Cameron and Poccia (1994) reported swelling of 100% of the sea urchin sperm nuclei occurring through morphological phases resembling those in vivo (Cothren and Poccia, 1993). Two phases of enlargement can be distinguished in vitro: a membrane-independent decondensation and a membrane-dependent swelling (Collas and Poccia, 1995b). The first phase occurs in cytosol devoid of membranes, converting the conical -1.5 X 4-ym nucleus through a transient ovoid intermediate to a sphere of -4 ym. This transformation requires ATP hydrolysis provided by addition of ATP and an ATP-generating system and cytosol (S100) and is inhibited by the protein kinase inhibitors 6DMAP and staurosporine (Cameron and

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Poccia, 1994). Decondensation is favored at alkaline pHs and in activated egg cytoplasm. If membranes are present, the nucleus will bind membrane vesicles and if GTP is provided they will acquire a nuclear envelope (lacking lamins). The second phase requires additional input of ATP, cytoplasmic lamins, and cytosolic factors sensitive to heat and NEM. The Ca2+ chelator BAPTA also blocks swelling. The rate and extent of swelling is similar to that in vivo. Final diameters are limited by suboptimal amounts of membrane vesicles and ATP or by depletion of lamins from the extract (Collas et a l . , 1995). Thus the first phase (chromatin decondensation) seems to be a property of the chromatin and cytosol; the second phase (swelling of the nucleus) seems to require a complete nuclear envelope with lamina and may be driven by lamina1 growth or import of karyophilic proteins after functional nuclear envelope formation.

C. Summary and Speculations

In vivo, decondensation of the sperm chromatin depends on the state of the maternal cytoplasm, but it is not always coordinate with the female or other male chromatin sharing the same cytoplasm. This presents an unsolved problem for interpretations based on common cell cycle signals inducing condensation. The contents of the germinal vesicle seem to be required for decondensation and at least one of these has been identified in amphibians, nucleoplasmin. It is not yet clear if nucleoplasmin-like molecules function in all organisms, however. There is in many cases a lack of species specificity to the decondensation. In some organisms, decondensation occurs from the periphery to the center, suggesting factors operating from outside to inside. Alternatively this may be a consequence of the mode of packing of the chromatin during fertilization. For example, late decondensation has been associated with specialized nuclear envelope structures (Yanagimachi and Noda, 1970; Ward and Coffey, 1989). Since some pronuclear chromatins disperse uniformly, they are unlikely to be limited by diffusion of decondensing factors. Whatever the pathway, in all cases the final state achieved is of chromatin much less condensed than sperm chromatin, a state more compatible with reactivation of nuclear activities. The transition to the fully decondensed state always seems to traverse more than one phase. Demembranated sperm chromatin is capable of an initial decondensation phase which can be completed only after the nuclear envelope is formed. The first phase of enlargement may be the result of removal of constraints laid down during spermatogenesis when special proteins package the chromatin into a unique compacted form, whether by removal of protamines, phosphorylation of histones, or other alterations. The second may depend less on the intrinsic tendency of chromatin to swell, to some degree

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driven by DNA backbone charge repulsion, and instead on the active participation of a growing envelope or import of karyophilic proteins and water. Superimposed on this pattern typical of amphibians and sea urchins are more complex changes whose timing depends on maturation and associated condensation signals operating on maternal chromosomes in organisms like clams and flies fertilized at earlier stages of meiosis. It is important to realize that sperm nuclei, even though condensed, are interphase nuclei when they enter the egg. Likewise their chromosomal protein composition is not identical to that of the maternal chromosomes. Thus maternal and paternal chromosomes may very well respond to common signals in different ways. This explanation does not apply to the problem of differential development of multiple male nuclei in physiological polyspermy, however. Among specific molecules likely to affect higher order chromatin transitions in the decondensing male pronucleus are H1 histones, HMG proteins, and topoisomerse 11. The relationship between chromosomal composition and condensationldecondensation as well as the nature of the signals regulating higher order chromatin structure in male pronuclei or mitotic chromosomes remain areas in need of fresh insight and investigation.

IV. Formation or Adjustment of Nucleosomes Somatic chromatin is organized into nucleosomes. A nucleosome contains two each of the histones H2A, H28, H3, and H4 which protect 146 bp of associated DNA. These are connected by a variable amount of linker DNA, depending on cell type, which is associated with histone H 1 . H1 protects an additional -20 bp of DNA against micrococcal nuclease digestion (- 166 bp) defining the chromatosome. Variable linker gives rise to variable average distances between nucleosomes or repeat lengths (up to 250 bp). The significance of variable repeats is not known, although short repeats tend to be found in active chromatins, perhaps as a result of biasing the distribution by regions where genes are active and structure is disrupted. All nucleosomal structure is lost in those sperm in which protamines completely replace histones. In sperm containing only histones, chromatin may have repeat lengths similar to somatic (such as goldfish) or different (such as in sea urchins or sea stars). For protamine sperm, reestablishing nucleosomal organization restores a repeat length; for histone sperm, the change from paternal to zygotic histones is likely to alter repeat length. Repeat length may depend on histone subtypes or secondary modifications. Histone H1 which binds to linker DNA is often considered a contributor to or determinant of repeat length. Alternatively, nonhistone proteins such as HMGs (high-mobility group proteins, chromatin nonhistone proteins with characteristic conserved amino acid sequences associated with active chromatin) may alter

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repeat lengths. Histone subtype changes are not necessarily accompanied by alterations of the repeat as during sea cucumber spermatogenesis (Cornudella and Rocha, 1979). Repeat length alterations may also be regulated by changes in histone secondary modification rather than subtype as in sea urchin spermatogenesis (Green and Poccia, 1988). Although in Xenopus and Drosophila one can reconstitute naked DNA with histones from ooplasmic extracts to form somatic-type repeat lengths, the only case in which in vivo alterations in male pronuclear repeat length have been measured is the sea urchin; the only system in which in vitro changes have been well documented is amphibians.

A, Amphibians

Chromatin has been assembled from DNA and histones in high-speed extracts of amphibian eggs (Rodriquez-Campos et al., 1989; Shimamura et al., 1989; Zucker and Worcel, 1990). This reaction requires ATP, Mg*+, an ATP-generating system, and S-150. In the presence of topoisomerase I or I1 and all four histones, the DNA forms a supercoiled chromatin, but the nucleosomes from this purified system are close packed. The 180-bp repeat can be altered by addition of H1, which increases the spacing up to 200 bp. An assembly activity was partially purified from Xenopus oocytes using such a system. It organizes regularly spaced nucleosomes with a repeat length of 165 bp and requires ATP (Tremethick and Frommer, 1992). If H1 is added it increases the repeat to 190 bp. HMG 14 and HMG 17 are present in extracts containing spacing activity (Tremethick and Drew, 1993). Adding back phosphorylated HMG 14 and 17 without H1 to histone cores is sufficient to achieve the short repeat of 160-165 bp. At least some of the sperm chromatin of Xenopus has a 180-bp repeat length, but chromatosome or core length DNAs do not accumulate upon micrococcal nuclease digestion (Dimitrov et al., 1994). This suggests that the sperm chromatin is not in a normal somatic-type configuration. (In contrast, Ohsumi et at., 1993, report chromatosome length protection in Xenopus sperm.) However, when male pronuclei form in vitro, clear core nucleosomal (146 bp) and chromatosomal lengths (168 bp) are generated upon digestion. In vitro male pronuclei have 180-200 bp repeat lengths (Dimitrov et al., 1994; Philpott and Leno, 1992). According to Dimitrov et al. (1994) core protection requires H2A and H2B assembly on the depleted sperm chromatin; chromatosome assembly requires the H1 protein B4. On the other hand, immunodepletion of B4 has no effect on generating repeat length spacing of 180 bp in vitro, so B4 appears to act as a linker histone in providing extra protection beyond the core, but is not responsible for adjusting repeat length. Ohsumi et al. (1993) report immunodepletion of H1.X (B4) does not interfere with the generation of a normal

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repeat length of 200 bp (or chromosome condensation) and, in the absence of another unidentified H1 subtype, would again suggest that an H1 is not necessary at all for generating a repeat pattern. Dephosphorylation of histones with phosphatase does not block histone assembly and remodeling and so this modification is apparently not required for chromatin assembly or spacing (Dimitrov et al., 1994). Dimitrov et al. (1994) do not find HMG14 or HMG 17 in pronuclear chromatin so rule out these as controlling repeat length, but do find substantial amounts of HMG 2 that may function in this way.

B. Fruit Flies

It has long been established that Drosophila embryo extracts can assemble closepacked nucleosomes on circular DNA (Nelson et d.,1979). Additionally, chromatin can be reconstituted with correct nucleosomal spacing from Drosophila embryo extracts using somewhat different techniques (Becker and Wu, 1992; Becker et al., 1994). S1.50 extracts from 0- to 90-min embryos will assemble chromatin from the maternal histone pool which lacks a typical H 1. If exogenous H1 is added, nucleosome repeat length increases from 188 to 200 bp. This reconstitute is transcriptionally repressed.

C. Sea Urchins

Since sea urchin sperm chromatin already arrives in the egg in a nucleosomal configuration, there is no need to establish a new structure. On the other hand, mature sea urchin sperm chromatin has the longest repeat length known (about 240-250 bp) (Keichline and Wasserman, 1979; Savic et al., 1981; Vodicka et al., 1990). During spermatogenesis, precursors to the spermatozoon have repeat lengths of 234 bp (Green and Poccia, 1988). In early embryos the repeat is only 195-205 bp at the four- to eight-cell stages (Savic et al., 1981). In vivo, the male pronucleus reestablishes the more typical somatic type repeat soon after fertilization. For the first 30 min, the chromatin repeat length remains essentially unchanged (Savic et al., 1981). Thus the change in repeat length is not tightly coupled to phosphorylation of the Sp histones which is completed within 2-3 min (Green and Poccia, 1985) or to the acquisition of CS H I . During replication which begins at 30 min, male pronuclear repeat lengths decline to levels approximately those of somatic cleavage stage nuclei (-210 bp). The decline in repeat length is slowed at high degrees of polyspermy, conditions in which the cell cycle, including S-phase, lengthens. Replication is not necessary for the decline in repeat length, however. In the presence of aphidicolin, which blocks DNA synthesis, or emetine, which blocks protein synthesis, the decline is

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retarded but still occurs. Since accumulation of CS core histone variants is also retarded, it was suggested that the change of these histone subtypes might control changes in repeat length (Poccia et af., 1984). In summary, although large changes in nucleosomal repeat length can occur in male pronuclei (or in spermatids differentiating into spermatozoa), neither controlling factors nor the significance of repeat length alterations are known. Increased linker length is correlated with dephosphorylation of SPKK regions of Sp histones during spermiogenesis in sea urchins, but whether histones drive the increase or merely bind to extra DNA made available is not apparent. Dephosphorylation of these histones in male pronuclei in vivo does not result in immediate reestablishment of short repeats. Although it is often assumed that the type of histone H1 or its state of phosphorylation is related to repeat length, experiments using sperm nuclei in immunodepleted frog lysates and in vitro reconstitution assays with naked DNA suggest that this may not always be the case. Of course, assembly reactions onto chromatin or onto naked DNA may not proceed by identical pathways, and adjustment of repeats taking place during replication may also differ from these nonreplicative assemblies.

V. Nuclear Envelope Disassembly and Assembly The sperm nuclear envelope typically lacks pores, reflecting its inactive state. This unusual nuclear envelope, however, is rapidly removed upon entry of the nucleus into egg cytoplasm. It is replaced by an envelope largely or entirely of maternal origin. Since DNA replication depends on the reconstitution of a nuclear envelope, this replacement is crucial to cell cycle progression. Envelope reformation along with chromatin decondensation mark the full transition of the sperm nucleus to a male pronucleus. All in virro systems mimic disassembly of the sperm nuclear envelope by permeabilization of input sperm nuclei with lysolecithin or nonionic detergents without which subsequent development is halted. Under conditions reported so far, egg cytoplasm does not appear to efficiently remove sperm nuclear envelopes. This may be due to the envelopment of the nuclei by resealed plasma membranes during isolation.

A. Removal of the Sperm Nuclear Envelope and Initiation of Nuclear Envelope Formation

Relatively little attention has been paid to the conditions of sperm nuclear envelope disassembly. It is likely that disassembly takes place rapidly and virtually completely in most organisms, but few have been carefully studied (see Longo, 1973). Rapid disassembly is characteristic of the four model organisms. Disassembly of nuclear envelopes during mitosis appears to require lamin phospho-

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rylation under the control of mitotic kinases. Whether similar mechanisms operate on the sperm nuclear envelope is not clear. There are conflicting data on whether nuclear envelope assembly is initiated at preferred sites or essentially randomly or whether it requires lamins.

1. Fruit Flies Nothing is known about the control of disassembly or reassembly of Drosophila sperm nuclear envelopes, since in vitro studies of Drosophila embryo extracts have used frog or chicken sperm. Studies of the nuclear envelope in embryonic cells of Drosophila reveal some unusual aspects of this process and it is possible that male pronuclear envelope formation in flies would be atypical. In mitotic embryonic cells of Drosophila, breakdown and reformation of the nuclear envelope does not involve complete disassembly. The chromosome periphery at mitosis comprises a structure surrounding each chromosome except in the region of the centromere which contains a variety of nuclear matrix proteins, nuclear and nucleolar proteins, and RNPs (Hernandez-Verdun and Gautier, 1994). Monoclonal antibodies against nuclear envelope antigens reveal particulate structures remaining near the chromosomes in prophase and metaphase (Fuchs et al., 1983). In telophase, nuclear envelope antigens begin to assemble near the chromosomal centromeres. It was suggested the centromere region constitutes a nuclear envelope organizing region. Lamin A binds first to the poles in regions depleted of perichromosomal material in telophase so its appearance is complementary to the perichromosomal material. Thus there is an apparent polarity to formation of the nuclear envelope in Drosophila somatic cells.

2. Surf Clams

In vivo, a small fertilization cone forms at the site of sperm entry and here the nuclear envelope of the sperm, which lacks pores, becomes vesiculated (Longo and Anderson, 1970). Sometimes the nucleus rotates in the cone. By 3 min, the nucleus, mitochondria, and centrioles are incorporated into the egg; the flagellum is only rarely seen. Vesicles initially associated with the male chromatin increase in number and then apparently fuse. The sperm nuclear envelope in surf clams vesiculates just before the first expansion of the sperm chromatin (Longo and Anderson, 1970). Factors controlling this loss are not known. Disassembly of the clam germinal vesicle envelope during meiotic maturation has been studied in vitro. Extracts of meiotic cells will disassemble the germinal vesicle nuclear envelope following phosphorylation of their 67-kDa lamin as occurs in vivo (Dessev and Goldman, 1988; Dessev et al., 1989). The system requires ATP and Mg2+ but not Ca2+. No effects of protease inhibitors are detected but the system is sensitive to dephosphorylation by alkaline phosphatase. Since the new male pronuclear envelope (and sperm aster) form only after

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the egg completes meiosis, if similar mechanisms operate for the male pronucleus, the two would have to be independently regulated. Some regions of the sperm nuclear envelope may escape disassembly in clams (Longo et al., 1994). Retention of the sperm nuclear envelope in the nuclear midregion associated with the acrosomal fossa is observed in vitro (Fig. 8). Although as yet undocumented in vivo, the nuclear envelope remnants have been suggested to be sites of chromatin-envelope interaction and potential nucleating sites for male pronuclear envelope assembly.

3. Sea Urchins At fertilization, the sperm nucleus enters a fertilization cone, a protrusion of egg cytoplasm containing ribosomes, vesicles, and filaments (Longo and Anderson,

Ag. 8 Nuclear envelope remnants of surf clam male pronucleus forming in vitro. Surf clam sperm nucleus taken 30 rnin after incubation in 15 niin activated oocyte extract. E, the portion of the sperm nuclear envelope which remains intact after permeabilization in lysolecithin. A, acrosome. Chromatin fibers (arrows) emanate from E. Bar, 0.5 pm. Taken from Longo e t a / . (1994) with permission.

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1968). Here within 2-3 min the nucleus rotates 180" from its initial orientation perpendicular to the egg surface so that the base of the conical nucleus points toward the center of the egg. The sperm nuclear envelope, lacking pores, is immediately disassembled, presumably through fusion of the inner and outer membranes. Even when disassembly is complete, two regions at the nuclear poles retain the nuclear envelope. The portions of the sperm nuclear envelope lining the acrosomal and centriolar fossae appear to be associated with electron dense cup-like structures and these envelope remnants persist and are subsequently incorporated into the male pronuclear envelope. These observations are mimicked in vitro. Isolated sperm nuclei permeabilized with either lysolecithin or 0.1% Triton X- 100 lose all nuclear envelope except for remnants at the acrosomal and centriolar fossae (Collas and Poccia, 1995a) that correspond to those portions retained in vivo (Longo and Anderson, 1968). The remaining membrane consists of a cup-shaped region surrounded by an osmiophilic thicker cup which also shows signs of membranous elements by electron microscopy (Fig. 9). It is to these poles that initial binding of cytoplasmic membrane vesicles is targeted in v i m (Fig. 10). Binding then continues toward the equator until the nucleus is covered with vesicles. Upon addition of GTP, the vesicles fuse with one another and with the lipophilic material at the poles as judged by separately labeling the structures with lipophilic fluorescent dyes emitting different wavelengths. It is unclear whether polarized binding occurs in vivo since the electron microscopy studies may be inconclusive due to the rapid kinetics in vivo and absence of three-dimensional reconstruction (Longo and Anderson, 1968). Extraction of 0.1% Triton X-100 washed nuclei with 1.0% Triton X-100 removes the polar lipophilic structures but leaves much of the thicker osmiophilic cup. The solubility characteristics of the polar material are not expected of

Fig. 9 Nuclear envelope remnants of sea urchin male pronuclei forming in vitro. Decondensed sperm nucleus made in ATP-depleted S100-containing membrane vesicles. (A) Lack of vesicle binding in the absence of ATP. Fossa lined with osmiophilic material. (B) Enlargement showing two views of the osmiophilic cup (arrowheads) associated with membranous material (arrows). Taken from Collas and Poccia (1995b) with permission.

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Fig. 10 Polarized membrane vesicle binding around decondensing sea urchin sperm chromatin in vitro. Membranes vesicles of cytoplasmic extracts were labeled with the fluorescent lipophilic dye DHCC. Vesicles bind initially at the two poles of the conical nucleus and then progressively around the perimeter. Time of incubation is indicated. Taken from Collas and Poccia (1995b) with permission.

normal membranes and may be due to either an unusual lipid composition or association with proteins. The stripped nuclei do not bind cytoplasmic membrane vesicles in vitro. If the lipophilic structures are added back to the stripped nuclei they reassociate specifically to the poles, probably targeted to the material of the osmiophilic cups, with a preference first for the acrosomal pole. Unipolar reconstitutes bind membrane vesicles only at the acrosomal pole. Bipolar reconstitutes bind membrane vesicles initially at both poles and direct complete nuclear envelope formation as assessed by exclusion of 150-kDa dextrans from the nuclear interior. Thus structures at both poles seem to be required for complete envelope formation. Binding to the nucleus of lipophilic material is sensitive to low levels of protease digestion of either. Washes of lipophilic material in 0.9 M KCl abolish binding, suggesting binding is mediated by loosely associated proteins (P. Collas and D. Poccia, 1996a). Binding of cytoplasmic membrane vesicles to the polar lipophilic structures is abolished by treatment of either with limited amounts of protease. The membrane vesicles are not salt sensitive, suggesting integral membrane proteins mediate their interaction with the lipophilic structures (P. Collas and D. Poccia, 1996a,b).

B. Nuclear Envelope Formation

The time of formation of the male pronuclear envelope varies with species. Neither nuclear envelope formation nor disassembly are necessarily coordinate for maternal and paternal chromatins. In Spisula, the male envelope forms only after the female pronucleus has completed meiosis (see Longo, 1973). In the mussel Mytilus, the male pronuclear envelope forms after decondensation while the female pronucleus is going through meiosis. In the sea urchin, the envelope forms during chromatin decondensation while the female pronucleus retains its envelope. Reports of experimental manipulations of the timing of envelope for-

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mation in vivo have been few. In vitro systems have opened up new ways of studying regulation of nuclear envelope formation, and the most useful of these are derived from egg lysates.

1. Amphibians At least two steps are required for nuclear envelope formation: precursor membrane vesicle binding and fusion. Binding of membrane vesicles to decondensed sperm chromatin does not require cytosol in the Xenopus system (Wilson and Newport, 1988; Boman et al., 1992a) and is insensitive to NEM (Newport and Dunphy, 1992). Vesicle binding is independent of ATP or GTP addition (Newport and Dunphy, 1992) but regulated by phosphorylation (Pfaller et al., 1991; Nigg, 1992; Foisner and Gerace, 1993). Binding is sensitive to protease treatment of chromatin or of vesicles, suggesting protein-mediated binding elements on both (Newport and Dunphy, 1992; Wilson and Newport, 1988). Fusion requires Ca2+ (Sullivan et al., 1993), ATP, and GTP hydrolysis (Boman et al., 1992a). The G-proteins appear to be membrane bound (Newport and Dunphy, 1992). Two nuclear membrane precursor populations have been separated: NEP-A and NEP-B (Vigers and Lohka, 1991). The two fractions look identical by electron microscopy but the endoplasmic reticulum enzyme marker a-glucosidase is 10-fold enriched in NEP-A. NEP-A is also sensitive to N-ethylmaleimide and insensitive to high salt, just the opposite to NEP-B. Binding of both is trypsin sensitive. NEP-B binds initially to chromatin. NEP-A is required for fusion and expansion of the envelope. Two receptors are postulated mediating NEP/chromatin binding and NEP-B/NEP-A binding. NEP-B seems to be involved in nuclear pore assembly which increases as the ratio of B/A increases. It was proposed that NEP-B contributes to the inner nuclear membrane and NEP-A to the outer, which is normally thought to be continuous with the endoplasmic reticulum.

2. Fruit Flies In D . melanogaster embryogenesis, mitosis is a modified closed form. An envelope of nuclear membrane proteins persists throughout mitosis (Strafstrom and Staehlin, 1984; Harel et al., 1989). The nuclear envelope remains intact except at the poles which rupture at prometaphase (Fig. 11). A second layer of envelope forms outside the structure which is completed by metaphase to give a so-called spindle envelope, which remains until the early stages of interphase. Nuclear pores disappear at metaphase but reappear beginning at telophase. At interphase a complete nuclear envelope is present but some of the extra membranes persist for a while (Strafstrom and Staehlin, 1984). Lamins, otefin, and a protein p53 remain in mitosis enclosing the spindle in early embryogenesis, but a pore molecule gp 188 is redistributed into the cytoplasm (Harel el al., 1989).

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Fig. 11 Partial retention of the nuclear envelope in Drosophila embryos. Grazing sections of Drosophila embryos in (a) interphase and (b) metaphase. Envelope not continuous near the spindle poles. Metaphase spindles are separated by infoldings of the plasma membrane (pm) called pseudocleavage furrows. Bar, 5 wm. Taken from Stafstrom and Staehelin (1984) with permission.

In spite of the unusual nuclear envelope formation in vivo and the syncytial nature of the early embryo, male pronuclear envelope formation in vitro, at least on heterologous sperm chromatin, is similar to the other model systems. Extracts from 0 to 5-hr embryos form nuclear membranes with pores and lamina on Xenopus nuclei (Berrios and Avilion, 1990) as judged by phase contrast and electron microscopy by 60-90 min (Fig. 12). Chicken or Xenopus sperm nuclei form a nuclear envelope in an S 14 Drosophilu embryo extract. Although chromatin decondensations require ATP and an ATP-generating system, nuclear envelope formation does not (Ulitzur and Gruenbaum, 1989; Ulitzuer et ul., 1992). Membrane vesicles bind to the surface of the chromatin and the new nuclear envelope contains pores. Both the membrane fraction and an S150 cytosolic fraction are required for envelope formation, suggesting the presence of soluble factors as well as membrane vesicle precursors.

3. Surf Clams In vivo, the male pronuclear envelope forms after completion of meiosis as vesicles assemble in the vicinity of the nucleus and then fuse (Longo and Anderson, 1970). Thus factors promoting formation of the male pronuclear envelope

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Fig. 12 Decondensation of Xenopus sperm chromatin and nuclear formation in Drosophila embryo extracts. Phase contrast images at: (a) 0 (b) 20-30 (c,d) 60-90 min. Bar, 25 pm. Electron micrographs: (e) demembranated sperm nucleus-bar, 0.5 pm; (f) 30-min nucleus with partially dispersed chromatin and forming nuclear envelope-bar, 1 pn; and (g) 60-min male pronucleus with nuclear membrane and pores (arrows)-bar, 1 pm. Taken from Berrios and Avilion (1990)with permission.

are not functional until completion of meiosis. The factors are not nascent proteins since the nuclear envelope forms in eggs blocked in protein synthesis, although male chromatin swelling is delayed (Longo et al., 1991). In vitro, postmeiotic extracts form nuclear envelopes on sperm nuclei (Fig. 13) following vesicle aggregation around the chromatin (Longo et al., 1994). The

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Fig. 13 Surf clam male pronuclear envelope formed in virro. Decondensed male pronucleus of surf clam in 65-min oocyte extract which has formed an intact nuclear envelope. A , basal portion of acrosome. Bar. 0.1 pm. Taken from Longo el u l . (1994) with permission.

vesicles are associated with ribosomes on their outer surfaces and undergo apparent fusion, forming pores. Earlier extracts do not form nuclear envelopes even if the chromatin has swollen. EDTA blocks nuclear envelope formation probably by interfering with Ca2+ metabolism. Additional histones, protamines, or 6DMAP also interfere with envelope formation. 4. Sea Urchins

Normal pronuclear envelope formation depends on the state of maturity of egg cytoplasm. In immature eggs, the chromatin does not disperse and membranous cisternae accumulate near its surface but do not fuse (Longo, 1978). During male chromatin decondensation in mature eggs, new nuclear envelopecontaining nuclear pores form, apparently by fusion of bound vesicles (Longo and Anderson, 1968). Even at this time, the single large toroidal mitochondrion of the sperm midpiece and tail remains attached to the nucleus as the sperm aster

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forms. The sperm aster originates from the region of the centriolar fossa following dissociation of the proximal centriole from the sperm flagellum. As it develops the distal centriole also dissociates from the male pronucleus and the two are found in a centrosomal region from which microtubules radiate excluding various ooplasmic organelles. Astral microtubules are needed for pronuclear migration (Zimmerman and Zimmerman, 1967). Endoplasmic reticulum and annulate lamellae concentrate (Longo and Anderson, 1968). Membranes surround the male pronucleus during its migration toward the female pronucleus (Fig. 14). The sperm mitochondrion and flagellum remain associated with the migrating structure. Upon encountering the much larger female pronucleus, the region of the centriolar fossa with mitochondrion and flagellum moves to one side. Fusion of the nuclear envelopes converts the male pronucleus to a portion of the zygote nucleus. The sperm nuclear envelope can only contribute 15% of the male pronucleus surface area, so most membrane must derive from the egg (Longo, 1976). In vivo the nuclear envelope is believed to arise primarily from preexistent endoplasmic reticulum. The rate of male pronuclear envelope development is inversely proportional to the amount of endoplasmic reticulum in the nuclear vicinity in fertilization of eggs cytoplasmically stratified by centrifugation (Longo, 1976). As in clams, protein synthesis inhibitors are ineffective in preventing male pronuclear envelope formation. In vitro, formation of the nuclear envelope requires a membrane fraction as well as cytosol (Collas and Poccia, 1995a). ATP but not ATP hydrolysis is required for membrane vesicle binding. The fusion step requires GTP hydrolysis and cytosolic factors, some of which are heat and N-ethylmaleimide sensitive (Collas and Poccia, 1995b, 1996b).

C. Role of Lamins

Most nuclei possess a fibrous lattice-like layer immediately subjacent to the inner nuclear membrane which extends from pore to pore. The proteins constituting this layer are the lamins, members of the intermediate filament class of proteins, which usually share a central rod domain largely a-helical. Dimerization is mediated by this domain. Dimers subsequently form higher order structures. Disassembly of the lamina at mitosis is due to phosphorylation in the N-terminal and C-terminal lamin domains. Although usually thought of as forming a relatively uniform layer at the chromatin periphery, lamins may not form a continuous interacting structure or be restricted to the periphery. For example, using three-dimensional light microscopy and electron microscopy with anti-lamin antibodies in early embryos of D . melanogaster, Paddy et al. (1990) report that only a small fraction of chromatin (two or three siteslchromosome equivalent) appears to be close enough to the lamina to interact with it. Goldman et al.

Fig. 14 Accumulation of membranes during pronuclear migration at various times after fertilization in a living sea urchin egg. Membranes stained by microinjection of fluorescent lipophilic dye DiI. Note the relatively symmetric clustering of membranes around the male but not the female pronucleus. Taken from Terdsaki and Jaffe (1991) with permission

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( 1992) report that human lamins microinjected into fibroblasts go quickly to the nuclei, but accumulate in the nucleoplasm, only slowly (5-6 hr) appearing in the peripheral lamina. However, Schmidt et al. (1994) find fluorescently labeled Xenopus lamin A microinjected into 3T3 cells is rapidly transported into the nucleus and incorporated into the lamina by 1 hr with no formation of nucleoplasmic foci. Lamins may be found in the nucleoplasm in G1 cells, perhaps as a store prior to assembly into lamina (Bridger et at., 1993). Alternatively, lamins may be part of a nucleoskeleton extending throughout the nuclear interior (Hozak et al., 1995). These may be inaccessible to antibodies and go undetected unless nuclei are extracted, but extraction itself may cause artifactual redistribution of lamins. Nuclear lamin chemistry and function in germ and somatic cells have been recently reviewed (Hutchison et al., 1994; Cox and Hutchison, 1994; McPherson and Longo, 1993). Somatic vertebrate cells normally express -70 kDa A-type lamins and -66 kDa C-type lamins formed by alternate splicing of the same gene. A-type lamins are usually expressed in differentiated cells. In addition, B-type lamins are constitutively expressed in embryos and adults. Lamins usually contain a sequence in the C-terminal domain (CaaX; a = aliphatic residue) which provides a site for isoprenylation and methylation believed to be involved in associating lamins with the nuclear membrane. At mitosis in somatic cells, A-type lamins become solubilized and B-type lamins remain associated with membrane vesicles, but in embryos much B-type lamin is also soluble (see Hutchison et al., 1994). Several male germ line-specific lamins have been reported, often of lower molecular weight than the somatic. These include L,, in Xenupus spermatids and sperm (Benevente and Krohne, 1985), lamins B and C in mouse spermatocytes of about 52 kDa (Furukawa and Hotta, 1993; Furukawa et al., 1994), a 52 kDa protein expressed during rat male meiosis (Moss e f al., 1993), and a 46-kDa protein in mouse (Furukawa and Hotta, 1993). The 46-kDa B3 lamin is produced from differential splicing and alternate polyadenylation of a B2 lamin (Furukawa and Hotta, 1993). Xenopus expresses multiple lamins. Single lamins have been well characterized in surf clams, fruit flies, and sea urchins (Dessev and Goldman, 1990; Gruenbaum et al., 1988; Holy e f al., 1995). However, it is likely that multiple lamins exist in these organisms as well (Maul er al., 1987; Riemer and Weber, 1994; Collas et al., 1995).

1. Amphibians Male germ line-specific lamins are known in Xenopus. The lamin L,, is specific to spermatids and sperm (Benevente and Krohne, 1985). During meiosis a nuclear lamina can be detected by electron microscopy and B 1 -type lamins can be detected by a broad-reacting anti-lamin antibody (Vester et al., 1993). Xenopus

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lamin L,,, (B3) exists in a soluble form in the oocyte as well as associated with the nuclear envelope (Stick and Hausen, 1985; Krohne and Benevente, 1986). Whether lamins are essential for nuclear envelope formation in frogs is controversial. Immunodepletion of lamin B3 does not prevent assembly of the nuclear envelope in vitro, but envelopes are fragile and nuclei small (Newport et al., 1990; Meier et al., 1991). Dabauvalle et al. (1991) report that depletion of lamin B3 interferes with nuclear envelope formation around chromatin, although nuclear envelopes lacking pores and lamina form around bacteriophage h DNA in extracts depleted of pore proteins (Dabauvalle et al., 1990). Jenkins et al. (1993) used magnetic beads and monoclonal antibodies recognizing three B-type lamins found in embryogenesis (Doring and Stick, 1990). Removal of >96% of the lamins did not prevent nuclear envelope assembly and transport was essentially unimpaired. However, Lourim and Krohne (1993) used several monoclonal antibodies and found a membrane fraction associated with a B2-type lamin and a smaller fraction of membrane associated with B3. They suggest that nuclear envelope precursors contain insoluble lamins that are needed for nuclear envelope formation. Lamins and a nuclear envelope will assemble around condensed Xenupus chromatin inhibited with the topoisomerase I1 inhibitor ICRF- 193 (Takasuga et al., 1995). Remarkably, the nuclear envelope-lamina complex continues to grow while in large part detached from the chromatin, suggesting it is independent. Thus swelling of chromatin is not forcing nuclear envelope enlargement which may depend on laminar growth or import of proteins.

2. Fruit Flies Initially only a single gene was characterized encoding Drosophila lamins, but the protein has several isoforms differing in phosphorylation state (Smith and Fisher, 1989). A 75-kDa soluble form is present in late oocytes and early embryos and serves as the lamina source in the early embryo (Smith and Fisher, 1989). During assembly of the nuclear envelope, the 75-kDa form is converted to 74- and 76-kDa forms. More recently, a second lamin gene for a lamin C was reported (Bossie and Sanders, 1993; Riemer and Weber, 1994). Its structure is related more closely to vertebrate lamin genes than to Dmo or the lamin from the nematode C . elegans. Riemer and Weber (1994) suggest that other invertebrates may have multiple lamins as yet uncharacterized. The protein otefin, localized to the inner nuclear envelope, may mediate membrane-lamin binding (Padan et al., 1990). The putative lamin B receptor is a 53-kDa hydrophilic protein abundant in serine and threonine. The hydrophobic C-terminal region may serve as a membrane anchor. In vitro,Xenopus lamins of the input nuclei disappear soon after addition to the lysate and are replaced by Drosophila lamins (Benios and Avilion, 1990). The

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lamins appear by immunofluorescence in phase 4 of decondensation, when the nuclear envelope forms and the nuclei swell (Ulitzur and Gruenbaum, 1989). If the extract is immunodepleted with anti-Drosophila lamin antibodies, no attachment of membrane vesicles or nuclear envelope formation is observed. The 75-kDa antigen is found both in the soluble and membranous fractions (Ulitzur et al., 1992). Soluble lamins can bind to the chromatin in the absence of membranes, including nuclear membrane precursor vesicles.

3. Surf Clams Spisula germinal vesicles have been reported to contain two lamins (Maul et al., 1987), one reacting with anti-lamin B antibodies (65 kDa) and one with antilamin A/C (67 kDa). The B-type lamin is soluble in the nucleoplasm. The A/C type is clam lamin G, a 67-kDa equivalent to lamin LI,, of Xenopus and located on the nuclear envelope. Spisula extracts phosphorylate the 67-kDa lamin (Dessev et al., 1989). A purified ~ 3 4 kinase-cyclin ~ ~ ~ 2 B complex from extracts of activated clam oocytes phosphorylates lamins and promotes disassembly of the germinal vesicle nuclear envelope (Dessev et al., 1991). It is not known whether this activity disassembles the sperm nuclear envelope. In vitro,condensed sperm nuclei in egg lysates fail to stain with an anti-lamin polyclonal antibody that recognizes the 67-kDa lamin of the female pronuclear envelope (Longo et al., 1994). About half of the nuclei from 15-min premeiotic extracts stain. All 65-min swollen male pronuclei in activated eggs stain. When inhibited with 6DMAP, EDTA, histone, or protamine, nuclei formed in premeiotic extracts show little or no lamin staining or nuclear envelope formation. In 65min extracts, 6DMAP- or EDTA-inhibited nuclei show no lamins, but half of the nuclei blocked with excess histone or protamine do, indicating possible assembly of lamins in the absence of an envelope. 4. Sea Urchins

Only one lamin has been characterized so far in sea urchins. It is a type B lamin of 65 kDa expressed in the embryo (Holy et al., 1995). Sea urchin sperm nuclei were reported to have A- and B-type lamins located only at the tip and base of the conical sperm nucleus (Schatten et al., 1985). More recent work suggests this localization may have been a preparation artifact. When isolated with low concentrations of detergent or no detergent, sperm nuclei exhibit uniform peripheral staining with four different anti-lamin antibodies (Collas et al., 1995). The antigens persist even when membranes have been disrupted or removed by lysolecithin or 0.1% Triton X-100. They are removed along the lateral aspects of the conical sperm nucleus with 1% Triton X-100, but even then persist at the poles in the regions of the acrosomal and centriolar fossae. All nuclear lamin staining is lost when sea urchin sperm nuclei are incubated

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in egg extracts (Collas er al., 1995). Lamins located on lateral aspects of the nucleus being to disappear almost immediately and after 5 rnin all lamin staining is gone, even from the poles (where the membranes of the nuclear envelope remnants are retained). No lamin assembly is detected during subsequent chromatin decondensation or nuclear envelope formation. Thus, as in Xenopus, lamins appear dispensable for initial formation of pronuclei in this system. However, when pronuclear swelling is induced by supplemental addition of ATP, lamins once again appear at the nuclear periphery. These immunofluorescence studies were confirmed by immunoblotting to rule out the possibility of antigen masking and to define the number and sizes of reacting antigens. Five distinct antigens were detected in sperm nuclei (p49, p54, p65, p72, and p84). All but p54 were detected by LS-1 human autoimmune serum which recognizes mammalian and sea urchin lamins AIC and B. A monoclonal anti-intermediate filament antibody (IFA) recognizing a conserved nineamino acid sequence epitope detected p54 and p65. A chicken polyclonal antibody W3-1 made against a fusion protein of the sea urchin B-type lamin recognized only p65. Blots showed the coordinate disappearance of all five antigens following incubation of sperm nuclei in egg extracts, and their coordinate reappearance only in nuclei induced to swell. If sperm nuclei are briefly exposed to egg cytoplasm to remove all sperm lamins, then reisolated and added to fresh extracts, p49 and p54 are not detected when they swell, although the other three antigens are. This indicates that p49 and p54 originate from the sperm and are probably absent from eggs. It also demonstrates that these two antigens are not necessary for pronuclear swelling, although if present they apparently reassociate with swollen nuclei. p54 is only recognized by IFA and thus may be an intermediate filament molecule associated with lamins, since it behaves completely in parallel. A requirement for lamins in nuclear swelling was tested by immunodepletion experiments. Irnmunodepletion of cytoplasmic extracts with anti-lamin antibodies completely prevented swelling of male pronuclei induced by supplemental ATP. N o lamins could be detected by immunofluorescence microscopy or immunoblotting of these nuclei except for small amounts of p54 associated with the unswollen male pronuclei. Both swelling and reacquisition of lamin p65 was demonstrated upon readdition of fresh undepleted cytoplasmic extract. Thus the sea urchin B-type lamin p65 seems to be essential for pronuclear swelling but not initial decondensation.

D. Nuclear Pores Nuclear pores interrupt the nuclear membrane at intervals and are required for bidirectional nuclear transport (for reviews, see Rout and Wente, 1994; Cox and Hutchison, 1994). They disappear at the last stages of spermatogenesis. The

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sperm nucleus loses its poreless nuclear envelope virtually immediately after fertilization, and the male pronuclear envelope forms with pores. Nuclear pores are constructed of eight subunit rings associated with the cytoplasmic face and eight corresponding subunits in a ring on the nucleoplasmic face. Between the two is a ring of spokes enclosing a central granule. The nucleoplasmic ring is connected to the lamina by fibers forming a basket. Several pore proteins have been partially characterized such as gp210 in Drosophila and rat, NSPl and NUPl of yeast, and a family of 0-linked glycoproteins called nucleoponns (Cox and Hutchison, 1994). Three nucleoporin analogues are known in Xenopus (Finlay and Forbes, 1990). In surf clams, pores are observed as soon as the male pronuclear envelope forms (Longo and Anderson, 1970). Pores also appear immediately in the completed male pronuclear envelope in sea urchins (Longo and Anderson, 1968). In Drosophila embryos nuclear pores are lost at mitosis when the perforated nuclear envelope remains. They then reform as mitosis completes (Stafstrom and Staehlin, 1984). Pore formation in vitro has been reported for Xenopus, Drosophila, and surf clams. Pores are present in the male pronuclear envelope of Xenopus within 2 hr (Lohka and Masui, 1983a), in surf clams in 65-min postmeiotic cytoplasms (Longo et al., 1994), and in Drosophila nuclei by 60 min (Berrios and Avilion, 1990). Whether pores are needed for nuclear envelope assembly was investigated by depleting Xenopus extracts of nucleoporins with wheat germ agglutinin treatment of the cytosol (Finlay and Forbes, 1990; Finlay et al., 1991). Envelopes formed. However, Cox (1992) reported that no intact envelopes formed if low speed extracts were treated with WGA. Dabauvalle et al. (1990) report antibodies against nucleoporins in Xenopus extracts prevent pore formation but not nuclear envelope assembly. Lack of pores may result in the lack of lamin import in this system. In Xenopus, immunoprecipitation of nucleoporin p68 is accompanied by coprecipitation of two other proteins in a large complex (Dabauvalle et al., 1990). The cytosolic fraction in Xenopus lysates contains N-acetylglucosaminemodified proteins that appear in the pores and are necessary for transport, though not for assembly of the envelope (Finlay and Forbes, 1990). Alternative models have been proposed for pore formation. Lohka (1988) suggests that pores arise upon fusion of the inner and outer portions of the chromatin-bound flattened membrane vesicles which attach to male pronuclei in vitro. Sheehan et al. (1988) propose a prepore model in which half pores on the inner nuclear membrane bind chromatin and serve as targets for vesicles associated with the remaining pore components. A chromatin-independent assembly mechanism in which pores originate from annulate lamellae abundant in eggs has also been put forth (Dabauvalle et al., 1991). Annulate lamellae are large cytoplasmic membranous cisternae containing pores which are not associated with chromatin and which may represent excess nuclear envelope material or may be precursors to the envelope. DNA or chromatin promote nuclear envelope assem-

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bIy rather than annulate lamellae assembly, suggesting alternate pathways for these materials. Annulate lamellae do not seem to be involved in formation of the sea urchin male pronuclear envelope (Longo, 1976).

E. Summary and Speculations

Initiation of nuclear envelope formation is not generally believed to occur at preferential sites. Membrane vesicles are usually described as binding uniformly to the chromatin surface of male pronuclei prior to fusion. However, in the sea urchin cell-free system, the lack of binding to chromatin from which all membranous material has been extracted and its restoration upon readdition of polar lipophilic structures suggest the possibility that specialized regions of membrane may act as nuclear envelope organizing sites. In this case, a strong correlation exists between the in vivo and in vitro observations of nuclear envelope remnants. The potential generality of such a mechanism needs to be considered at two levels: male pronuclear and somatic chromosomal. Specialized regions of the sperm nuclear envelope, retained in vivo, associated with late decondensing chromatin or resistant to detergent, have been reported for sea urchins, annelids (Colwin and Colwin, 1961), surf clams (Longo et a l . , 1994), and mammals (Yanagimachi and Noda, 1970; Dooher and Bennett, 1973; Ward and Coffey, 1989). Longo and Anderson (1969) suggest that during sea urchin spermatogenesis these specialized regions contain evaginations of nuclear envelope in the fossae and may represent nuclear envelope. Preliminary data indicate detergent resistant lipophilic material is present in the implantation fossae of fish, frog, mouse, rabbit, fox, and bull sperm (P. Collas and D. Poccia, 1996a). Although such structures have not been reported in somatic cells, aspects of nuclear envelope breakdown and reformation in Drosophila (Stafstrom and Staehlin, 1984; Hernandez-Verdun and Gautier, 1994) and membranous spindles in other organisms (Wolf, 1995) raise the possibility of polarized assembly/ disassembly. Why then would specialized envelope not be generally apparent in somatic cells? One possibility is that such regions are more obvious in sperm because they are collected together when chromatin is reorganized during spermatogenesis, but that in somatic cells, the organizing regions are more uniformly distributed in the interphase envelope or with each chromosome during mitosis. Thus a biochemical or immunological search might prove more telling than a morphological one, especially considering the likelihood that membranes are not always well preserved by conventional electron microscopy (Wolf, 1995) and that the structure may consist of both membranous and chromatin binding elements. The formation of nuclear envelopes around individual anaphase chromosomes in the sea urchin embryo (Wolf, 1995) and in somatic cells (Gerace et al., 1978) implies that such organizing centers might be associated with each chro-

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mosome rather than an intrinsic property of the nuclear envelope. Clearly, further molecular characterization of polar lipophilic structures as well as electron microscopy of nascent envelopes during mitosis, pronuclear development, and spermatogenesis are in order. A possible objection to the idea of an organizing center carried by chromosomes is the observation that purified A DNA can be assembled into a nucleus (Newport, 1987). However, this process only occurs after a lag in which assembly and condensation of chromatin take place. It may be that factors from the egg are able to reassemble a structure onto chromatin prior to initiation of the nuclear envelope. It is clear that the nuclear envelope arises from fusion of membrane vesicles it? vitro. Cell-free systems offer the possibility of reconstructing this process with the kind of molecular detail now being achieved in studies of cytoplasmic membrane vesicle traffic such as secretory mechanisms. In these processes, the roles of G-proteins and N-ethylmaleimide-sensitive factors (both operative in nuclear envelope assembly in vitro) are being elucidated (reviewed by Whiteheart and Kubalek, 1995). A variety of intracellular fusion events are mediated by soluble factors for membrane vesicle docking and fusion. Factors similar to the N-ethylmaleimidesensitive fusion protein, NSF, and the soluble NSF attachment proteins in addition to a set of membrane bound receptors that bind these proteins and provide specificity of fusion, would seem likely to operate in nuclear envelope formation as well. The participation of some form of coated vesicle in male pronuclear envelope formation is not yet clear, but G-proteins may have a role in unidirectional membrane vesicle fusion or membrane vesicle uncoating similar to their fusion roles in secretory processes (Bourne, 1988). An ADP ribosylation factor reminiscent of those involved in Golgi vesicle fusion has been identified in Xenopus (Bowman et a!., 1992b). The role of lamins in formation of the nuclear envelope is still controversial. In some systems, lamins seem required (Lourim and Krohne, 1993; Ulitzur e t a / . , 1992). In others, they appear to be involved in growth of the membrane, but not its initial formation (Jenkins et al., 1993; Collas et al., 1995). Hutchison et a / . (1994) outline three models of nuclear envelope assembly: (1) soluble lamins bind to chromatin, nuclear envelope precursor vesicles with associated B-type lamins bind to the chromatin initiating fusion, and then pores are inserted; 2) precursor vesicles bind to chromatin without need of soluble lamins, fusion results in a double membrane, pores assemble, and soluble lamins are imported forming the lamina; or (3) soluble lamins and precursor vesicles with or without attached lamins bind cooperatively to the chromatin and form an envelope after which pores assemble and soluble lamins are imported and the lamina assembles. no sequential early pathway being required. Differences in reported requirements for lamins in nuclear envelope assembly between and even within the model systems may be rationalized in several ways, and precautions have not always been taken (Hutchison et al., 1994). Although

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imrnunodepletion experiments show that membrane vesicles can bind to chromatin in the absence of lamins, and although A-type lamins can bind to chromatin in the absence of vesicles (Burke, 1990), neither experiment shows the order of events when both are present as in an intact cell. Also different antibody preparations might recognize different classes of antigen such as soluble or membrane bound. For example, if one immunodepletes soluble lamins there might still be enough of the membrane-associated form to assemble an envelope or, alternatively, the rate might be slowed to the point where the system deteriorates before assembly is complete. Thus stability of extracts as well as size of precursor pools may affect results of in vitru reconstructions. Furthermore, lamin incorporation evaluated by antibody reactivity is not necessarily the same as lamina assembly. Whether one or more of the models of assembly are correct in all cases is subject to further investigation. The dependence of nuclear swelling on lamina assembly offers at least two possibilities (Newport et al., 1990; Collas and Poccia, 1995). The lamina may drive growth independently of the chromatin as in the experiments of Takasuga and Yagura (1993) and Takasuga et al. (1995), increasing in size whether by thinning of the layer or by accumulating and inserting additional lamins. Alternatively, swelling may occur by import of macromolecules and subsequent increase in water content, import depending on a functional envelope with lamina and pores. In either case the lamins would undergo restructuring of local interactions. The nature of lamin binding both to chromatin and to membranes is currently of great interest. It is unlikely that lamins directly associate with histones, because no lamin binding to histone or DNA-histone beads is observed in vitro (Glass and Gerace, 1990). Several integral membrane proteins which bind to lamins are known to be localized to the inner nuclear membrane (Foisner and Gerace 1993; Chaudhary and Courvalin, 1993). These lamin-associated proteins can also bind to chromosomes before B-type lamin-membrane vesicles do in mitosis. Other lamin receptors may include peripherin (Chaley et al., 1984), proteins of the nuclear scaffold (Fields and Shaper, 1988), and the lamin B receptor (Schuler et al., 1994; Ye and Worman, 1994). It is clear that all receptors and ligands need to be defined for the in vitru systems. This identification will be greatly aided by isolation of mutants in these proteins. Another unsolved problem is how two nuclei in a common cytoplasm may undergo differential envelope assembly/disassembly. In some cases lamina and envelope disassembly occur in parallel (Dessev et al., 1989); in other cases, the nuclear envelope remains when lamina are disassembled (Newport and Spann, 1987). A situation such as when the sea urchin sperm nuclear envelope is disassembled while the female pronuclear envelope remains intact could be explained in this way but seems unlikely since the female pronucleus retains lamins (Schatten et al., 1985). Alternatively, it might result from different lamins, different modifications of the lamins, or different lamin-associated proteins in the two pronuclei.

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The relationship between the lamins and intermediate filaments as potential sources of communication and organization between nucleus and cytoplasm should be considered more carefully as well. Lamins may extend throughout the nuclear interior (Hozak et al., 1995). Intermediate filaments can associate with the nuclear envelope (Cook, 1988) and may serve an integrating function with the endoplasmic reticulum or cytoskeleton. It is worth noting that lamins are assumed to be evolutionary progenitors of intermediate filaments (Klymkowsky, 1995) and cytoplasmic intermediate filaments of invertebrates are closely related to lamins in having central and tail domains (see Reimer and Weber, 1994; Klymkowsky, 1995). Although in vitro, nuclear envelopes are formed from membrane vesicles, the vesicles are probably mostly created in the process of egg lysis. A major future task will be to sort out the origin of the vesicles and to fractionate populations carrying out different steps of the assembly reaction as begun by Vigers and Lohka (1991). Much of the membrane population undoubted derives from the endoplasmic reticulum. The endoplasmic reticulum seems to be the major source of envelope (Longo, 1976; Collas and Poccia, 1996b), although the role of annulate lamellae needs further exploration (Dabauvalle et al., 199 1). Just as endoplasmic reticulum may give rise to nuclear envelope, envelope may give rise to endoplasmic reticulum. For example, the biogenesis of certain forms of endoplasmic reticulum in a CHO cell-derived cell line (UT-1) may initiate at the nuclear envelope (Pathak et al., 1986). Formation of a specialized smooth endoplasmic reticulum, triggered by a fungal metabolite, derives as a set of lamellar stacks from the outer nuclear membrane with which it shares an enzyme marker. These kinds of observations make the data of Terasaki and Jaffe (1991) particularly intriguing. The accumulation of endoplasmic reticulum and/or vesicles about the male pronucleus during migration toward the female pronucleus and their envelopment of the zygote nucleus and persistence into the first division imply a cytoplasmic organizing role for the male pronucleus, the function of which is for now a matter of speculation (Fig. 14). Although the authors describe this organization as coincidental with the sperm aster, which acting through microtubules can certainly organize cytoplasmic domains, the positioning of the membranous “cloud’ is not eccentrically disposed, as the sperm aster is, emanating from the centriolar fossa or off to one side at pronuclear fusion. Continuity between the outer nuclear envelope and portions of the endoplasmic reticulum could account for organization of the cloud and offer additional roles for the male pronuclear envelope.

VI. Male Pronuclear Activities Once the chromosomal proteins can support nucleosome formation, chromatin is sufficiently decondensed, and a new envelope has formed, the male pronucleus is

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capable of activity. Resumption of DNA synthesis must occur within the first cycle. Transcriptional activation may or may not be delayed to later embryonic stages. A. Replication

In vivu, initiation of replication in male pronuclei normally follows formation of the nuclear envelope. In vitro, extensive replication has only been reported in amphibian systems. Limited success was reported with fruit flies and sea urchins. No reports are available for surf clams. 1. Amphibians

In Xenopus, the first DNA synthesis occurs at about 20 min postfertilization and depends on germinal vesicle breakdown as evaluated by microinjection experiments (Gurdon and Woodland, 1968). Activated eggs replicate microinjected nuclei or DNA, but stage VI oocytes do not. Extracts from immature oocytes inhibit replication when mixed with extracts of activated eggs in v i m (Zhao and Benbow, 1994). Nuclei are also smaller than controls. It was proposed that oocyte cytoplasm contains negative regulators of DNA synthesis. In vitro, one round of semiconservative replication of male pronuclei takes place after nuclear envelope formation (Blow and Laskey, 1986). Permeabilization by lysolecithin or passage through mitosis makes them capable of replication, again suggesting that the block to rereplication is attributed to utilization of a “licensing factor” which must be renewed from the cytoplasm each cycle (Blow and Laskey, 1988). Extracts treated with 6DMAP are unable to replicate male pronuclei (Blow, 1993). This was attributed to a lack of a replication (licensing) factor which alters G1 chromatin before the nuclear envelope forms but which cannot cross the nuclear envelope. The factor affects initiation not elongation. The male pronuclei have normal looking nuclear envelopes but their chromatin is not uniformly decondensed. Nuclei which had seen uninhibited cytoplasm for 15 min prior to addition of 6DMAP could replicate. A revised licensing factor model was presented. Multiple replication centers can be detected in male pronuclei which contain newly synthesized DNA labeled with bromodeoxyuridine. These remain constant throughout S-phase (Cook, 1991; Almouzni and Wolffe, 1993). These sites do not seem to require specific replication origin sequences or chromatin structure, and it has been suggested that the requirements for origins in rapidly replicating early nuclei are relaxed.

2. Fruit Flies Replication in Drosophila embryo nuclei is exceeding fast, requiring as little as 4 min. Crevel and Cotterill (1991) report that in vitro, 0 to 2-hr embryo extracts

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support DNA synthesis (bromodeoxyuridine incorporation) in added Xenopus sperm nuclei. In their hands, up to 30% of input nuclei round up and form nuclear envelopes if the embryos were cold treated prior to embryo disruption. Extracts from 0 to 5-hr embryos did not work as well. They estimate 30-50% of the DNA of each nucleus replicated. They suggest mostly repair synthesis occurs without a lag period in untreated cytoplasms and replication occurs after a lag period in treated extracts. The nuclei never enter mitosis, but further incorporation is seen if the envelope is permeabilized.

3. Sea Urchins

In vivo, replication is initiated in the fused male and female pronuclei 30 min postfertilization, but does not require pronuclear fusion (Longo and Plunkett, 1973). It initiates at the same time in all nuclei in polyspermic eggs (Poccia et al., 1978, 1984). In vitro, extracts from fertilized but not from unfertilized sea urchin eggs were reported to support incorporation of deoxyribonucleotide triphosphates into DNA in permeabilized Xenopus but not sea urchin sperm nuclei (Zhang and Ruderman, 1993). Incorporation is sensitive to aphidicolin, suggesting it is semiconservative replication due 10 a or 6 DNA polymerases. The extent of completion of S-phase was not reported. Cell cycle regulation seems to be absent since 3-min GI and 30-min S-phase extracts were similar in supporting incorporation.

B. Reinitiation of Transcription RNA synthesis in Xenopus and Drosophila is activated during cleavage stages. Thus male pronuclei and early embryo nuclei are devoted primarily to replication and mitosis, but not transcription, and the issue is one of transcriptional repression rather than activation. In contrast, sea urchin RNA synthesis is initiated in the first cell cycle (Poccia et a l . , 1985) and surf clam transcription in the one to two cell stage (Firtel and Monroy, 1970). In these organisms, the inactive sperm genome must undergo rapid transcriptional activation.

1. Amphibians Transcriptional activation in Xenopus occurs at the midblastual transition. In fact, somatic nuclei which are engaged in RNA synthesis cease when injected into fertilized eggs (Gurdon and Woodland, 1968). In vitro transcription systems from Xenopus have been developed to study regulation of a variety of genes (see Almouzni and Wolffe, 1993). For example, the role of HMG 17 in preventing transcriptional repression is under investigation with such systems. Many tran-

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scription factors are maternally stored and also amenable to analysis as is the relationship of chromatin structure to activation.

2. Fruit Flies Transcriptional activation occurs in nuclear cycle 10 in Drumphila (Edgar and Schubiger, 1986). Histone gene transcription is activated at cycle 10 and is restricted to S-phase. Repression in Drusophila embryos has been studied in reconstituted chromatin made from extracts. Chromatin assembled onto plasmid DNA from embryo extracts is transcriptionally repressed but correctly spaced, offering the possibility of analyzing factors involved in activation of specific genes in the early embryo (Becker et al., 1994). It is possible to purify the chromatin with magnetic bead technology (Sandaltzopoulos et a l ., 1994). Histone H I seems not to be involved in transcriptional inhibition in these studies. It is not clear if a B4 homolog exists in early Drosuphila nuclei which plays a role in repression. A soluble nuclear fraction extracted at low salt concentrations from Drosuphila embryo nuclei contains basal RNA polymerase I1 transcription factors and supports active transcription of naked DNA or reconstituted chromatin templates (Kamakaka and Kadonage, 1994). The fraction is deficient in H1 and other nonspecific repressors of transcription. It lacks some sequence-specific transcription factors that are extracted in low salt.

3. Sea Urchins Autoradiography experiments suggested that RNA synthesis is activated in the early embryo (Selvig et al., 1972). In polyspermic eggs, incorporation of labeled uridine into RNA commences by 30 min or the time of DNA replication, but is independent of replication (Poccia et al., 1985). Transcripts for the a-class of histones are predominantly expressed. The pattern of transcription does not change for at least 4 hr postfertilization.

C. Summary and Speculations In order to reactivate replication and transcription in male pronuclei, a new nuclear envelope must be formed, chromatin must be decondensed, and somatic histones must be assembled. These transitions are established just before (sea urchins, surf clams) or well in advance (amphibians, fruit flies) of reactivation. Cell-free systems have provided the basis of a useful model concerning the block to rereplication, that of the “licensing factor” required once per cycle and available only at mitosis or when the nuclear envelope is artificially permeabilized in vitro. This attractive model also suggests a possible requirement

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for disassembly of the sperm nuclear envelope soon after fertilization, namely, to permit the dormant nucleus to acquire licensing factor. It is important to establish whether the licensing factor model applies to flies, clams, and urchins as well as amphibians, but none of these systems yet exhibits robust replication in cell-free extracts. Identification of the cytoplasmic factor(s) should allow a direct approach to establishing their role in the other systems. The lack of extensive replication in three of the model systems might be due to deficiencies in the input nuclei or the extract. At least in the case of Drosophila, limited replication cannot be attributed to an intrinsic defect in added Xenopus nuclei which replicate in Xenupus extracts. On the other hand, it is not clear whether histone modeling of frog chromatin is normal in fly extracts. Lack of substantial replication may result from incorrect nuclear envelope formation or degradative activities in some preparations. Reconstituted chromatin made from extracts and defined templates will prove useful in the future for studies of transcriptional regulation. A potential problem is that nucleosomes may not position correctly with respect to promoters and enhancers (see Dimitrov and Wolffe, 1995). A second is that lysates are a rather complex mixture of factors. Nonetheless, such reconstitutes from flies and amphibians will be valuable in forming fully repressed chromatin to be activated. In addition, it may be possible to form sea urchin or clam male pronuclei in vitro which will be active in transcription, although this has not yet been addressed. These nuclear templates could provide a second type of chromatin for studies of transcriptional regulation.

VII. Conclusions Several in vitro systems which more or less faithfully mimic in vivo transitions occurring during formation of male pronuclei from sperm nuclei are now available. Studies using cell-free systems will contribute greatly to our understanding not only of events involved in pronuclear formation, but of those occurring in other nuclei as well, such as nuclear envelope dynamics, gene regulation and replication, and alterations of chromatin structure and histone chemistry. Cell-free systems provide a powerful tool for investigating the relationship between the three-dimensional architecture of chromatin and its composition, since the transitions are difficult to analyze in living cells and impossible to reproduce in highly purified systems. In vitro studies, especially those used in conjunction with genetic and molecular tools, will help to identify major players such as lamin membrane receptors and chromatin receptors in envelope assembly/disassembly reactions. Generation of repressed and activated chromatin templates should provide illuminating comparisons of chromatin structures and protein factors involved in transcriptional control and replication. The comparative approach will confirm the generality of pathways uncovered.

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Longo, F. J. (1980). Reinsemination of fertilized sea urchin (Arbacia punctulata) eggs. Dev. Growth Diyer. 22, 219-227. Longo, F. (198 I ). Regulation of pronuclear development. In “Bioregulators of Reproduction” (G. Jagiello and C. Vogel, Eds.), pp. 529-557. Academic Press, NY. Longo, F. (1985). Pronuclear events during fertilization. In “Biology of Fertilization” (C. B. Metz and A. Monroy, Eds.), Vol. 3, pp, 251-298. Academic Press, NY. Longo, F. J. (1991). Gamete interactions and the fate of sperm organelles in fertilized echinoderm eggs. J. Electron. Microsc. Tech. 17, 246-265. Longo, F. J ., and Anderson, E. (1968). The fine structure of pronuclear development and fusion in the sea urchin, Arbacia punctulata. 3. Cell Biol. 39, 335-368. Longo, F. J . , and Anderson, E. (1969). Sperm differentiation in the sea urchin Arbacia punctulata and Strongylocentrotus purpuratus. 3. Ultrasrr. Res. 21, 486-499. Longo, F. J . , and Anderson, E. (1970). An ultrastructural analysis of fertilization in the surf clam, Spisula solidissima. 11. Development of the male pronucleus and the association of the maternally and paternally derived chromosomes. 3. Ultrastruct. Res. 33, 5 15-527. Longo, F. J., and Kunkle, M. (1978). Transformations of sperm nuclei upon insemination. Curr. Top. Devel. Biol. 12, 149-184. Longo, F. J., and Plunkett, W. (1973). The onset of DNA synthesis and its relation to rnorphogenetic events of the pronuclei in activated eggs of the sea urchin, Arbacia punctufuta. Dev. Biol. 30, 56-67. Longo, F. J., and Schuetz, A. W. (1982). Male pronuclear development in starfish oocytes treated with I-methyladenine. Biol. Bull. 163, 453-464. Longo, F. J., Cook, S . , Mathews, L., and Wright, S . J. (1991). Nascent protein requirement for completion of meiotic maturation and pronuclear development, examination of fertilized and A-23187-activated surf clam (Spisula solidissima) eggs. Dev. Bio. 148, 75-86. Longo, F. J., Mathews, L., and Palazzo, R. E. (1994). Sperm nuclear transformations in cytoplasmic extracts from surf clam (Spisula solidissima) oocytes. Dev. B i d . 162, 245-258. Lourim, D., and Krohne, G. (1993). Membrane-associated lamins in Xenopus egg extracts, Identification of two vesicle populations. 3. Cell Biol. 123, 501-512. Luttmer, S., and Longo, F. (1986). Examination of living and fixed gametes and early embryos stained with supravital fluorochromes (Hoechst 33342 and 3,3’-dihexyloxacarbocyanineiodide). Gamete Res. 15, 267-283. Luttmer, S . J., and Longo, F. J. (1987). Rates of male pronuclear enlargement in sea urchin zygotes. J . Exp. 2001.243, 289-298. Luttmer, S. J., and Longo, F. J. (1988). Sperm nuclear transformations consist of enlargement and condensation coordinate with stages of meiotic maturation in fertilized Spisula solidissima oocytes. Dev. Biol. 128, 86-96. Masui, Y., Lohka, M., and Shibuya, E. (1984). Roles of CaZ+ ions and ooplasmic factors in the resumption of metaphase-arrested meiosis in Rana pipiens oocytes. Symp. SOC. Exp. Biol. 38, 45-66. Maul, G.G.,Schatten, G . , Jimenez, S . A,, and Carrera, A. E. (1987). Detection of nuclear lamin B epitopes in oocyte nuclei from mice, sea urchins, and clams using a human autoimmune serum. Dev. Biol. 121, 368-375. McPherson, S., and Longo, F. J. (1993). Chromatin structure-function alterations during mammalian spermatogenesis. DNA nicking and repair in elongating spermatids. Eur. J. Histochem. 37, 109-128. Meier, J . , Campbell, K.H.S., Ford, C. C., Stick, R., and Hutchison, C. J. (1991). The role of lamin 111 in nuclear assembly and DNA replication in cell-free extracts of Xenopus eggs. J . Cell. Sci. 98,271-279. Mita, K., Takamune, K . , and Katagiri, C. (1991). Genes for sperm-specific basic nuclear proteins in Bufo and Xenopus are expressed at different stages in spermatogenesis. Dev. Growth Di’jer. 33, 491-498.

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Moss, S . B., Burnham, B. L., and Bellve, A. (1993). The differential expression of lamin epitopes during mouse spermatogenesis. M u / . Reprod. Dev. 34, 164-174. Myake-Lye, R., Newport, J. W., and Kirschner. M. W. (1983). Maturation promoting factor induces nuclear envelope breakdown in cycloheximide-arrested embryos of Xenopus laevis. J . CellEiol. 97, 81-91. Nelson, T., Hsieh, T. S . , and Brutlag, D. (1979). Extracts of Drosophila embryos mediate chromatin assembly in vitro. Proc. Nut/. Acud. Sci. USA 76, 5510-5514. Ner, S . S . , and Travers, A. A. (1994). HMG-D, the Drosophila melanogusrer homologue of HMG 1 protein, is associated with early embryonic chromatin in the absence of H I . E M 5 0 J . 13, 1817-1822. Newport, J. W. (1987). Nuclear reconstitution in v i m . Stages of assembly around protein-free DNA. Cell 48, 205-217. Newport, J., and Dunphy, W. (1992). Characterization of the membrane binding and fusion events during nuclear envelope assembly using purified components. J . Cell Biol. 116, 295-306. Newport, J. W., and Spann. T. (1987). Disassembly of the nucleus in mitotic extracts. Membrane vesicularization, lamin disassembly and chromosome condensation are independent processes. Cell 48, 219-230. Newport, J . W., Wilson, K. L., and Dunphy, W. G . (1990). A lamin independent pathway for nuclear envelope assembly. J . Cell Biol. 111, 2247-2259. Nigg, E. A. (1992). Assembly and cell cycle dynamics of the nuclear lamina. Semin. Cell. B i d . 3, 245-253. Ohsumi, K., and Katagiri, C. (199la). Characterization of the ooplasmic factor inducing decondensation of and protamine removal from toad sperm nuclei: Involvement of nucleoplasmin. Dev. Biol. 148, 295-305. Ohsumi, K., and Katagiri, C. (1991b). Occurrence of H1 subtypes specific to pronuclei and cleavage-stage cell nuclei of anuran amphibians. Dev. Biol. 147, 110-120. Ohsumi. K., Katagiri, C . , and Kishimoto, T. (1993). Chromosome condensation in Xenopus mitotic extracts without histone H I . Science 262, 2033-2044. Padan. R . , Nainudel-Epszteyn, S . , Goitein. R., Fainsod, A., and Gruenbaum, Y. (1990). Isolation and characterization of the Drosophila nuclear envelope otefin cDNA. J . B i d . Chem. 265, 7808-7813. Paddy, M. R . , Belmont, A. S . , Saumweber. H., Agard, D. A,, and Sedat, J. W. (1990). Interphase nuclear envelope lamins form a discontinuous network that interacts with only a fraction of the chromatin in the nuclear periphery. Cell 62, 89-106. Pathak, R. K.. Luskey, K. L . , and Anderson, R.G.W. (1986). Biogenesis of the crytalloid endoplasmic reticulum in UT- 1 cells: Evidence that newly formed endoplasmic reticulum emerges from the nuclear envelope. J . Cell B i d . 102, 2158-2168. Pfaller, R., Smythe. C., and Newport. I. W. (1991). Assemblyidisassembly of the nuclear envelope membrane, cell-cycle-dependent binding of nuclear membrane vesicles to chromatin in wirro. Cell 65, 209-217. Philpott, A,, and Leno, G. H. (1992). Nucleoplasmin remodels sperm chromatin in Xenopus egg extracts. Cell 69, 759-767. Philpott, A,, Leno. G. H., and Laskey, R. A. (1991). Sperm decondensation in Xenopus egg cytoplasm is mediated by nucleoplasmin. Cell 65, 569-578. Poccia, D. ( 1986). Remodelling of nucleoproteins during gametogenesis, fertilization, and early development. Int. Rev. Cytol. 105, 1-65. Poccia, D. (1987). Regulation of chromatin condensation and decondensation in sea urchin pronuclei. In “Molecular Regulation of Nuclear Events in Mitosis and Meiosis” (R. A. Schlegel, M. S . Halleck, and P. N. Rao, Eds.). pp. 149-177. Academic Press, NY. Poccia, D. L. (1989). Reactivation and modelling of the sperm nucleus following fertilization. In “The Molecular Biology of Fertilization” (H. Schatten and G. Schatten, Eds.). pp. 115-139. Academic Press, NY.

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3 Paternal Investment and lntracellular Sperm-Egg Interactions during and Following Fert iIi zation in Drosophda Timothy L. Karr Department of Organismal Biology and Anatomy University of Chicago Chicago, Illinois 60637

I. Introduction II. Sperm Structure and Production in Drosophilu 111. Sperm Transfer, Storage, and Utilization IV. Syngamy (Sperm Penetration), Pronuclear Maturation, Migration, and Karyogamy A. Syngamy B. Pronuclear Maturation and Migration C. Karyogamy V. Structural Analysis of a “Sperm-Derived Structure” in the Developing Zygote A. The Sperm Forms a Stereotypical Structure in the Fertilized Egg B. The Early Cleavage Divisions C. Sperm Fate in Later Stages of Embryogenesis V1. Genetics and Molecular Biology of Fertilization and Early Embryonic Development in Drosophila A . Maternal-Effect Mutations B. Paternal-Effect Mutations VII. Cytoplasmic Incompatibility VIII. Speculative Models of Sperm Function in the Fertilized Egg A. Model 1-Nutritive Protein Import (Fig. 7A) B . Model 2-Specific Protein Importation (Fig. 7 8 ) C. Model 3-DiffusioniGradient Production (Fig. 7C) D. Model 4-Structural Role (Fig. 7D) IX. Conclusions and Perspectives References

1. Introduction The predominant mechanism for sexual reproduction among eukaryotic organisms involves fertilization of one specialized cell type, the egg, by another specialized cell type, the sperm. The evolutionary mechanisms that gave rise to sexual reproduction based on two sexes have been studied and debated by scientists for over a century (Parker, 1982). While the seminal evolutionary events that led to the development of a two-sex-based system of reproduction are not known, they ultimately gave rise to anisogamy, i.e., two highly disparate cell types, 89

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sperm and egg (Parker, 1982). These two cell types bear little resemblance-egg cells are usually large and spherically or elliptically shaped and contain large quantities of stored products, while sperm cells are almost invariably elongated, thin cells containing little cytoplasm and are specialized for motility. They apparently share only one common theme: both carry the haploid DNA complement of each parent. Over the past 2 decades, significant strides have been made in understanding some of the molecular mechanisms of fertilization. Particularly impressive has been the discovery of specific receptors in mammals and echinoderms responsible for species recognition and specificity (Wasserman, 1987). Thus, at least in those organisms for which such molecules have been identified, we can hope to eventually begin to understand how these highly differentiated cell types: (1) find each other, ( 2 ) interact and fuse at their surfaces, and (3) ultimately form a diploid zygote capable of realizing the developmental program. A more thorough understanding of fertilization would benefit greatly from study of a variety of animal species. However, fertilization has historically been studied in only a highly restricted set of animals-mainly, chordates and echinoderms. Ironically, insects, which arguably represent the most diverse group of animals, have received very little attention from developmental biologists interested in fertilization. As pointed out by Sander (1983, this bias in the field is, for the most part, a practical one: insects usually fertilize their eggs internally and generally produce smaller numbers of egg and sperm, making laboratory studies difficult, if not impossible. Nonetheless, the potential for studying fertilization in insects is enormous, considering the rich genetic heritage of Drosophilu and the recent advances made in understanding the cellular biology and developmental genetics in this model system. Also, from an economic and health perspective, knowledge of the molecular mechanisms of fertilization in insects could represent a powerful and effective point of attack for the biocontrol of insects. The potential involvement of the sperm and/or sperm-derived products in the egg during and following fertilization was implied from our laboratory’s cytological and biochemical studies of Drosophila (Karr, 1991; Graner et al., 1994). Our interest stems from the general observation by numerous investigators over the years that sperm “gigantism” is a common feature in insects (Counce, 1963; Hildreth and Lucchesi, 1963; Warn et al., 1984; Karr, 1991). For example, D. melanogaster sperm, measuring 1.8 mm, are approximately the same length as the adult males. The recent demonstration that these very large sperm are completely engulfed into the egg, persist intact during and following fertilization, and coil into a stereotypical structure may reflect a previously unappreciated role(s) of the sperm in fertilization (Karr, 1991). While this claim remains to be proven, it would, if true, significantly change our view of the role of the sperm following egg penetration and provide new insights into the evolution of sperm gigantism. An even more controverial idea, that extragenic paternal investments participate in the development of the early embryo, will be discussed later.

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From a cell biological viewpoint, the real importance of these results is the suggestion that intercellular sperm-egg interactions following sperm penetration are central to fertilization, particularly in those insects where sperm gigantism has evolved. Recent work by Schatten and colleagues (Simerly et al., 1993) has also shown that the entire mouse sperm enters the egg and also persists for some time after fertilization. More recent work has shown sperm tail entrance and persistence to be a common feature in a number of related mammals (Schatten, personal communication). The challenge now will be to integrate these new and seemingly general findings into the overall picture of fertilization. Our laboratory is engaged in the biochemical and cellular analysis of some of the proteins associated with fertilization in Drosophila. The approach has been to characterize sperm-associated proteins identified using monoclonal antibodies. These antibodies have identified a large family of proteins, many of which are specific to testes (Graner et al., 1994), related by their antibody reactivity. Monoclonal antibodies have also allowed us to study sperm structure and fate in the egg following fertilization. The extraordinary size of the sperm in D . melanogaster aided in this description and has revealed previously unrecognized aspects of sperm behavior and fate. The evolutionary and developmental consequences of sperm structure in the egg, and the potential importance of spermegg interactions during and following fertilization, will be discussed. In this context, I will also discuss recent advances that have led to a deeper understanding of early development, particularly the isolation of maternal-effect mutations affecting fertilization and/or the very earliest stages immediately following fertilization. We are also currently studying a biological phenomenon related to fertilization and early embryonic development known as cytoplasmic incompatibility (CI; Karr, 1994). The phenomenon is characterized by blockage of the normal process of fertilization in particular crosses of strains within the same insect species (Jost, 1970; Werren et al., 1987; O’Neill and Karr, 1990). CI is closely associated with the presence of a bacterial endosymbiont, Wolbachia pipientis, found in a wide variety of insect species (Breeuwer et al., 1992; O’Neill et al., 1992; Boyle et al., 1993). interestingly, CI only occurs when infected males are mated to uninfected females. Therefore, C1 can be viewed as a unique form of male sterility similar to known patemal-effect lethal mutations in Drosophila. This review relies heavily on previous excellent reviews of Sander (Sander, 1985, 1990), to which the reader is referred for a more comprehensive and general assessment of insect fertilization. This review will focus on new results and information since that time, particularly as they relate to paternal contributions, including extragenic contributions, to fertilization. Accordingly, this review will only briefly describe the fundamentals of insect fertilization, focus on fertilization and early embryonic development in the dipteran D . melanogaster, and, where appropriate, refer to related insect species. Another important purpose of this review is to provide a forum for speculation

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about the significance and purpose of the evolution of sperm gigantism in insects. In this context, four speculative models that may be relevant to this unique and intriguing biological conundrum will be presented.

I I . Sperm Structure and Production in Drosophila Although outwardly different in size and shape from their better-known mammalian, amphibian, and echinoderm counterparts, insect sperm and eggs contain essentially the same components necessary for embryonic development. Insect eggs are usually ellipsoidal and large relative to their body size. For example, the mature Drumphila egg measures approximately 0.5 mm in length (approximately the length of the adult abdomen) and 0.2 mm in width. The egg is invested with the same cellular components as those found in all other eggs, including large stores of mRNA, lipids, and proteins. Insect sperm, in addition to the highly condensed chromatin in the head, also contain a flagellar axoneme composed of a 9 2 structure and an acrosome (or a rudimentary acrosome; Lindsley, 1980; Sander, 1985). However, unlike that found in most other animal groups, insect sperm length can in some cases reach monumental proportions. For example, D . melanugaster males produce sperm that are 1.8 mm in length or about the length of the entire adult fly. This is hardly the record-sperm in excess of 20 mm have been recorded in D . hydei, and the record now stands at at least 50 mm for D . bijiurcu (Pitnick et d., 1995). During spermatogenesis, stem cell divisions occur at the proximal end of paired testes, and fully differentiated sperm appear at the distal end, as shown in Fig. 1 (Gonczy et al., 1992). Due to the highly regular and stereotypical develop-

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Fig. 1 (A) Phase contrast view of an adult testis. The apical end (api) is left in all figures unless otherwise noted: ter, terminal end. Bar = 50 Fm. ( 9 ) Schematic representation of five stages of spermatogenesis. Arrows pointing to part (A) indicate where approximately in the testis each stage begins; cells are displaced in an apical-to-terminal direction as they mature within each stage. Germ line stem cells and somatic cyst progenitor cells are anchored around a hub of somatic cells (hub) at the apical tip of the testis. Only one germ line stem cell (ste) and two cyst progenitor cells (cyp) are represented for clarity. asy. asymetric divisions of a germ line stem cell and two neighboring cyst progenitor cells give rise to one primary gonial cell (spg) and two cyst cells (cyc), respectively. mit, the spermatogonial cell undergoes four mitotic divisions, while the cyst cells no longer divide. gro, the resulting 16 spermatocytes (spe) grow dramatically. mei, the two meiotic divisions occur. mor, the 64 haploid spermatids (spt) undergo dramatic morphological changes. Only 6 elongating spermatids are shown for clarity. Because of the length of the sperm tail, fully elongated spermatids have their nucleus at the terminal end of the testis, while the tail extends almost to the apical end. During this last stage, the two cyst cells become structurally distinct, the head-cyst cell (cyh) being associated with the sperm heads and the tail-cyst cell (cyt) elongating with the growing sperm tails. The head-cyst cell then becomes entrapped by a specialized epithelial cell (tec) located in the terminal part of the testis. Coiling of the sperm bundle ensues, followed by release of motile spermatozoa (spz) into the seminal vesicle. Only one spermatozoon is shown for clarity. See text for additional information. Reprinted, with permission, from Giinczv et al. (1992)

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mental pathway of gametogenesis in Drosophilu and other insects, spermatogenesis (and oogenesis) has been a favorite subject of structural biologists over the years. Gametogenesis has been elegantly described in great detail at the light and electron microscopic levels; the reader is referred to reviews of this subject (Lindsley, 1980; Mahowald and Kambysellis, 1980; Henning and Kremer, 1990). Recent excellent reviews of oogenesis (Spradling, 1993), spermatogenesis (Fuller, 1993), and embryogenesis (Foe e f al., 1993) have appeared in the literature, to which the reader is referred for additional information.

111. Sperm Transfer, Storage, and Utilization Since sperm of D . melunogaster are 1.8 mm in length, it seems unlikely that sperm actively swim through the duct to the female. Instead, as discussed by Sander (1990), some as yet unexplained force propels sperm into the female genital tract. It has been estimated that D. melanoguster females can store, on average, about 700 sperm (Lefevre and Jonsson, 1962; Fowler, 1973; Gilbert, 1981). This is in stark contrast to the number of sperm transferred and stored by other animals and other species of insects. For example, a queen bee can store an estimated 4 to 6 million sperm. Honeybee sperm are, of course, much shorter in length. The extreme variation in sperm numbers is undoubtedly due, at least in part, to the evolution of sperm gigantism in Drosophilu (Pitnick and Markow, 1994b). This extraordinary variation in sperm size and numbers raises many intriguing questions about how and why sperm gigantism evolved and how this is beneficial to those animals where it has occurred. Once sperm transfer is completed, sperm move from the uterus into the sperm storage organs of the female. Following sperm transfer and storage, the female controls the patterns of sperm utilization, as documented in a variety of inspect species (Sander, 1990). The efficiency of sperm utilization in insects is remarkable. For example, the Drosophilu female lays about the same number of fertilized eggs in her lifetime as sperm stored (Gilbert, 198I), indicating that virtually every sperm stored is utilized. Obviously, adaptation of such efficient utilization of gametic resources is a survival strategy employed by many species of insects. The gametic strategies that have evolved in insects are, of course, quite different from the reproductive strategies used by many animal groups utilizing both internal (e.g., mammals) and external (e.g., echinoderms) strategies, where enormous numbers of sperm are produced, but only a minute fraction are utilized (Parker, 1982).

IV. Syngamy (Sperm Penetration), Pronuclear Maturation,

Migration, and Karyogamy At the completion of oogenesis, the egg arrests in meiotic metaphase I (Huettner, 1924; Sonnenblick, 1950; Davring and Sunner, 1973). In response to either

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sperm penetration or egg hydration at the time of ovulation, or both, the egg is activated, protein synthesis begins, and meiosis I1 is completed (Doane, 1960; Mahowald et al., 1983). The time from ovulation to the first mitotic division has been estimated at about 20 min in D . melanogaster (Rabinowitz, 1941). It has not been possible to observe the earliest events of fertilization, particularly sperm entry, as they occur inside the female. Therefore, only those events present after eggs are laid are detectable, and the percentage of eggs in these very early stages of fertilization represents a small percentage of the total (females tend to “hold’ their eggs for some time following fertilization). Although incomplete, some of the basic events during this stage have been documented as described below.

A. Syngamy

Syngamy usually refers to the fusion of sperm and egg membranes that initiates the subsequent events leading to karyogamy. However, very little is known about syngamy in insects. The limited data available on this subject indicate that syngamy in insects occurs by very different mechanisms than those employed by other animals. The elegant electron microscopic study of Perotti (1975) has shown that sperm penetration in D . melanogaster does not involve sperm-egg membrane fusion, in direct contrast to what is known to occur in other animal groups. Thus, at least in Drosophila, other mechanisms for sperm entry have evolved that d o not include sperm-egg membrane fusion, and, technically, syngamy does not occur (at least not at the cell surfaces). The electron microscopic evidence indicates that the sperm enters by puncturing a hole in the egg oolemma (Perotti, 1975). This opening is soon closed, apparently by a “curing” of the membrane (Perotti, 1975). This raises intriguing and important questions about the precise mechanism of sperm entry and the fate of the sperm membrane following sperm entrance.

B. Pronuclear Maturation and Migration Migration of the female pronucleus would appear to rely on an extensive array of microtubles nucleated by the sperm aster (W. Theurkauf, persona1 communication). Some of the events of pronuclear maturation and migration are shown in Fig. 2. The fertilized eggs shown in Fig. 2 were fixed and stained to reveal both the maturing nuclei and sperm tail. Shortly after sperm entry, meiosis 1 and 11 are completed, and, as shown in Fig. 2A, the four haploid products of meiosis are aligned normally to the egg surface in the anterior-dorsal region of the egg. Following sperm entry, maturation of the sperm nucleus and female pronucleus commences (Figs. 2B and 2C). The female pronucleus migrates to the center part of the egg near the anterior end at approximately 75% egg length (the posterior end of the egg is considered to be 0% egg length). The rates at which the two

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pronuclei mature are apparently different, as shown by a comparison of Figs. 28, and 2C. Both the shape and extent of DNA decondensation are different in the two pronuclei. The female pronucleus (top arrow in Fig. 2A) appears more expanded and decondensed than the male pronucleus (bottom arrow in Fig. 2A). The differences in the kinetics of maturation probably reflect the very different nuclear structures involved. The sperm nucleus must first decondense from its very compacted state and then import and/or exchange proteins. Presumably, by the end of this process, both nuclei are identical with respect to their protein compositions, both are invested with nuclear membranes, and both begin DNA synthesis.

C. Karyogamy

In order to form the diploid zygote, the nuclear membranes surrounding the two pronuclei must fuse (this fusion is known as karyogamy). During the entire period of pronuclear decondensation and migration, DNA replication occurs and presumably is completed by, or shortly after, the time the two nuclei complete migration (Shamanski and Om-Weaver, 199 1). Following maturation and migration, the two pronuclei lie next to each other in the interior of the egg at approximately 75% E.L., as shown in Fig. 2D. At this stage, nuclear membranes have formed or are in the process of forming around each individual pronucleus as they each condense following replication (Fig. 2E). The first mitosis ensues (Fig. 2F) resulting in two diploid zygote nuclei. (Lin and Wolfner, 1991; Lopez et d.,1994). The exact nature of the ensuing events of mitosis is only poorly understood. These events have been recorded at the light microscopic level in great detail using conventional sections and stains (Huettner, 1924; Rabinowitz, 1941; Sonnenblick, 1950) and, more recently, through confocal microscopy and indirect immunofluorescence antibody staining (Karr, 1991; Lopez et al., 1994). An excellent review of the current state of our understanding of these crucial early events has recently been published (Foe et al., 1993). Other than these classical descriptive studies, we know very little about the molecules mediating these events. However, as discussed below, new insights are being provided by genetic and biochemical studies of fertilization.

< Fig. 2 Pronuclear maturation, migration, and fusion in Drosophila melunogaster. Young fertilized eggs were fixed and stained with a DNA-specific fluorochrome. (A) Five products of meiosis, three polar bodies (bracket) and two pronuclei (arrows) are observed in the anterior region. ( B and C) High magnification views of female (B) and male (C) pronuclei showing the initial stages of pronuclear decondensation. (D-F) Formation of the zygote nuclei. Fully decondensed and replicated nucici apposed and touching (D); fully condensed nuclei lying immediately next to each other (E): anaphase of the first mitosis (F).

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V. Structural Analysis of a “Sperm-Derived Structure” in the Developing Zygote As previously shown (Karr, 1991), the sperm enters the egg intact and localizes within the anterior region of the egg. This structure has some interesting features that may provide clues to its function in the egg. It is important to keep in mind that, because the sperm persists in the egg throughout early embryogenesis, the sperm structure observed in the egg is more accurately referred to as a spermderived structure. Although we know very little about the biochemical changes occurring in, on, and around the sperm, it is safe to assume that proteins in the sperm are degraded, modified, or bound by specific components in the egg. Presented below are some of the data, accumulated in the laboratory over the past 5 years, that are relevant to the behavior and interaction of the sperm during and following fertilization.

A. The Sperm Forms a Stereotypical Structure in the Fertilized Egg

Figure 3A shows a three-dimensional reconstruction of the sperm tail. The image was produced from confocal optical sections of the sperm using the DROP-I . I antibody (Karr, 1991; Graner et a / . , 1994). A unique feature of this structure is the highly stereotypical folding and coiling of the sperm in the anterior end of the egg. Observation of numerous fertilized eggs confirms the regularity of this structure, suggesting that sperm-egg interactions are necessary for the observed folding and coiling. Further indirect evidence of sperm-egg interactions comes from numerous structural changes observed during and immediately following sperm entrance (Karr, 1991). Presumably, sperm receptors and/or other interacting molecules are present in this region. As discussed further below, some maternal-effect mutations in D . melanogaster disrupt this structure, suggesting that the proteins affected by these mutations interact with the sperm. The length of the sperm tail in the egg was directly measured from three-dimensional reconstructions like the one shown in Fig. 3A. These measurements confirm that the entire sperm enters the egg. Similar results have now been found in 10 other species of Drosophila (Karr and Pitnick, 1996).

Fig. 3 Localization of sperm tail during and following fertilization in Drosophila simulans. Sperm in fertilized eggs were visualized using a mouse polyclonal antisera and a Auorescently labeled goat anti-mouse antibody (A) or using an HRP-based detection system (B,C). Anterior is to the left. (A) The entire sperm tail was computer reconstructed from confocal optical sections (A) and is seen as a thin long string at one end of the egg (the image was contrast-enhanced to accentuate the faint outline of the egg). (B and C) Arrows point to the close association of the end of the sperm tail to one nucleus in the developing zygote at nuclear cycle 4 (B) and nuclear cycle 6 (C). Note that the sperm is always found in the anterior end of the egg and that the sperm tailinucleus association is at or near the anterior boundary of the dividing nuclei.

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B. The Early Cleavage Divisions

An even more striking (and perplexing) aspect of sperm persistence in the developing egg was discovered using polyclonal antibodies that stain the entire length of the sperm tail, including the midpiece. Examination of embryos at various stages postfertilization revealed that the sperm tail remains associated with a single zygotic nucleus (Figs. 3B and 3C). During each nuclear division, the sperm migrates and remains closely associated with the centrosome. Nothing presently is known about how this attachment site is formed or how or why it persists during embryonic development. This structure has no known correlates in other animals, and it remains to be seen if similar behavior can be found in other animal groups. However, one conclusion is inescapable: a paternally derived structure persists in the developing zygote long after fertilization. Some possible roles for the unique sperm-nucleus association are discussed below.

C. Sperm Fate in later Stages of Embryogenesis

The entire sperm structure appears to remain intact throughout much of embryogenesis (Karr, 1991; Graner et al., 1994). During cellularization of the blastoderm, the sperm tail is sequestered in the yolk, excluded from the forming cells (Karr, 199 1). Much later in embryogenesis, the sperm tail fragments and eventually disappears (unpublished observations).

VI. Genetics and Molecular Biology of Fertilization and Early Embryonic Development in Drosophila Over the past few years, the identification of maternal gene products essential for embryonic viability by classical genetic and more recent enhancer-trap methodologies has provided valuable new information about the genetic systems controlling early development (Nusslein-Volhard and Wieschaus, 1980; Driever, 1993; Johnston, 1993). Analysis of mutations and the proteins encoded by them forms the foundation of our understanding of the molecular mechanisms of pattern formation in the embryo. As discussed below, mutations that affect very early events, including the most proximal events following sperm entry, have been identified. A. Maternal-Effect Mutations

1. Young Arrest (fs(1)Ya) Recently, the fs(1)Ya gene product has been shown to be necessary for pronuclear fusion and possibly for the early embryonic mitoses (Lin and Wolfner, 1989). The

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fs(Z)Ya protein accumulates in the sperm nucleus and female pronucleus prior to pronuclear fusion, suggesting a role at this early stage of zygote formation (Lin and Wolfner, 1991). Thefs(l)Ya gene product is localized to the forming nuclear lamina and is speculated to be involved in the signal processes that regulate entry into S-phase (Lopez et al., 1994).

2. Deadhead (dhd) Another maternal-effect gene product thought to act early is deadhead (dhd). Fertilized dhd eggs almost never initiate nuclear divisions (Salz et al., 1994). The predominant phenotypes observed are anaphase-like mitotic figures associated with meiosis I, suggesting that dhd function is involved in the completion of meiosis. In these eggs, the sperm nucleus does not undergo nuclear decondensation (H. K. Salz and T. L. Karr, unpublished communication), suggesting that dhd is involved in some aspect of pronuclear maturation prior to DNA synthesis. The cellular function of dhd is currently unknown. However, the predicted amino acid sequence of dhd has extensive homology with thioredoxin, a multifunctional protein implicated in a variety of cellular processes (Holmgren, 1989), including the regulation of the rate of DNA synthesis (Muller, 1991) and microtubule assembly (Khan and Ludena, 199 1 ). Two intriguing phenotypes observed in the small percentage of dhd embryos that develop are: (1) defects in nuclear migration in the anterior end of the egg and ( 2 ) defects in some of the segmental structures of the head (Salz et al., 1994). Perhaps the two events are related, suggesting that dhd either directly or indirectly acts specifically in the anterior region of the egg.

3. Plutonium (plu) and Pan Gnu (png) The plutonium (plu) and p a n gnu (png) genes are involved in the regulation of DNA synthesis in the fertilized (or activated) egg (Shamanski and Orr-Weaver, 1991). plu and png have nearly identical phenotypes to a previously identified maternal gene giant nudeus (gnu) also thought to regulate DNA synthesis (Freeman and Glover, 1987). In all cases, mutant eggs indiscriminately synthesize DNA without accompanying mitoses, resulting in giant, endoreplicated nuclei. To date, little is known about the proteins encoded by these genes. 4. Maternal Haploid (mh)

The rnh mutation was originally recovered in genetic screens designed to detect maternal-effect mutations (Gans et al., 1975). The mh mutation results in abortive embryonic development, and the large majority of eggs die prior to blastoderm formation; the few eggs that make it past this stage die soon afterward. Sperm enter mhlmh eggs, but the sperm does not form the typical folded and coiled structure seen in wild-type eggs (T. L. Karr, unpublished observations).

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The mh gene has not been cloned, but could be an excellent candidate for an egg product that interacts with the sperm.

B. Paternal-Effect Mutations

The existence of a major structural entity, derived from the father, in the fertilized egg suggests that mutations affecting this structure will influence the course of fertilization. Additional factors brought in by the sperm also represent potential paternal elements that may be involved in development of the zygote. As argued below, the purpose for this structure, if any, may be revealed by the study of sperm-egg interactions. To date, only two mutations have been characterized that affect the paternal genome, paternal loss (pal [Baker, 19751 and ms(3)KBl (Fuyama, 1984, 1986a,b). The low number of paternal-effect genes isolated so far is not surprising, since no systematic genetic search has yet been accomplished. However, large-scale genetic screens are being pursued that are designed to identify genes involved in fertilization (B. Wakimoto, personal communication). With use of the DROP-1 antibody to assess the state of fertilization, paternal genes affecting early development can be screened. To date, no new paternal genes have been identified, but it will be interesting to see the nature and number of mutations isolated by such screens in the future.

1. Paternal Loss (pal) Homozygous pal males produce progeny that, in a small percentage of cases, lack the X, Y, or fourth chromosome (Baker, 1975). Also, high levels of embryonic lethality were observed, presumably due to paternal chromosome loss during embryogenesis. The mutation is a strict paternal-effect, with no known effects on the female. Thus, a gene product, not necessary for sperm development but necessary for chromosome maintenance in the embryo, is defined by this mutation. The eventual molecular cloning should yield interesting new data concerning this gene and its product. 2. ms(3)K81

A strict paternal-effect mutation resulting in almost 100% embryonic lethality was isolated and described by Fuyama ( 1986a,b). Homozygous ms(3)K81 males produce motile sperm fully capable of fertilization (T. L. Karr, unpublished observations). However, shortly after fertilization, a variety of developmental defects are found. These range from defects in chromatin structure to mitotic spindle structure and sperm structure, as shown in Fig. 4. The nature and timing of the defects indicate a role in very early stages of zygote formation and

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Fig. 4 Cytological defects associated with the male-sterile mutation, ms(3)K81. Shown is a high magnification view of a fertilized egg arising from a cross between a wild-type female and a male homozygous for the ms(3)K81 mutation. The sperm tail has been visualized using an antisperm antisera and counterstained with a DNA-fluorescent dye to reveal chromosome structure. The arrowhead shows the location of the end of the sperm tail and the arrows point out various types of chromatin defects. The majority of fenilized eggs show similar aberrant chromatin structures that almost always lead to early embryonic death.

maintenance. We are currently comparing and contrasting the similarities of defects induced by ms(3)K81 mutations to those seen in a related male-sterile phenomenon, cytoplasmic incompatibility (see below). The application of classical genetics and newer molecular genetic techniques to the study of fertilization promises to yield valuable new information about sperm entry, syngamy, and karyogamy. The available data already strongly suggest that paternal products play a central role in fertilization and, perhaps, postfertilization processes in insects. As more attention is drawn to this area, other genes associated with fertilization will certainly be discovered. Given the size of the sperm and the interactions observed between sperm and egg (Karr, 1991; Graner et a l . , 1994), we can expect genetic screens to identify mutations affecting these interactions. In the future we can hope to see this area continue to blossom and Drosophila become an integral member of the family of organisms used as model systems to study fertilization.

VI I. Cytoplasmic lncompatibility Cytoplasmic incompatibility is a phenomenon that disrupts fertilization in particular crosses of strains within the same insect species. CI occurs in at least five orders of insects (Saul, 1961; Ryan and Saul, 1968; Yen and Barr, 1973; Wade

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and Stevens, 1985; Hoffman et al., 1986; O’Neill and Karr, 1990). Interestingly, cytoplasmic incompatibility is intimately associated with an endocellular symbiotic bacterium. Wulbachia pipientis (Saul, 1961; Yen and Barr, 1973; Breeuwer er al., 1992; O’Neill et al., 1992). This association was revealed by use of antibiotics, which remove the bacteria and the incompatibility simultaneously. CI has been discussed as a possible mechanism of speciation (Laven, 1959) and as a tool for the biocontrol of insect pests (Laven, 1967; Karr, 1994). As shown in Fig. 5 , cytoplasmic incompatibility is expressed in an asymmetrical manner-only infected males mated to uninfected females are incompatible. The reciprocal cross of infected females mated to uninfected males yields normal progeny counts, as do crosses between either infected males and females or uninfected males and females. In an incompatible cross, the sperm enters the egg, and karyogamy and zygote formation does not occur (Ryan and Saul, 1968; Jost, 1970; Breeuwer and Werren, 1990; O’Neill and Karr, 1990). The physical entry of the sperm into the egg cytoplasm is sufficient to trigger the early develr Uninfectedegg

,

II 1I Fertilized by spenn from infected Wale

II ‘

’

Fig. 5 Cytoplasmic incompatibility. An uninfected egg fertilized by sperm from an infected male fails to complete karyogamy and/or the first few cleavage divisions. By contrast, an egg from an infected female fertilized by sperm from an infected male completes karyogamy and develops normally. Reprinted with permission.

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opmental stages of cell division, but then most haploid embryos die at an early stage. lncompatible crosses therefore result in very few, if any, viable adults. An incompatible cross is formally equivalent to the paternal-effect mutation ms(3)K81 in D . melanogaster discussed previously. It is also important to note that mature sperm are devoid of detectable Wolbachia, implying that the bacterium exerted its effect during spermatogenesis and that the effect was transmitted in or on the sperm (again, formally analogous to a genetic lesion). The molecular mechanism(s) of incompatibility is (are) currently unknown, but the phenomenon raises intriguing questions about the role of the sperm in fertilization and early embryonic development. Cytoplasmic incompatibility also raises important questions about when, how, and why this form of symbiosis arose initially. Most important for the present discussion, cytoplasmic incompatibility is an extragenic, paternally transmitted form of sterility that has profound effects on the reproductive success of its host. In this respect, cytoplasmic incompatibility is consistent with the idea that a sperm-derived or sperm-delivered product(s) can provide important factors to the egg during and/or following fertilization in insects. Cytoplasmic incompatibility also has important practical implications for strategies of biological insect pest management (Karr, 1994). The rationale for using C1 comes from the unique male-sterile effect described in Fig. 5-only infected males mated to uninfected females result in inviable embryos. Therefore, release of infected males into an indigenous uninfected population should rapidly reduce the number of offspring in the next generation. Of course, care must be taken not to release infected females, which would result in the spread of infected progeny, rendering CI ineffective. A number of laboratory experiments with the tropical warehouse moth Ephestia cuutella, an agricultural pest of stored grain, have shown that cytoplasmic incompatibility can be successfully used as a means of control of this insect (Brower, 1979, 1980). Our laboratory has recently become interested in the cell biology of this intriguing phenomenon. We are using as a model system Drosophila simulans, a sibling species of D . melanogaster. Cytoplasmic incompatibility in D . simulans was first discovered in crosses between strains of D . simulans from southern and northern regions of California (Hoffmann et al., 1986). By applying immunocytochemical techniques originally developed for observation of cellular substructure in the D. melanogaster embryo (Foe and Alberts, 1983; Mitchison and Sedat, 1983; Warn et al., 1984; Karr and Alberts, 1986; Karr, 1991), we are examining the cellular defects associated with cytoplasmic incompatibility. As shown in Fig. 6, our preliminary results suggest that cytoplasmic incompatibility disrupts the normal behavior of chromosomes during the mitotic cycle. One rarely observes normal chromatin figures-instead, only fragmented and aberrant chromosomes are observed. This has lead us to speculate that cytoplasmic incompatibility disrupts the normal process of protein incorporation into the sperm head during maturation (Lassy and Karr, 1996). The

fig. 6 Early embryological defects associated with cytoplasmic incompatibility. Eggs fertilized by sperm from (a) tetracycline-treated parents (normal development) or (b and c) untreated males. Shown in b is an aberrant cytoplasmic bridge formed during the first cleavage division (gonomeric division). Parts of two embryos in c show (left) highly fragmented DNA and (right) two condensed DNA bodies that are probably the result of failed karyogamy. Reprinted with permission from O’Neill and Karr (1990).

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disruption of the normal system of condensation and decondensation related to the presence of a prokaryotic organism could arise by two (not necessarily mutually exclusive) mechanisms: ( 1 ) incorporation of a bacterial protein or proteins into the sperm during spermatogenesis, and (2) modification of specific sperm proteins. Modification of sperm proteins has been recently observed by high-resolution, two-dimensional gel electrophoresis, suggesting candidate proteins for further study (W. Chang and T. L. Karr, unpublished observations). In addition to providing molecular clues to the mechanism of cytoplasmic incompatibility, the eventual elucidation of the exact molecules involved in the expression of cytoplasmic incompatibility promises to yield new information about general mechanisms of fertilization in insects.

VIII. Speculative Models of Sperm Function in the Fertilized Egg For the purposes of illustration, and to provide a forum for speculation and discussion, four possible models suggesting roles of the sperm and/or spermderived structure in early development are shown in Fig. 7. As in all disclaimers that appear with speculative models, each model is not necessarily exclusive, and the “real” model could be an amalgation of some, all, or none of the models presented. With this proviso in mind, each will be discussed separately.

A. Model 1-Nutritive

Protein Import (Fig. 7A)

One obvious consequence of the entrance into the egg of sperm of such extraordinary length is the importation of a fairly significant amount of paternally derived proteins. For example, it has been estimated that the total tubulin delivered to the egg in the sperm is at least 0.1-0.5% of the total tubulin in the egg (Karr, 1991). Other proteins, unique to the sperm, would introduce a new set of proteins into the egg, and as such would represent an infinite change in protein concentration from the egg’s perspective. It is logical to assume that molecular mechanisms in the egg have evolved to utilize, alter, or degrade these paternal contributions. For example, the disposition, fate, and possible function (if any) of sperm proteins in the egg might be highly regulated and essential elements of early development. In support of this idea, using a library of monoclonal antibodies, we have observed differential patterns of antigen loss from the sperm following sperm entry (T. L. Karr, unpublished observations). The eventual identification of these proteins and their ultimate fate may provide important clues about their function in the egg. The evolution of sperm gigantism, in the context of male provisioning and reproductive mating strategies in dipterans, has been examined extensively by Markow and colleagues (Markow and Ankney, 1984, 1988; Pitnick e t a l . , 1991;

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1

1

1

1 D

Fig. 7 Four speculative models of postfertilization sperm function in Drosophila. A-D show possible roles for, and consequences of, sperm persistence during early embryonic development. The first two can be broadly classified as provisioning models: (A) general and (B) specific provisioning. The last two can be broadly categorized as structural models: (C)sperm structure participates in gradient formation and (D) sperm structure interacts with anterior migrating nuclei. Also note that inherent in all four scenarios is a possible fifth functional aspect-marking of an anterior boundary of the early cleavage nuclei by the sperminucleus structure. See text for further details.

Pitnick and Markow, 1994a,b). In certain Drosophilidae, large amounts of accessory gland proteins are taken up through the female reproductive tract and utilized by females in somatic tissue maintenance and oogenesis. However, because of multiple female matings, males run the risk of investing in progeny they do not actually sire (Markow, 1988). One mechanism by which males may provision eggs but remain assured of their paternity is to provision directly through their

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gametes in the form of long, protein-rich sperm (Pitnick and Markow, 1994b). Collaborative experiments, designed to follow the fate of sperm proteins in the developing egg using a library of monoclonal antibodies against sperm, are planned. Proteins with interesting or novel patterns of utilization or localization will be candidates for further study. Once identified, mutations of genes encoding such proteins may provide insights into their functional significance vis a vis provisioning.

B. Model 2-Specific

Protein Importation (Fig. 7B)

Sperm may deliver to the egg specific molecules essential for either fertilization or early zygote viability. Although many possibilities exist, three examples are mentioned: (i) proteins involved in the generation of a functional centrosome (i.e., the sperm basal body), (ii) enzymes necessary for the initiation or maintenance of DNA synthesis, or (iii) proteins that either regulate or directly participate in cell cycle regulation (i.e., cyclins, protein kinases). These could include as yet unknown proteins in addition to the known regulators of the cell cycle, that are unique to the initiation of the first zygote division. Although Fig. 7B shows a hypothetical factor surrounding the early cleavage nuclei, this factor (or factors) may work at any stage during postfertilization development of the egg. Further studies of the specific fate(s) of sperm proteins in the developing egg may identify candidate proteins.

C. Model 3-Diffusion/Gradient

Production (Fig. 7C)

Diffusion of a soluble factor has long fascinated biologists as a mechanism for the generation of a gradient of morphological information during development (Wolpert, 197 1). Over the years, models invoking diffusion-induced gradients have become increasingly sophisticated. However, with one prominent exception in insects, discussed below, very little substantive data on the molecular mechanisms involved have been forthcoming. Nonetheless, diffusion models continue to dominate the theoretical landscape of developmental biology, and, in lieu of equally intellectually attractive and intuitive alternatives, they will continue to do so. The current leading candidate for such a molecule is the morphogen bicoid (Driever and Nusslein-Volhard, 1988a,b; Driever, 1991). Bicoid RNA is localized to the anterior tip of the egg, where it is transcribed. The bicoid protein produced at the anterior end then diffuses throughout the anterior half of the egg where it later becomes incorporated into embryonic nuclei at the time of blastoderm formation (Driever and Nusslein-Volhard, 1988a,b). In the nucleus, the bicoid protein acts as a regulatory molecule directing the transcription of other

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regulatory genes (Driever and Nusslein-Volhard, 1989; Driever, 1993). Therefore, the available data strongly suggest that the bicoid protein gradient in the egg is generated by simple diffusion and that this gradient is intimately involved in directing the overall segmented body plan of the embryo. The sperm tail in the D . melanagaster developing egg forms a “natural” gradient by virtue of its structure in the anterior end (Fig. 3). In terms of sperm volume occupied per unit volume of egg, inspection of the distribution of the sperm in the anterior end clearly indicates a “gradient” of sperm tail. It will be interesting to see if experimental manipulation of the position of the sperm in the egg (e.g., via magnetic microbeads attached to the sperm by anti-sperm antibodies) may affect some aspects of early development.

D. Model 4-Structural

Role (Fig. 7D)

There are two general ways that the sperm tail could provide an essential structural element to the developing egg. The first is more general and relates to the concentration of the sperm tail in the anterior end of the egg. Its mere existence suggests that the egg cytoplasm is organized differently in the anterior region of the egg. This is consistent with the biochemical differences that exist in the egg (see discussion of the bicoid protein above). For example, the sperm tail could bind and organize specific egg proteins in the anterior region of the egg either during or shortly following sperm penetration. This binding could in principle organize other components in the anterior end. In doing so, this reorganization would result in a gradient of proteins similar in shape to that of the sperm tail, as alluded to in Model 3 (Fig. 7C). Another structural role may involve the regulation of nuclear migration into the anterior region of the egg. The pattern of nuclear movement to the egg periphery on first inspection appears synchronous and uniform (Zalokar and Erk, 1976; Foe and Alberts, 1983; Karr and Alberts, 1986). However, upon closer inspection, nuclei are slightly delayed in arriving in the anterior end of the egg, as depicted in Fig. 7D (T. L. Karr, unpublished communication). Because nuclei must pass by, over, and around the sperm tail in route to the egg surface, they may be expected to interact either physically and/or biochemically. Preliminary data suggest that microtubules associated with migrating nuclei and the sperm tail interact. Double-label immunofluorescence using anti-sperm and anti-tubulin antibodies demonstrates that microtubules and the sperm tail come into extremely close contact (physically touching at the resolution of the light microscope), suggesting that this may be the mechanism of the delayed nuclear movements. Therefore, the sperm may serve to physically impede the free movement of nuclei into this region. Another consequence of these interactions would be the incorporation of either sperm proteins or egg proteins bound to the sperm into the advancing nuclei or

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into the domains of organized cytoplasm that surround them (Foe and Alberts, 1983; Karr and Alberts, 1986).

IX. Conclusions and Perspectives Essentially, nothing is known about the evolution of sperm length variation. However, several recent comparative studies examining the adaptive significance of sperm length in a variety of taxa (Gomendio and Roldan, 1991; Roldan et al., 1992; Briskie and Montgomerie, 1992, 1993; Gomendio and Roldan, 1993) including Drosophila (Pitnick and Markow, 1994b; Karr and Pitnick, 1996), have led to the following conclusion. Relatively long sperm either provide an advantage in sperm competition, which is postcopulatory male-male competition for fertilization of the eggs of a specific female during a single fertile period (Parker, 1970), and/or they represent the provisioning of sperm as a form of paternal investment (Sivinski, 1984; Pitnick and Markow, 1994b). At present, no hypotheses consistent with sperm competition theory to explain sperm length evolution in Drosophila have been supported (Pitnick and Markow, 1994b). Although the idea that sperm provide a functional role following sperm entrance into the egg, particularly a role after formation of the zygote, is controversial, if true it has intriguing and important implications for not only our understanding of fertilization and development in insects, but for theories of the evolution of sperm gigantism. Taken together, the following facts support the notion of a functional role for the sperm in the egg in Drosophila: (1) the sperm is always found asymmetrically localized to the anterior region of the egg; (2) it persists intact during embryonic development; (3) the sperm remains associated with a single zygotic nucleus; and (4) the position of this unique sperm tail/nucleus structure within the egg marks the anterior-most boundary of the dividing nuclei during the early cleavage stages. A complete understanding of fertilization in Drosophila awaits the explanation and integration of these observations. Preliminary data suggest that gigantic sperm other than those of D . melanogaster fully enter the egg. For example, in collaboration with Scott Pitnick and Therese Markow at Arizona State University, we have visualized the 17 min long sperm of D.pachea in a fertilized egg (Karr and Pitnick, 1996). We are currently using confocal microscopy and indirect immunofluorescence to create three-dimensional reconstructions of this enormous sperm. The entrance and persistence of a sperm 10 times the length of the D . melanogaster sperm in an egg that is essentially the same size (both eggs are approximately 0.5 mm in length) strongly suggest that specific mechanisms coevolved in the egg to accomodate such gigantic sperm. Given the considerable investment in energy and resources needed to construct a sperm of this length (Pitnick and Markow, 1994a), it is unlikely that this occurred without some sort of adaptive and func-

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tional significance. The models provided (Fig. 7) point out some of the possible functions for this structure and will hopefully spark renewed interest in fertilization in insects in general and in this paternal structure in particular. Our understanding of factors responsible for the maintenance of anisogamy suggests that sperm gigantism should not exist (Parker, 1982), and conventional wisdom based on sexual selection theory fails to explain the occurrence of sperm gigantism. Moreover, no theory of fertilization would predict the presence or persistence of such a structure in the fertilized egg. Assuming an adaptive nature of design, discerning the functional significance of sperm gigantism will change our concept of direct paternal investment during reproduction in insects. The challenge to biologists is to provide an explanation for the design and functional significance of such adaptations,

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4 Ion Channels: Key Elements in Gamete Signaling Alberto Darszon, Arturo Lievano, and Carmen Beltrdn Departamento de GenCtica y Fisiologia Molecular Instituto de Biotecnologia, Universidad Nacional Autdnoma de MCxico Cuernavaca, Morelos 6227 1 , MCxico

I. Why Are Ion Channels Important in Fertilization? 11. Gamete Generalities A . Spermatozoa

B. The Egg 111. Influence of the Ionic Environment on Spermatozoa A . Sea Urchin and Fish Spermatozoa

B. Mammalian Spermatozoa IV. Long-Range Communication between Gametes A. Sea urchins B. Mammals V. Short-Range Communication between Gametes: The Acrosome Reaction A. Sea Urchin Sperm Acrosome Reaction B. The Starfish Acrosome Reaction C. The Mammalian Sperm Acrosome Reaction VI . Do Ion Channels Turn the Egg On? VII. Concluding Remarks References

1. Why Are Ion Channels Important in Fertilization? During the past 10 years it has become apparent that ion channels are not only fundamental to excitable cells, but play a key role in cell signaling in general (Tsien and Tsien, 1990; Hille, 1992; Brown, 1993). Because of this, interest in them has grown explosively, Fertilization, a crucial event in the generation of a new individual, requires communication between sperm and egg. The male and female gametes must be fully mature and competent for fertilization. The success of fertilization depends on gamete information processing from the environment. There are long- and short-range signals emitted by the egg that influence sperm function and lead to proper gamete interaction and finally to fertilization. Although the factors that mediate the sperm-egg dialogue have been studied for close to a century (Lillie, 1919), the detailed molecular mechanisms involved in these events remain elusive. However, there is growing evidence that ion channels are deeply involved in gamete signaling. For instance, in echinoderm, fish, Currrnr Topics in Dsvelopmrnrol Bk,/o,qv. Vol. 3 4 Copyrighl D 1996 by Academic Press. Inc. All right? or reproduction in Any form reserved

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and mammalian spermatozoa, the acrosome reaction (AR),’ a necessary process for fertilization in many species, is inhibited by ion channel blockers (reviewed in Ward and Kopf, 1993; Darszon et al., 1994). On the other hand, egg activation can be stopped by compounds known to interfere with the ion channels responsible for Ca*+ release from intracellular stores (reviewed in Shen, 1992; Swann et al., 1994). This review will focus on the participation of ion channels in the information exchange between gametes themselves and with the environment. Since the best known sperm ion transport systems have been described in the sea urchin, a marine invertebrate (Schackmann, 1989; Darszon et al., 1994), and in mouse, bull, and pig in mammals (Florman and Babcock, 1991; Ward and Kopf, 1993), these species will be referred to more extensively. Without doubt, because of the authors’ partial view, many important contributions will not be mentioned; they can be found in several excellent reviews on general aspects of gamete interaction and function (Trimmer and Vacquier 1986; Eddy, 1988; Yanagimachi, 1988; Garbers, 1989; Saling, 1989; Schackmann, 1989; Nuccitelli et a / . , 1989; Kopf and Gerton, 1990; Jaffe, 1990; Florman and Babcock, 1991; Nuccitelli, 1991; Shen, 1992; Foltz and Lennarz, 1993; Ward and Kopf, 1993; Whitaker and Swann, 1993; Myles, 1993; Miyazaki et al., 1993; Swann et al., 1994).

11. Gamete Generalities A. Spermatozoa

The general design of many animal spermatozoa is quite similar. These small cells are mainly constituted by a head, containing condensed packages of chromosomes in the nucleus, which occupies a significant part of its volume, and the acrosome in many species, a membranous structure sitting as a cap over the nucleus in the anterior part of the sperm head. The tail, which varies in length among different species, is universally composed of a characteristic “9 + 2” complex of microtubules found in eukaryotic flagella and cilia. A few mitochondria at the base of the tail are the power source for movement. The total amount of cytoplasm in sperm is very small. Spermatozoa are very specialized ‘Abbreviations used: AR, acrosome reaction; BCECF, 2’,7‘-bis(2-Carboxyethyl)-5(6)-carboxy fluorescein; [Ca2+],, intracellular calcium concentration; cADPr, cyclic-ADP ribose; CICR, calciumacid; E,, Nemst poinduced calcium release; DIDS, 4,4’-diisothiocyanatostiIbene-2,2’disulfonic tassium equilibrium potential; EM,membrane potential; FSP, fucose sulfate polymer; GDPPS, guanosine 5’-0-(2-thiadiphosphate); GTPyS. guanosine 5’-0-(3-thiotriphosphate); IBMX, 3-isobuthyl-I-methyl xantine; IICR. IPJnduced calcium release; InsP,, inositol I ,4.5-trisphosphate; InsP,R, inosytol trisphosphate receptor; [K+],, external potassium concentration; pH,, intracellular pH; pS, pic0 Siemens; PTX, pertussis toxin: RyR, ryanodyne receptor; TEA+, tetraethylammonium ion.

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cells committed to find, fuse, and deliver their genetic information to the egg. They lack the machinery for protein or nucleic acid synthesis (which is lost during spermatogenesis) and contain only few mRNAs (Kumar et al., 1993). In the sea urchin spermatozoa, the head is conical, as in many species, with a length of -4 pkl and a diameter of 1 pM. It contains a single mitochondrion, the acrosomal granule, the haploid nucleus, a pair of centrioles, G-actin in the nuclear fossa, and a flagellum (0.1 pkf in diameter and -50 pM long). After full development and differentiation in the seminiferous tubules of the testis, mammalian spermatozoa also end up being very long cells composed of heads, midpieces, and tails (Fawcett, 1975; Eddy, 1988). Mitochondria are spirally arranged in the midpiece and contain the distal centriole. The proximal centriole, which serves as the male pronuclear centrosome after sperm-egg fusion, is found at the most posterior end of the head. The sperm nucleus, with its compacted chromatin and the acrosome, occupies the rest of the head space. Mammalian spermatozoa must undergo changes after leaving the testis to become competent for fertilization (reviewed in Florman and Babcock, 1991). These changes occur in the male reproductive tract (epididymal maturation) and in the female reproductive tract (capacitation and AR). During maturation, the sperm surface is modified, there are surface charge changes, the reactivity and distribution of plasma membrane components is altered, and epididymal-secreted proteins are added. It is not known how spermatozoa develop motility as they traverse the epididymes (Bleil, 1991). Mature cauda epididymal or vas defferens spermatozoa cannot bind to or fertilize eggs. They must be incubated in particular media or spend some time in the female genital tract to become capable of binding to the zona pellucida of eggs. This time period necessary for sperm to acquire their fertilizing capacity has been termed “capacitation,” and is not well understood (Yanagimachi, 1988). During this process, motility changes occur, and there are further protein and lipid membrane component rearrangements and modifications (Morton and Albagli, 1973; Berger and Clegg, 1983; Stein and Fraser, 1984; Stein et al., 1986). It is thought that epididymal surface components added to the sperm surface throughout maturation are removed during capacitation, leaving the cell ready to undergo the AR (Yanagimachi, 1988; Rochwerger and Cuasnicu, 1992). Efforts have been made to redefine the various sperm states in ways that could allow their better understanding (Florman and Babcock, 1991). Ion transport systems and, specifically, ion channels probably participate in these events, but their roles remain unestablished.

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B. The Egg In general, eggs are fairly large cells; however, their structure varies greatly from one species to another. The eggs of many invertebrates and vertebrates are filled

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with lipid and protein nutrients which provide the energy and building blocks for embryonic development. Although dormant until fertilization, they contain a repertoire of maternal mRNA and ribosomes ready to initiate development. Sea urchin eggs are, as in other species, large cells (-80 pA4 diameter). They are surrounded by a thick 10 to 30-nm extracellular matrix, named the “vitelline coat,” which is bound to the plasma membrane (Kidd, 1978; Chandler and Heuser, 1980, 1981). A second extracellular matrix, the egg jelly, of -40 pI4 thickness, covers the vitelline layer. This outermost layer triggers the sperm AR (Dan, 1967) and dramatic permeability changes in sea urchin spermatozoa that will later be discussed at length. Sea urchin eggs, in contrast to many other animal eggs, have completed meiosis when released from the ovary. A complete nuclear envelope surrounds the haploid interphase nucleus of decondensed chromatin. Eggs at this stage are metabolically arrested, mainly due to their low intracellular pH (pH,) (Epel, 1978; Shen, 1983). Sea urchin egg jelly can be easily solubilized at low pH. Early fractionation of nondissociated solubilized egg jelly, using ion exchange and gel filtration chromatography, yielded a fucose sulfate polymer (FSP) and a sialoglycoprotein. The FSP fraction displayed all the AR-inducing activity (SeGall and Lennarz, 1979, 1981). However, the activity of egg jelly is reduced when treated with proteases, indicating that a protein component of FSP is important (Ishihara and Dan, 1970; Garbers et al., 1983; Yamaguchi et al., 1989; Shimizu et al., 1990). These results, and new ones (Suzuki, 1990; Mirakami-Takei et al., 1991; Keller and Vacquier, 1994), have led to the notion that the AR-inducing activity in the egg jelly may reside in a glycoprotein and that egg jelly is formed from globular glycoproteins bound to a fibrous fucan superstructure (Bonnell et al., 1994). In starfish eggs, two components have been found to induce the AR: a high molecular weight glycoconjugate (ARIS) and a steroid-based saponin (Co-Aris [Hoshi et al., 1990, 19911). The mouse oocyte grows in the ovary during 16 days from a diameter of 10 to 80 pA4. When halted at the first meiotic prophase (germinal vesicle stage), oocytes grow and then follow through meiosis until they are arrested at the second metaphase. For most mammals, ovulation and the fusion of sperm and egg occur during this second metaphase arrest (Wassarman, 1988a, b). The cumulus matrix and the zona pellucida, a porous, extracellular glycoprotein envelope -5 pA4 thick, surround the unfertilized egg. Before fertilizing the egg, spermatozoa must transverse these surroundings. The cumulus, the outermost layer, apparently selects potentially fertile sperm. Uncapacitated and acrosome-reacted sperm are excluded from the cumulus, while capacitated sperm can enter it (Cherr et al., 1986; Cummins and Yanagimachi, 1986). The zona pellucida plays a crucial role during mammalian fertilization: it mediates sperm-egg recognition, sperm entrance into the egg, and the slow block to polyspermy (Bleil, 1991). This extracellular coat can be easily purified and has been intensively charac-

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terized structurally and functionally (Bleil and Wassarman, 1980a, b, c, 1983; Bleil et al., 1981, 1988; Florman et al., 1984; Florman and Wassarman, 1985; Wassarman 1987, 1988a, b). Three glycoproteins containing N- and 0-linked oligosaccharide chains, ZP1, ZP2, and ZP3, constitute the mouse zona pellucida. Their apparent molecular weights determined by SDS-polyacrylamide gel electrophoresis are 200, 120, and 83 kDa, respectively (Bleil and Wassarman, 1980a, b, c). These proteins are synthesized and secreted by the oocyte during its growth. Dimers of ZP2 and ZP3 form filaments that appear to be crosslinked by ZP1, which exists as a disulfide-linked homodimer in the zona pellucida.

111. Influence of the Ionic Environment o n Spermatozoa Before gametes communicate, they exchange information with their environment. Marine, fish, and mammalian spermatozoa are exposed to important alterations in their ionic milieu as they progress in their journey toward the egg.

A. Sea Urchin and Fish Spermatozoa

Sea urchin spermatozoa are immotile in the male gonads due to the high CO, tension in semen, which keeps pH, at -7.2 (Johnson et al., 1983). Below pH 7.3, dynein, the ATPase that drives the flagella, is inactive, and motility and respiration are repressed (Schackmann et al., 1981; Christen et al., 1982; Lee et al., 1983). Upon spawning, sperm dilution in sea water lowers the concentration of CO,, H+ are released, pH, increases to -7.4, and motility is initiated (Nishioka and Cross, 1978; Christen et al., 1982, 1983a, b, c; Johnson et al., 1983). Dynein can hydrolyze ATP at this pH,, producing ADP that activates mitochondria1 respiration 50-fold. A phosphocreatine shuttle allows the energy produced in the mitochondria to reach the flagella (Tombes and Shapiro, 1985). The regulation of pH, in spermatozoa is under the influence of the ion composition of the surrounding environment. In Na+-free sea water, sea urchin sperm activation is inhibited; it can be restored by adding Na+ or NH4+ (Schackmann et al., 1981; Christen et al., 1982, 1983c; Johnson et al., 1983; Lee et al., 1983; Bibring et al., 1984). The increase in pH, that occurs upon sperm dilution is mainly due to the activation of an unusual, and yet not fully characterized, Na+/H+ exchange, which is amiloride-insensitive and Mg2+- and voltage-dependent. This Na+/H+ exchange has been studied in isolated sperm flagella and in vesicles derived from them (Lee, 1984a,b, 1985). It has also been reported that this Na+/H+ exchange is somehow modulated by Zn2+ (Clapper and Eppel, 1985). As in other cells, the Na+-K+ATPase maintains intracellular Na+ low and participates in regulating pH, (Gatti and Christen, 1985).

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The increase in pH, that occurs when spermatozoa activate is also sensitive to the concentration of external K+ ([K+],); sperm activation is inhibited by 100 mM [K+],. The resting membrane potential of sea urchin spermatozoa (-36 to -56 mV; Schackmann et a l . , 1981; Garcia-Soto et al., 1987) is somewhat sensitive to [K+], (particularly in Lyrechinus pictus); its increase could depolarize sperm and inhibit the voltage-dependent Na+/H+ exchange. These results can be explained by the presence of K + channels in the plasma membrane of these cells (LiCvano et a l . , 1985; Guerrero et al., 1987). Thus, sperm motility is regulated by pH,, which is set mainly by the Na+/H+ exchange. This exchange is governed by membrane potential and depends on the Na+ ionic gradient established by the Na+/K+ ATPase. For as long as pH, is above 7.3, dynein ATPase hydrolyzes ATP, and protons produced in this reaction are released from the cell; if the Na+/H+ exchange stops, the cell acidifies and motility is detained. What activates the Na+/H+ exchange upon sperm dilution? It is known that [K+], is higher in semen than in sea water; possibly, the decrease in [K+1, could hyperpolarize and stimulate the voltage-dependent Na+ IH+ exchange. Spermatozoa from many species are immotile in the seminal tract. It has been known for a long time (Schlenk and Hahaman, 1938) that high [K+], IS responsible for keeping trout sperm inactive. This has been further investigated in trout (Morisawa and Suzuki, 1980; Morisawa et al., 1983) and shown to be the case for sea urchins (Christen et al., 1982, 1983b, c; Schackmann et a l . , 1984). In rainbow trout spermatozoa, a decrease in [K+], initiates motility (Morisawa and Okuna, 1982) and causes an immediate transient increase in CAMP (Morisawa and Ishida, 1987). Activation of motility requires a CAMP-dependent phosphorylation of axonemal proteins (Morisawa and Hayashi, 1985). More recently it was reported that hyperpolarization leads to activation of motility and membrane potential depolarization to inactivation in a pH,-independent fashion (Boitano and Omoto, 1991). A later report by these authors indicates that a sixfold transient increase in [Ca2+], coming from intracellular stores may mediate motility activation (Boitano and Omoto, 1992). Also in marine teleosts, puffer, and flounder, motility activation ensues upon hypertonic dilution in nonelectrolyte solutions. This activation appears to involve an increase in pH, and [Ca2+],; however, the way these ion movements are coupled to motility is not known (Oda and Morisawa, 1993). What is the relationship between hyperpolarization and the [CAMP] increase? A couple of years ago it was reported that an adenylyl cyclase not modulated by G proteins from paramecium is directly stimulated by hyperpolarization (Schultz et al., 1992). The similarity in the characteristics of this enzyme with the sea urchin sperm adenylyl cyclase, which also appears insensitive to G proteins (Hildebrandt et al., 1985; Garbers, 1989), made the authors of this chapter wonder if the latter could also be modulated by membrane potential. Interestingly, Garbers and Hardman (1975) observed a twofold increase in [CAMP] 1 min after diluting L. pictus sea urchin spermatozoa in sea water, which lowers

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[K+], from 27 mh4 in semen to 10 mM. At the time they did not pay much attention to this stimulation. Preliminary results indicate that it is possible to show that hyperpolarization of sea urchin spermatozoa stimulates increases in CAMPthat are independent of [Ca2+Ii,pH,, and phosphodiesterase activities and thus can only be explained by the membrane potential stimulation of adenylyl cyclase (Beltran et al., 1995). The sea urchin sperm adenylyl cyclase is also modulated by [Ca2+],and pH, (reviewed in Garbers, 1989; Cook and Babcock, 1993a, b; Beltran er al., 1995). It seems worthwhile to further consider the interplay among membrane potential, cyclic nucleotide metabolism, pHi, and [Ca2+I, in order to better understand the mechanisms that regulate sperm motility.

B. Mammalian Spermatozoa

In the process of acquiring the capacity to fertilize, sperm [Ca2+],may play an important role. Progressive increases in [Ca2+], have been reported to occur during maturation of some sperm species, leading to hyperactivated motility (White and Aitken, 1989) and spontaneous AR (Langlaic and Roberts, 1985; Bavister, 1986; Yanagimachi, 1988). The influence of pH, on the process of maturation and capacitation is still unsolved (Meizel and Deamer, 1978; Working and Meizel, 1983; White and Aitken, 1989). Since it has been described that pH, may modulate Ca2+ permeability in sea urchin (Garcia-Soto and Darszon, 1985; Guerrero and Darszon, 1989b) and mammalian spermatozoa (reviewed in Florman and Babcock, 1991), it is possible that an acidic pH, contributes to the maintenance of membrane potential (Calzada er al., 1988) and low [Ca2+],thus preventing untimely AR. In their path through the epididymis, spermatozoa encounter important variations in ion concentrations. For instance, K+ increases from -20 mM in the caput to -40 mM in the cauda. On the other hand, the Na+ concentration decreases from more than 100 mM in the caput to less than 50 mM in the cauda (Jenkins et al., 1980). Increases in [K+] may depolarize and open voltagedependent Ca2+ channels known to be present in sea urchin and mouse spermatozoa (Cox and Peterson, 1989; Florman et al., 1992; Beltran et al., 1994). This situation in turn could trigger premature exocytosis. However, it could also be thought that the low [Ca2+]in epididymal fluids (Jenkins et al., 1980) and the decrease in “a+], which may acidify pHi, would counterbalance the tendency to open Ca2+ channels and in this manner prevent spontaneous AR. There must be a subtle balance between conditions that push sperm to a premature AR in transit through the epididymis and the female reproductive tract, which may serve to choose the tightest and fittest, and those conditions that counterbalance the environmental changes, so that the chosen population may survive and be properly stored till fertilization.

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IV. long-Range Communication between Gametes The exchange of information at a distance serves an important role in fertilization. In organisms of external fertilization, where gametes undergo an immense dilution after spawning, it appears crucial to activate sperm and inform them about the whereabouts of the egg. On the other hand, in organisms where gametes interact in the female reproductive tract (internal fertilizers), long-range signaling prepares gametes for fertilization and promotes preferential interactions of the egg with the fittest subpopulations of sperm. Some of these signals stimulate the directed movement of sperm toward the egg (chemotaxis) andlor enhance their motility and metabolism (chemokinesis). Egg factors have been described to cause chemotaxis in spermatozoa from plant and animal species (Miller, 1985). It can be difficult to distinguish between chemotaxis and chemokinesis; here, they will both be considered as long-range gamete signaling processes (Ward and Kopf, 1993). Many small peptides that diffuse from the egg outer layer of a wide variety of echinoderm species, such as sea urchins, starfish, and sand dollars, have been isolated and shown to alter sperm behaviour (Suzuki and Yoshino, 1992). On the other hand, various egg components purified from the hard coral Montipora digitata (Coll et a l . , 1990) and from the horseshoe crab Limulus polyphemus (Clapper and Eppel, 1982, 1985) act in concert to induce chemotaxis and activation in the homologous sperm. In the Pacific hemng, Clupea pallasi, sperm are immobile until they are exposed to a 105-kDa glycoprotein isolated from egg micropyles (Pillai et al., 1993). Chemoattractants isolated from starfish ovaries appear to induce responses involving phospholipid methylation in homologous spermatozoa (Tezon et al., 1986). Little is known about the second messengers and ion permeability changes involved in these responses.

A. Sea Urchins

Although peptide structural differences frequently account for species specificity, egg-conditioned media from a particular sea urchin species may contain several variants of a peptide (Shimomura et al., 1986b; Suzuki er al., 198Xa, b). Speract (also named SAP- l), a decapeptide (Gly-Phe-Asp-Leu-Asn-Gly-Gly-GlyVal-Gly) of this kind isolated from S. purpuratus and Hemicentrotus pulcherrimus (Hansbrough and Garbers, 1981a; Suzuki et al., 1980, 1981), has been cloned. The DNA sequences from an ovary cDNA library of the putative precursor protein contained multiple speract and speract-like structures. Thus, the diverseness in these peptides can be explained by synthesis of variants from a single mRNA (Ramarao et al., 1990). What is the physiological function of these peptides; how do they relay information from the egg to the sperm? They stimulate sperm phospholipid metabo-

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lism, respiration, and motility (Hansbrough et al., 1980; Hansbrough and Garbers, 1981a, b; Suzuki et al., 1982; Ward et al., 1985a; Shimomura and Garbers, 1986; Shimomura et al., 1986a, b; Suzuki et al., 1988a, b; Suzuki and Yoshino, 1992). For instance, pM speract and resact (Cys-Val-Thr-Gly-Ala-Pro-GlyCys-Val-Gly-Gly-GlyArg-Leu-NH,), a similar peptide isolated from Arbacia punctulata (Suzuki et al., 1984), induce these changes in spermatozoa suspended in acidified sea water. However, this barely occurs at the physiological pH, where 100- to 1,000-fold higher peptide concentrations are required (Kopf et al., 1979; Hansbrough et al., 1980; Hansbrough and Garbers, 1981a; Suzuki and Garbers, 1984; Mita et al., 1990). However, since induction of the AR with egg jelly reduces respiration, it was shown that these peptides can overcome this inhibition and enhance fertilization (Suzuki and Garbers, 1984). Along this line, speract promotes AR in H . pulcherrirnus, acting in concert with the main inductor of the reaction in egg jelly (Yamaguchi et a l . , 1988; Shimizu et al., 1990). These results indicate that cooperativity between egg factors may be important in improving the success rate of fertilization. In addition to the changes discussed above, these peptides profoundly alter the plasma membrane permeability of sea urchin spermatozoa. Initially it was shown that at nanomolar concentrations speract and resact stimulate uptake of ,,Na+, 45Ca2+,release of H+, K+ efflux, and increases in [Ca2+],and pH, (Hansbrough and Garbers, 198 1b; Repaske and Garbers, 1983; Lee and Garbers, 1986; Schackmann and Chock, 1986). This decapeptide triggers a K+-dependent hyperpolarization in S. purpuratus sperm flagella and flagellar plasma membrane vesicles, probably mediated by the opening of K + channels (Lee and Garbers, 1986; Garbers, 1989). In these membranes, GTPyS stimulates the speract-induced hyperpolarization, suggesting the participation of G protein (Lee, 1988). Indeed, Gi has been detected in sea urchin spermatozoa (Kopf et al., 1986; Bentley et al. , 1986a). More recently, Gs was identified in sperm flagellar membranes by cholera toxin [3,P]ADP ribosilation and immunoprecipitation with anti-Gs. In addition, three low molecular weight G proteins were spotted using [32P]GTP blotting in flagellar and head membranes, and a fourth one found only in head membranes with a monoclonal antibody against Ras p21 (Cuellar-Mata et al., 1995). The speract-induced hyperpolarization stimulates a Na+ /H+ exchange which, even though having a 1I1 stoichionietry, is voltage-dependent. This stoichiometry was determined using methods with very different time resolutions, influx of 22Na+, and a fluorescent dye (BCECF), to determine changes in pH, (Lee, 1984a,b; Schackmann and Chock, 1986); it is therefore worthwhile to reexamine this point. Concomitantly, these peptides increase the levels of cGMP and CAMP (Kopf et al., 1979; Hansbrough and Garbers, 1981a,b; Garbers et al., 1982; Shimomura et al., 1986b; Yoshino et a / . , 1989). Although all these peptides have chemokinetic effects, it is attractive to think that they may mediate chemotaxis (Ward et al., 1985a; Miller, 1985; Brokaw,

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1987; Cook et a l . , 1994). However, chemotaxis has been clearly demonstrated only in A . punctulatu, where spermatozoa are attracted by nanomolar concentrations of resact changing their swimming pattern from a circular to a straighter trajectory. This chemotactic response requires Ca2+ in sea water (Ward et a l . , 198Sa). The species specificity and water solubility of these egg peptides suggest that they interact with sperm plasma membrane receptors. In crosslinking experiments, labeled functional analogs of speract allowed identification of a 77-kDa transmembrane peptide (Dangott and Garbers, 1984). This plasma membrane speract receptor was purified, sequenced, and cloned in S . purpuratus; it has a short predicted cytosolic domain (Dangott e f al., 1989) and is also present in A . punctuluta spermatozoa (Dangott, 1991). The receptor is thought to interact with and modulate the membrane guanylyl cyclase (Garbers, 1989; Schultz et al., 1989; Yuen and Garbers, 1992). In A . punctulatu, nanomolar resact induces the dephosphorylation of a membrane protein, changing its apparent molecular weight from 160 to 1.50 kDa (Ward and Vacquier, 1983). This protein has been isolated and identified as guanylyl cyclase (Ward et al., 1985b). It was shown that the phosphorylated enzyme had a higher activity and that dephosphorylation decreased it (Ramaro and Garbers, 1985; Ward et al., I985b). The phosphorylated state of guanylyl cyclase is pH-dependent in vitro and in vivo (also "a+],-dependent; Ward, 198.5), dephosphorylation being enhanced at alkaline pH (Ward, 1985; Ward et a!., 1985b; Suzuki et al., 1984; Vacquier and Moy, 1986; Ward et al., 1986; Bentley et al., 1986b). The phosphatases and kinases which regulate the phosphorylation state of guanylyl cyclase have not been identified, and they could also be pHi-dependent. Using a similar crosslinking strategy, it was later found that nanomolar resact binds to a 160-kDa plasma membrane protein identified as a guanylyl cyclase (Shimomura et al., 1986a). This cyclase was cloned; its predicted amino acid sequence is homologous, in the amino-terminal region, to the atrial natriuretic factor receptor (Singh et al., 1988). The resact receptor turned out to be the first cloned and sequenced member of a family of guanylyl cyclases that are surface receptors participating in a new signal transduction pathway (Garbers, 1989, 1992; Drewett and Garbers, 1994). So far it has not been possible to functionally express the sea urchin sperm guanylyl cyclase in heterologous systems like COS-7 cells (Drewett and Garbers, 1994). It is surprising that the speract and resact receptors are significantly different, even though they trigger similar changes in sea urchin spermatozoa. Although speract analogs are not crosslinked to the S. purpuratus membrane guanylyl cyclase from sperm, it is 77% identical to that of A . punctuluta (Thorpe and Garbers, 1989). However, in L. pictus sperm, speract induces an apparent molecular weight change from 160 to 150 kDa, in a similar fashion to that in A . punctulata. Considering the limitations of crosslinking experiments, the gua-

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nylyl cyclase peptide receptor could be formed from several subunits (Garbers, 1989, 1992; Drewett and Garbers, 1994). Recently it was reported that a radiolabeled analog of another sperm-activating peptide, SAPIII, binds to two classes of receptors with different affinities having K,s of 3.4 and 48 nM. Crosslinking experiments with the analog revealed labeling of three proteins of 126, 87, and 64 kDa of apparent molecular weight (Yoshino and Suzuki, 1992). As explained above, egg peptides like speract and resact induce important ion permeability changes in sea urchin spermatozoa. How these changes are related to those that occur in the cGMP and CAMPlevels holds the key to understanding the information processing system used by sea urchin gametes to effectively meet and fertilize. The authors believe that the mechanisms that are being uncovered will probably contribute to our understanding of gamete signaling in general and to some of the most interesting problems of signal transduction in biology. Though incomplete, a picture emphasizing the fundamental role that various ion channels play in the speract-induced sperm responses is emerging. Spermatozoa are small cells. This has precluded their electrophysiological characterization and motivated the use of complementary strategies to understand how their ion transport systems participate in gamete communication (Darszon et al., 1988; Schackmann, 1989; Cox and Peterson, 1989; Florman and Babcock, 1991; Darszon e t a l . , 1994). In vivo measurements of [Ca2+Ii,pH, (Schackmann and Chock, 1986; Guerrero and Darszon, 1989a, b), membrane potential (EM [Schackmann et al., 198 1; Gonzdez-Martinez and Darszon, 1987; Garcia-Soto et nl., 1987; Babcock el al., 1992]), and patch clamp techniques (Guerrero er al., 1987; Babcock et al., 1992), together with reconstitution in planar and spherical bilayers (reviewed in Darszon et al., 1994) have revealed the presence and participation of Ca2+, K + , and C1- channels in sea urchin spermatozoa responses to the egg coats. These combined strategies are allowing researchers to explore their regulation. An alternative to circumvent the size limitation of sea urchin sperm is to swell them in 10-fold-diluted sea water plus 20 mM MgSO,. The swollen cells are spherical (-4 pM diameter) and regulate their EM,pHi, and [Ca2+],. Their main virtue is that they can be patch-clamped (Babcock et al., 1992), a difficult endeavor with normal sperm (Guerrero et al., 1987). Picomolar speract provokes a long-lasting, K+-selective, and TEA+-sensitive permeability increase in swollen sperm, mediated by K + channels, as indicated by patch clamp experiments (Babcock et al., 1992; see Fig. 1). Higher concentrations (>25 pM) transiently hyperpolarize the cells close to the K+ equilibrium potential (EK)and immediate repolarize them toward the resting potential (ER).The hyperpolarization appears to activate Na+/H+ exchange. A direct link between membrane potential and pH, in the response to speract in whole sperm has been suggested in experiments where a valinomycin-induced hyperpolarization increases pH, (Lee, 1984b; Gonzalez-Martinez et al., 1992; Reynaud et al., 1993). In addition to the changes described above, nanomolar speract increases

b

Na' ?

L**117w**)Control + CAMP

400 pM

sw msec Fig. 1 Schematic diagram of the sea urchin sperm responses to speract. Guanylyl cyclase in sea urchin sperm is indirectly activated by binding of speract to its receptor (1) in S . purpuratus or by the direct binding of resact in A. punctulata (2). The transient increase in [cGMP], after an unknown number of steps (X,,), opens a K + channel (3) probably responsible of the initial transient hyperpolarization. This later change initiates other important alterations in membrane potential (A&,), and a Na+/CaZ+ exchange (4) is also activated. At appropriate [speract], the hyperpolarization activates a Na+/H+ exchange (5) which increases intracellular pH (ApH,). The changes in pH, directly or indirectly modulate the activities of adenylyl cyclase (AC, 6 ) , guanylyl cyclase, and probably also some kinases, phosphatases, and phosphodiesterases. AE, also participates in AC (6) regulation. The increase in [CAMP] activates a CAMP-dependent K + channel involved in the Speruct-induced depobdrization (7). Concomitant changes in pH, and [CAMP] may modulate a Ca2+ channel (8). In addition, the increase in [cGMP] would regulate a CaZ+ influx through a cCMP-regulated channel (9). like the one found in photoreceptors and recently in mammalian sperm. At least two different receptors to speract-like peptides are present in sea urchin sperm. One of them (10) could activate a G protein that might modulate K+ channels (3). The traces on the right show simultaneous measurements of EM (A), pH, (B), and of [Ca2+],(C), upon addition of 100 nM speract to S. purpuratus sperm in ASW. Upward deflections indicate depolarization (A), alkalinization (B), and [Ca2+], increase (C). EM (A) was measured with the fluorescent probe Dis-C,-(5), and the changes are expressed in mV, after titration of the record with valinomycin and K + additions. BCECF was used to measure pH, and Fura-2 for [Ca2+],. The single-channel records shown correspond to the K + channels activated by speract addition in swollen sperm (3) and the CAMP-regulated K + channels (7) in planar lipid bilayers.

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[Ca2+], and a CaZ+-dependent depolarization occurs beyond E, (Babcock et al., 1992; Reynaud et al., 1993; Cook and Babcock, 1993a). The permeability to Na+, Ca2+, and Mg2+ contributes to the resting membrane potential of swollen sperm (Reynaud et al., 1993). It is likely that the Ca2+-dependent depolarization triggered by nanomolar speract and at least part of the [Ca2+Iiincrease (de De Latorre and Darszon, unpublished results) involve channels, since they are inhibited by Ca2+ channel blockers like Co2+, Ni2+ and Zn2+ (Darszon et al., 1990; Reynaud et al., 1993; Cook and Babcock, 1993b). As shown before for a Ca2+ channel opened during the AR (Guerrero and Darszon, 1989b), these Ca2+permeable channels allow MnZ+through and are regulated by cAMP (Cook and Babcock, 1993b). Exploiting papaverin and isobuthyl-methylxantine (IBMX), two inhibitors found to preferentially act upon the cAMP or cGMP sea urchin sperm phosphodiesterases, respectively, the relationship between cyclic nucleotide metabolism, membrane potential, pH,, and [Ca2+], was further examined using swollen sperm (Cook and Babcock, 1993a, b). With the available information, a working model for the action mechanism of speract can be delineated (see Fig. 1). This decapeptide activates guanylyl cyclase (reviewed in Trimmer and Vacquier, 1986; Garbers, 1989). The increase in [cGMP] opens TEA+-insensitive, K+-selective channels that hyperpolarize sperm (Lee and Garbers, 1986; Babcock et a l . , 1992; Cook and Babcock, 1993a). Since there is not much evidence at present for direct cGMP modulation of K+-selective channels, K+ channels activation by cGMP could be indirect in sperm, perhaps through phosphorylation. Furthermore, or alternatively, one of the speract receptors could be coupled to a G protein which might directly or indirectly activate K+ channels (Lee, 1988). It is not clear if a threshold value of hyperpolarization or its rate of change, both of which are dependent on the speract concentration, activate Na+/H+ exchange (Lee and Garbers; 1986, Babcock et a l . , 1992; Reynaud et al., 1993). At an appropriate speract concentration (>100 pM), the resulting increase in pH, inhibits guanylyl cyclase (Suzuki et al., 1984; Ward et al., 1985a, b; Vacquier and Moy, 1986; Ward el al., 1986; Bentley et a / ., 1986b) and stimulates adenylyl cyclase, which is pHi- (Cook and Babcock, 1993a,b), membrane potential- (Beltrh et al., 1995), and [Ca2+]-sensitive (reviewed in Garbers, 1989). The decrease in [cGMP] would diminish K+ permeability (Cook and Babcock, 1993a) and repolarize sperm. Two (or more) ion channels with distinct selectivity and pharmacology might contribute to the depolarization triggered by nanomolar speract in normal sea urchin spermatozoa: a CAMP- and/or pH,-regulated Ca2+ channel (Darszon et al., 1990; Babcock er al., 1992; Cook and Babcock, 1993b) and a CAMP-regulated K+ channel that allows Na+ flux into sperm (Labarca et al., 1995). In fact, a K+-selective channel, directly modulated by CAMP, has been detected in planar lipid bilayers with incorporated flagellar sperm membranes. This channel is blocked by TEA+ (30 mM) and Ba2+ and has a P,+/P,,+ of 5; therefore, in sea water, its opening would depolarize sperm (Labarca et a/.,

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1996). The CAMP-regulated channel could also contribute to the repolarization and explain part of its Na+-dependence (Reynaud et al., 1993; Labarca et al., 1996). As in photoreceptors, cGMP could also up-regulate a cation-selective channel permeable to Ca2+ (Yau, 1994). Recently, a member of this ion channel family, found in mammalian spermatozoa, has been cloned and functionally recorded (Weyand et al., 1994). In sea water, such a channel would not hyperpolarize sea urchin sperm but depolarize it. A Na+/Ca2+exchanger may participate in the increase in [Ca2+Ii triggered by speract and the regulation of [Ca2+Ii (Schakmann and Chock, 1986). Recently it was shown that the simultaneous addition of speract (50 nM) and a phosphodiesterase inhibitor (100 ph2 IBMX), but not speract alone, produces in S. purpuratus spermatozoa a hooked flagellar waveform, characteristic of high flagellar asymmetry (Cook et al., 1994). Nonetheless, the fact that in the presence of IBMX, nanomolar speract induces a significant percentage of AR (Schackmann and Chock, 1986) is not discussed. This may influence the swimming behavior of sperm in a nonphysiological manner. Based on previous information and the analogy between the resact responses, where chemotaxis has been demonstrated (Ward et al., 1985), and those of speract, an interesting model is presented to explain how sperm can detect an increasing egg peptide gradient over a broad concentration range (Cook et al., 1994). It is worth pointing out that Cook et a1 (1994) used the speract-induced changes in [Ca2+], and membrane potential of swollen sperm to build their model for chemotaxis in normal spermatozoa (see their Fig. 6). However, the speract responses of swollen and normal spermatozoa differ significantly (Labarca et al., 1995a). For instance, the ion selectivity and pharmacology of the depolarizing phase, seen with nanomolar speract, is different in normal and swollen sperm. This is most likely due to the dissimilar ion composition of the external media and the loss of compartmentali zation in swollen sperm. In the latter, the depolarization is Ca2+-dependentand sensitive to divalents which block Ca2+-selective channels (Darszon et al., 1990; Babcock et a/., 1992; Reynaud et a l . , 1993; Cook and Babcock, 1993b). In normal sperm, the depolarizing phase is only partly diminished in the absence of external Ca2+, depends on external Na+, is poorly sensitive to Ca2+ channel blockers, and is blocked by TEA+ and Ba*+, (Labarca et al., 1995a).

B. Mammals Mammalian spermatozoa are delivered in the female reproductive tract, having an arranged pathway toward the egg. In spite of this, long-range communication with the egg may also be important. A significant fraction of ejaculated spermatozoa from rabbits, pigs, hamsters, sheep, and cattle appears to have reduced motility when stored in the caudal isthmus of the oviduct (Harper, 1973; Flechon and Hunter, 1981; Hunter and Nichol, 1983, 1986; Hunter and Wilmut, 1984).

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Minutes after ovulation, sperm become motile, leave their storage sites, and reach the ampullary region (Harper, 1973; Overstreet and Cooper, 1979; Flechon and Hunter, 1981). In the case of women, ejaculated spermatozoa are stored in the cervix (Zinaman eta!., 1989). These results have led to the belief that eggs or follicle cells release factors that activate motility and guide sperm toward the ovulated egg. These factors may improve productive encounters of the fittest gametes, particularly considering that the spermlegg ratio is low ( I / 1 to 10/ 1) at the site of fertilization (reviewed in Yanagimachi, 1988; Ward and Kopf, 1993). A recent report has indicated that human follicular fluid from women undergoing in vitro fertilization contains compounds involved in chemotaxis and/or chemokinesis (Villaneuva-Diaz et al., 1990). Another study has shown that diluted human follicular fluid can change the swimming pattern of human spermatozoa and may contain a chemoattractant (Ralt et al., 1991). Even though chemotaxis is difficult to determine in mammalian spermatozoa, it would be very interesting to determine the nature of these factors, their receptors, and how they alter sperm ion permeability.

V. Short-Range Communication between Gametes: The Acrosome Reaction The acrosome reaction is an absolute requirement for successful fertilization in all sperm species possessing an acrosome. This organelle, found in the sperm head, is synthesized and assembled as a product of the Golgi complex during spermiogenesis. Its basic function is similar among many species. Triggering of this fundamental process involves short-range interactions between sperm and signals coming from the egg’s outer layers and, in the case of internal fertilizers, also from the female reproductive tract. This section will focus on those species where more information about the participation of ion channels during the AR is available.

A. Sea Urchin Sperm Acrosome Reaction

Contact of spermatozoa with the egg jelly layer triggers the AR (Dan, 1952; Tilney, 1985). As mentioned in section IIB, the egg jelly component responsible for inducing the AR is a FSP (SeGall and Lennarz, 1979; Garbers et al., 1983). The latest results indicate that a glycoprotein(s) in FSP is the AR-inducing factor (Suzuki, 1990; Mirakami-Takei et al., 1991; Keller and Vacquier, 1994). This reaction involves acrosomal vesicle exocytosis (Dan, 1952; Summers and Hylander, 1975, which exposes material required for sperm-egg binding (Vacquier and Moy, 1977; Glabe and Lennarz, 1979) and lytic enzymes (Levine and Walsh,

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1979; Green and Summers, 1980). These events lead to the extension of the acrosomal tubule, which is surrounded by the membrane destined to fuse with the egg (Trimmer and Vacquier, 1986). The AR requires external Ca2+ and Na+ in seawater at pH 8.0 (Dan, 1954; Collins and Epel, 1977; Shackmann and Shapiro, 1981). Exposure of spermatozoa to FSP induces, within seconds, Na+ and Ca2+ entry and H+ and K + efflux (Schackmann et al., 1978; Garbers and Kopf, 1980; Schackmann and Shapiro, 1981; Garbers, 1989; Schackmann, 1989). These ion fluxes lead to interrelated changes in EM (Schackmann et al., 1984; Gonzalez-Martinez and Darszon, 1987; Garcia-Soto et al., 1987), [Ca2+],(Trimmer et al., 1986; Guerrero and Darszon, 1989a,b), and pH, (Lee et al., 1983; Guerrero and Darszon, 1989b). FSP also raises the concentration of cAMP (Garbers and Kopf, 1980), protein kinase A activity (Garbers et al., 1980; Garcia-Soto et al., 1991), tumover of inositol 1,4,5-trisphosphate (InsP, [Domino and Garbers, 1988]), and stimulation of a phospholipase D (Domino and Garbers, 1989), changes which bear an unclear relationship to the FSP-induced permeability changes. Apparently the increase in [CAMP] precedes the AR (Garbers, 1981) and depends on Ca2+ uptake (Garbers and Kopf, 1980). The FSP-induced accumulation of cAMP results from stimulation of a sperm adenylyl cyclase (Watkins et al., 1978). A 2 10-kDa plasma membrane protein is, at the present time, the best candidate for the FSP receptor in S. purpuratus spermatozoa. This protein has specific affinity for FSP (Podell and Vacquier, 1984a), and some monoclonal antibodies to it block the AR (Trimmer et al., 1985) and inhibit the increase in pHi, yet cause large increases in [Ca2+],(Trimmer et al., 1986) and also activate sperm adenylyl cyclase (Vacquier et al., 1988). Immunofluorescence localization of these monoclonal antibodies shows that they bind to a narrow collar of plasma membrane over the acrosome and also with the entire flagellum (Trimmer et al., 1985). Wheat germ agglutinin also binds to the 210-kDa protein and blocks the AR (Podell and Vacquier, 1984b). These results strongly suggest that the 210kDa protein is involved in regulation Ca2+ channels activated by FSP during the AR (Trimmer et al., 1986). The first membrane potential response observed in L . pictus spermatozoa exposed to FSP is a transient hyperpolarization, after which they depolarize. This hyperpolarization is K+-dependent and probably mediated by K+ channels (GonzAlez-Martinez and Darszon, 1987). It is believed that the FSP-induced hyperpolarization leads to an increase in pH,, at least in part activating Na+/H+ exchange (Gonzalez-Martinez et al., 1992). In agreement with this idea, a small increase of [K+] in seawater (from 10 to 30-40 mM) blocks Ca2+ uptake, the AR (Schackmann, 1978), pHi increase (Guerrero and Darszon, 1989b), and hyperpolarization (Gonzalez-Martinez and Darszon, 1987). Antagonists of Ca2+ channels (verapamil and dihydrophyridines) (Schackmann et al., 1978; Garbers and Kopf, 1980; Kazazoglou et al., 1985; GarciaSoto and Darszon, 1985) and K+ channels (TEA+) (Shackmann et al., 1978)

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inhibit Ca2+ uptake and the AR, indicating their mandatory participation in this process (reviewed in Garbers, 1989; Schackmann, 1989; Darszon et al., 1994). K+ single channels were first recorded in bilayers made at the tip of patch pipettes from monolayers generated from a mixture of lipid vesicles and isolated sperm flagellar membranes. Three types of K+ channels, with conductances of 22, 46, and 88 pS, were identified. Two of them were blocked by TEA+, which inhibits the AR (LiCvano et al., 1985). Thereafter, with great difficulty, single channels were recorded directly from sea urchin sperm heads using the patch clamp technique. Single channel events of 40,60, and 180 pS were detected, and one of the channels observed was a K+ channel (Guerrero et al., 1987). As mentioned in section IVA swelling S. Purpuratus sperm improves significantly the success rate of patch formation and allows the detection of a 2 pS K+ that is activated by picomolar speract. Swollen sperm have opened new possibilities for directly studying the ion channels modulated by egg components and their regulation (Babcock et al., 1992). Through the use of fura-2-loaded sea urchin sperm it was found that two different Ca2+ channels participate in the AR (Guerrero and Darszon, 1989a, b; Schackmann, 1989). The first type of Ca2+ channel is activated when egg jelly binds to its receptor and inactivates. The key question of how the receptor activates the channel remains unanswered; the possibilities are: (a) through a G protein directly or indirectly using second messengers; (b) by opening a built-in channel like the ACh receptor; or (c) by some new unknown mechanism. Verapamil and dihydropyridines block the first type of channel. The second channel is not blocked by these compounds, does not inactivate, and allows Mn2+ to permeate. Conditions that inhibit the egg jelly-induced pH, increase, block the second type of channel and the AR, but still support a transient increase in intracellular Ca*+ due to the opening of the first channel. Opening of the second channel requires activation of the first type of channel. Development of a normal AR apparently require both channels (A. Darszon and M. T. Gonzalez-Martinez, unpublished), but how they are coupled is still a mystery. Could a type of Ca2+induced Ca2+ release be the answer (Endo, 1977; Fabiato, 1985) or could it be a modification of the protein, like a change in its phosphorylation status or even proteol ysis? Fusion of isolated S. purpuratus sperm plasma membranes into planar lipid bilayers has revealed the presence of two types of Ca*+ channels: (1) a 50 pS voltage-dependent channel observed in 10 mM Ca2+; and (2) a high-conductance, voltage-dependent channel having a main conducting state (172 pS in 10 mM CaCI,) and several subconductance states (for reviews on planar bilayer reconstitution, see Darszon, 1986; Darszon et al., 1994). The high-conductance channel discriminates poorly between divalent and monovalent cations (Pca2+lPNa+ = 5.9 [Likano et al., 1990)). It remains to be seen if the small Ca2+ channel detected in bilayers is the first type of Ca2+ channel which opens during the AR, and whether it is a ligand-gated channel. The high-conductance

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Ca2+ channel detected in bilayers is insensitive to verapamil and nisoldipine and is blocked by Cd2+ and Co2+ at concentrations similar to those required to inhibit AR and Ca2+ uptake induced by egg jelly. Because of this, it might be the second type of channel that participates in the AR, allows Mn2+ influx, and is pH,-modulated (LiCvano et al., 1990). Recent collaborative work from the authors’ laboratory and Pedro Labarca has shown that functional ion channels can be transferred to planar lipid bilayers directly using spermatozoa from S . purpuratus and L . pictus and from mouse. Spermatozoa from these species possess conspicuous Ca2+-selective, high-conductance, multistate, voltage-dependent channels similar to the one described previously from isolated S. purpuratus sperm membranes (LiCvano et al., 1990). In the three species, the channel displays similar voltage dependence and equal PBal+/PK+ (-4). The presence of this Ca2+ channel in such diverse species suggests it is a relevant ion transport mechanism in spermatozoa. The high sensibility of planar bilayers to detect single ion channels can now be exploited further to study ion channel regulation and gamete interaction (Beltran et a / . , 1994). As anion channel of 150 pS was also identified in planar lipid bilayers fusing either sperm plasma membranes or vesicles formed from an enriched preparation in anion channel activity. The anion channel selectivity sequence found was: NO,> SCN- > Br- > C1-. This anion channel has a high open probability at the holding potentials tested, is partially blocked by the stilbene disulfonate DIDS, and often displays substates (see Fig. 2). DIDS blocks the AR in S. purpuratus sea urchin sperm by a still unknown mechanism. These results suggest that this C1- channel could be involved in the events that lead to the AR or in determining the resting potential of sperm, which modulates this reaction (Morales et d., 1993). Figure 2 illustrates a working model for the sea urchin AR. It is remarkable that still so many fundamental questions remain unanswered. How does binding of egg jelly to its receptor start the transduction cascade? What are the molecular properties of this receptor? What activates the K + channel responsible for the hyperpolarization that stimulates Na+/H+ exchange; could it be a Ca2+-regulated channel? So far, no Ca2+-dependent K+ channels have been described in spermatozoa, and even though this type of channel is present in most cells (Latorre et al., 1989; Brown, 1993), the regulation could be indirect. What entity(ies) is responsible for the peculiar amiloride-insensitive, electroneutral but voltage-dependent Na+/H+ exchange found in sea urchin spermatozoa? Why is the increase in pH, critical for the AR, and what are the targets? Possibly they are the high-conductance Ca2+ channel, a protease (Farach et al., 1987; Matsumara and Aketa, 1990), adenylyl cyclase, some kinase, or a phosphatase. How are the increases in CAMP and InsP, related to the permeability changes that occur during the AR and how are they so exquisitely orchestrated? A CAMP-regulated channel, which could very well be one of the pieces of the puzzle, has been

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